methods

TECHNIQUES IN CELL AND MOLECULAR BIOLOGY

dna tech test

OBJECTIVES

 Describe the way that various techniques are used.

 Provide examples of the types of information that can be obtained by using these techniques.

 Define magnification and resolution and explain their relative importance to light microscopy.

 Describe the components of the light and electron microscopes and their functions.

 Emphasize the importance of contrast to specimen visibility in the light and electron microscope.

 Outline different light and electron microscopy techniques emphasizing the information they reveal and their advantages and           disadvantages.

 Emphasize the reason for the much higher resolution obtained with electron microscopy.

 Outline the factors that cause the formation of different radioisotopes and describe the major methods used to detect    radioactivity.

 Describe cell culture techniques and the advantages of working with a cell culture system.

 Explain purification of cellular organelles by differential centrifugation.

 Discuss various methods used to purify proteins and nucleic acids and the traits of these molecules by which the    methods effect purification.

 Present the principles of ultracentrifugation and explain velocity sedimentation and equilibrium density centrifugation.

 Outline nucleic acid hybridization techniques and recombinant DNA technology.

 Discuss amplification of DNA sequences by polymerase chain reaction.

 Describe the techniques by which DNA fragments can be sequenced.

 Discuss the use of antibodies in research and the production and advantages of monoclonal antibodies.

LECTURE OUTLINE

The Light Microscope: Principles

I.  Components of the light microscope – a microscope produces an enlarged image of an object

A.  Light source - external to microscope or built into base; illuminates specimen

B.  Substage condenser lens - gathers diffuse rays from light source; illuminates specimen with a small cone of bright light   intense enough to allow very small parts of a specimen to be seen after magnification

C.  Objective lens – collects light rays focused on specimen by condenser lens; collects 2 kinds of light rays passing through slide/specimen

1.  Rays altered by specimen – emanates from the many parts of the specimen; focused by objective lens in scope column forming real, enlarged image of the object

2.  Rays not altered by specimen - pass directly into objective; form visual field background light

D.  Ocular lens - image formed by objective lens is used as an object by ocular lens (a second lens system); picks up real image formed in scope column & forms enlarged, virtual image

E.  Eye lens system - uses virtual image formed by ocular as an object to form a real image on the retina

F.  Focusing knob – when turned, knob alters relative distance between specimen & objective, focusing the final image precisely on plane of retina

II.  Magnification = product of magnification produced individually by ocular & objective lenses

III.  Resolution – the ability to see 2 neighboring points in a field as distinct entities

A.  If an image is magnified beyond ability to resolve (see additional detail), result is empty magnification

B.  Degree of resolution defines objective lens quality; the extent to which fine detail in specimen can be discriminated or resolved

C.  Resolution attained by a microscope is limited by diffraction

1.  Because of diffraction, light emanating from a point in specimen can never be seen as a point in the image, but only as a small disk

2.  If the disks produced by 2 nearby points overlap, the points cannot be distinguished in the image

3.  Thus, the resolving power of a microscope can be defined in terms of the ability to see 2 neighboring points in the visual field as 2 distinct entities

4.  If 2 parts of a specimen are not separated by sufficient distance —> images merge (not resolved)

D.  Resolution is determined by the following equation & is limited by the wavelength of illumination & numerical aperture

1.  d = minimum distance that 2 points in specimen must be separated by in order to be resolved

2.   = the wavelength of light (527 nm used for white light)

3.  n = refractive index (RI) of medium present between the specimen & the objective lens

4.   = half the angle of cone of light entering objective lens (angle between normal to base of light cone & its side);  is a measure of light-gathering ability of lens; directly related to its aperture

5.  N. A., the denominator of the equation, is a measure of the light-gathering qualities of the lens (a constant for each lens)

E.  For an objective designed for use in air, largest N. A. possible is 1.0; for an oil immersion objective, the maximum possible N. A. is ~1.5

1.  In air, largest possible angle of  = 90°, so the maximum sin  = 1 & n (RI of air) = 1; thus,       N. A. = n sin  = 1 x 1 = 1

2.  Rule of thumb – maximum useful magnification of a microscope = objective N. A. x 500 - 1,000; magnification beyond this amount, you get empty magnification & quality of image deteriorates

3.  Achieve high N. A. by using short focal length lens; requires lens placement very close to specimen

F.  Limit of resolution obtained by using lowest possible wavelength & highest possible N. A. in equation

1.  Standard light microscope - slightly < 0.2 µm (200 nm); resolves larger cell organelles (nuclei, mitochondria) 

2.  Naked eye - N. A. = ~0.004; so the limit of resolution = ~0.1 mm (0.0036 inches)

G.  Optical flaws or aberrations (7 important ones) also affect resolving power - lens makers must overcome these handicaps so that actual resolving power approaches that of the theoretical limits

1.  Objective lenses are made of a complex series of lenses, rather than a single convergent lens - eliminates the aberrations

2.  One lens unit typically magnifies; the rest compensate for errors in first lens (corrects overall image)

IV.  Visibility – factors that allow an object actually to be observed; largely determined by contrast 

A.  Example – clear glass bead placed in immersion oil with same refractive index as glass —> bead disappears from view since it no longer affects light in any obvious way that is different from background fluid

B.  Contrast is the difference in appearance between adjacent parts of object or object & its background; example - clearly visible stars in night sky; during day, they cannot be seen against bright background

1.  In microscope, you view light transmitted (or diffracted) through the object; to do this, object must be translucent (nearly transparent) & such objects are difficult to see

2.  To make thin, translucent specimens visible under scope —> you must stain them with dye so the specimen appears colored (dye should absorb only certain wavelengths within visible spectrum)

3.  Those wavelengths not absorbed by dye are transmitted to eye, causing object to appear colored

4.  Different dyes bind to different types of biological molecules; they not only heighten specimen visibility, but also effectively localize that molecule in cells & tissues

C.  Many staining procedures, like Feulgen staining (DNA), which causes chromosomes to appear colored in scope, generally cannot be used with living cells

1.  Usually stain or staining conditions are toxic; also, some stains cannot penetrate plasma membrane

2.  In Feulgen staining, tissue must be hydrolyzed in acid before stain is applied

D.  Vital dyes stain organisms without killing them, but their use is limited

E.  To enhance visibility, different types of light scopes use different types of illumination (see next section)

The Light Microscope: Techniques - Bright-Field Light Microscopy and Specimen Production

I.  Bright-field microscopy - condenser converges light on specimen forming bright cone of light that enters objective; cone seen as bright background against which image of specimen is contrasted

A.  Ideally suited for high contrast specimens (stained tissue sections); not good for all specimens

B.  May not provide optimal visibility for other specimens

II.  Two categories of specimens are observed in a light microscope – whole mounts & sections

A.  Whole mounts - intact living or dead object (whole organism [a protozoan] or a small part of a larger organism); view with transmitted light if transparent enough; clear opaque specimen with organic solvents 

1.  Make opaque objects translucent by substituting water in the sample with alcohol & immersing the object in solvents, like toluene & xylene, in which they become clear

B.  Sections – examine as very thin slice of organism; use if organism is opaque & cannot be cleared

1.  Most plant & animal tissues are much too opaque for microscopic analysis unless examined as a very thin slice or section

III.  Section production – first kill cells by immersing in chemical solution called a fixative (formaldehyde, alcohol, acetic acid are the most common for the light microscope)

A.  A good fixative rapidly penetrates cell membrane & immobilizes all of its macromolecular material, maintaining cell structure as close as possible to that of the living state

B.  Tissue is then dehydrated by transfer through a series of alcohols & embedded in paraffin (wax), which provides mechanical support during sectioning

1.  Paraffin is used for embedding because it is readily dissolved by organic solvents

C.  Slides containing adherent paraffin sections are immersed in toluene, which dissolves the wax

D.  The thin slice of tissue is thus left attached to slide where it can be stained or treated with enzymes, antibodies or other agents

E.  After staining, a coverslip is mounted over tissue using a mounting medium with the same RI as glass slide & coverslip

The Light Microscope: Techniques – Phase-Contrast Microscopy

I.  Phase-contrast (P-C) microscopy - good for small, unstained specimens like living cells & those that are hard to see in bright-field; type of interference microscopy; makes highly transparent objects more visible

A.  Can distinguish different parts of object since they affect light differently from one another; organelles differ in refractive index (RI) since they differ in molecular composition 

1.  Organelles are made up of different proportions of various molecules: DNA, RNA, protein, lipid, carbohydrate, salts, water; such regions of different composition will likely differ in refractive index

2.  P-C turns differences in RI into differences in intensity (relative brightness & darkness) that are visible to the eye, while RI differences are not visible to the eye

B.  Their ability to do this centers on ability of light waves to interact with one another (interference); P-C scopes perform 2 functions bright field scopes do not:

1.  P-C scope separates direct light waves (background) entering objective from diffracted light waves emanating from specimen

2.  Causes light rays from the 2 sources to interfere with one another; relative brightness or darkness of each part of image reflects way in which light from that part of specimen interferes with direct light 

3.  Puts 2 light ray types ~0.5 wavelength out of phase; they interact destructively (interfere with one other) —> light intensity (relative brightness or darkness) is altered point to point

C.  Useful in examination of intracellular components of living cells at relatively high resolution

1.  Dynamic motility of mitochondria, mitotic chromosomes & vacuoles followed & filmed with P-C

2.  Watching these organelles moving around in living cell, in seemingly random fashion, conveys excitement about life better than observing stained, dead cells

3.  Greatest benefit of P-C microscopy has not been discovery of new structures, but in its everyday use in research & teaching labs for observing cells in a more revealing way

D.  Disadvantages & limitations of P-C  - only suitable for observing single cells or thin cell layers 

1.  P-C has optical handicaps that result in loss of resolution, and

2.  The image suffers from interfering halos & shading produced where sharp changes in RI occur

E.  Other interference scopes minimize these optical artifacts by completely separating direct & diffracted beams using complex light paths & prisms

II.  Differential interference contrast (DIC) microscopes or Nomarski interference (after its developer) – another type of interference system

A.  Delivers an image that has an apparent 3-D quality (like bas relief - Lincoln's head on a penny)

B.  Contrast in DIC microscopy depends on rate of change of RI across specimen; edges of structures (where RI varies markedly over relatively small distance) are seen with especially good contrast

The Light Microscope: Techniques - Fluorescence Microscopy

I.  Over past couple of decades, the light microscope has gone from being an instrument designed primarily to examine sections of fixed tissues to being able to observe dynamic events at molecular level in living cells

A.  These advances in live-cell imaging have been made possible to a large extent by innovations in fluorescence microscopy

B.  Fluorescence microscope allows viewers to observe the location of certain compounds called fluorochromes or fluorophores

II.  Fluorochromes absorb invisible, UV light & release a portion of the energy as longer, visible light wavelengths called fluorescence

A.  Presence of fluorochromes in cell is observed in fluorescence microscope fitted with UV light source; fluorescent stain then glows against a dark background

B.  Fluorescence microscope light source produces a beam of UV light that is passed through a filter that blocks all wavelengths except the one that excites the fluorochrome

C.  The beam of monochromatic light is focused on the specimen containing the fluorochrome, which becomes excited & emits light of visible wavelength that is focused by objective lens into an image

D.  Because the light source produces only UV (black) light, fluorochrome-stained objects appear brightly colored against a black background, providing very high contrast

III.  Applications of fluorescence – use of fluorescence to localize molecules in the cell; there are many different ways that fluorescent compounds can be used in cell & molecular biology

A.  In one of its most common applications, one covalently links or conjugates a fluorochrome (fluorescein or rhodamine) to an AB to make a fluorescent AB

1.  Fluorescent AB is used to determine location of a specific protein in the cell (immunofluorescence)

B.  Fluorochromes can also used in studies to locate DNA or RNA molecules that contain specific nucleotide sequences by using fluorescently labeled probes

C.  Fluorochromes can be used to study the sizes of molecules that can pass between cells, as indicators of transmembrane potentials or as probes to determine the free Ca2+ concentration in the cytosol

IV.  Applications of fluorescence – use of fluorescently labeled proteins to study dynamic processes as they occur in a living cell (use of green fluorescent protein (GFP) from the jellyfish Aequorea victoria)

A.  A specific fluorochrome can be linked to a cellular protein (e.g., actin, cytoplasmic dynein or tubulin) & the fluorescently labeled protein can then be injected into & followed through the living cell

B.  A noninvasive approach has been widely employed that utilizes a fluorescent protein (green fluorescent protein or GFP) from the jellyfish Aequorea victoria

1.  In most of these studies, researchers construct a recombinant DNA in which the GFP-coding region is joined to the coding region of the protein being studied —> transfect recombinant DNA into cells

2.  Cells then synthesize a chimeric protein containing GFP fused to protein under study

C.  In all the abovementioned strategies, the labeled proteins take part in normal cell activities —> location can be followed microscopically to reveal dynamic activities in which the protein participates

V.  Applications of fluorescence – simultaneous usage of GFP variants 

A.  Studies can often be made more informative by the simultaneous use of GFP variants that exhibit different spectral properties from one another

1.  GFP variants that fluoresce in shades of blue (BFP), yellow (YFP) & cyan (CFP) have been generated through directed mutagenesis of the GFP gene

2.  A distantly related red fluorescent tetrameric protein (DsRed) has also been isolated from a sea anemone

3.  Monomeric variants of DsRed (e.g., mBanana, mTangerine & mOrange), which fluoresce in a variety of distinguishable colors, have also been generated by mutagenesis experiments

B.  Example - recently, researchers generated strains of mice whose neurons contained differently colored fluorescent proteins

1.  When a muscle of one of these mice was exposed surgically, investigators could observe the dynamic interactions between the variously colored neurons & the neuromuscular junctions being innervated

2.  They watched as branches from a CFP-colored neuron competed with branches a YFP-colored neuron for synaptic contact with the muscle tissue

3.  When 2 neurons compete for innervation of different muscle fibers, all of the winning branches belong to one of the neurons, while all of the losing branches belong to the other neuron

C.  Another example labeling proteins with 2 different fluorochromes

1.  The dual-label strategy has allowed investigators to follow the simultaneous movements of two different proteins in real time as they occur within the boundaries of a single cellular organelle

VI.  Applications of fluorescence – Fluorescence Resonance Energy Transfer (FRET)

A.  GFP variants have been particularly useful in a technique called fluorescence resonance energy transfer (FRET), which can measure distances between fluorochromes in the nanoscale range

1.  FRET is typically employed to measure changes in distance between two parts of a protein (or between two separate proteins within a larger structure)

2.  FRET can be used to study such changes as they occur in vitro or within a living cell

3.  FRET is based on fact that excitation energy can be transferred from one fluorescent group (donor) to another fluorescent group (acceptor), as long as the 2 groups are very close together (1 – 10 nm)

4.  This transfer of energy reduces the fluorescence intensity of the donor & increases the fluorescence intensity of the acceptor

5.  The efficiency of transfer between 2 fluorescent groups that are bound to sites on a protein decreases sharply as the distance between the two groups increases

6.  Thus, determination of changes in fluorescence of the donor & acceptor groups that occur during a process provides a measure of changes in the distance between them at various stages in the process

B.  Example – 2 different GFP variants (ECFP & EYFP) have been covalently linked to 2 different parts of a cGMP-binding protein kinase (PKG)

1.  In the absence of bound cGMP, the 2 fluorochromes are too far apart for energy transfer to occur

2.  Binding of cGMP induces a conformational change in the protein that brings the 2 fluorochromes into close enough proximity for FRET to occur

3.  FRET can also be used to follow many different processes including protein folding or the association & dissociation of components within a membrane

The Light Microscope: Techniques – Video and Laser Scanning Confocal Microscopy

I.  Video microscopy and image processing – view field electronically & tape using a video camera; video cameras have several advantages over the human eye for viewing specimens

A.  Special types of video cameras (called charge couple devices or CCD cameras) are constructed to be very light-sensitive —> can film field under very low illumination —> less heat damage

1.  Live specimens are easily damaged by heat from light source, so the lower heat of video is good

2.  Fluorescently stained specimens fade rapidly upon exposure to light, so low light important here, too

B.  Video cameras can detect & amplify very small differences in contrast within a specimen so that very small objects become visible

1.  For example, an individual microtubule (0.025 mm in diameter) is far below the limit of resolution of a standard light microscope (0.2 mm) but it can be seen with video microscopy

     C.  Video images can be converted to digital electronic images & processed by computer, thus greatly increasing their information content

1.  Conversion of analog electronic image obtained by a video camera to a digital electronic image like that obtained by a digital camera & its storage on a compact disc is called digitization

2.  Digital images consist of a discrete number of picture elements (pixels) each of which has an assigned color & brightness value corresponding to that site in the original image

3.  Distracting, out-of-focus background in visual field is stored by computer & then subtracted from image containing the specimen —> greatly increases clarity of image

4.  Image brightness differences can be converted to color differences —> makes them much more apparent to eye & also allows mathematical analysis

II.  Laser scanning confocal microscopy - replaces old serial section technology (slice embedded specimen); we now examine sections of specimen without cutting it with knife (like CAT scanner)

A.  Old method - look at individual sections & reconstruct 3D structure in mind's eye or with photos

1.  When a whole cell or section of organ is viewed in scope, you focus up & down through specimen & view the specimen at different depths; different parts of specimen go into & out of focus

2.  Specimens have different planes of focus —> reduces ability to get crisp image

3.  Parts of specimen above & below the plane of focus interfere with light from part in focus

B.  Marvin Minsky (MIT, late 1950s) – invented laser scanning confocal microscope, which produces an image of a thin plane situated within a much thicker specimen

1.  It illuminates specimen with finely focused laser beam that rapidly scans across the specimen at a single depth, illuminating only a thin plane (optical section) within the specimen

2.  Short-wavelength incident light is absorbed by specimen & reemitted at longer wavelength; often used with fluorescently stained specimens; light emitted from section forms image

3.  Light emitted from specimen is brought into focus at a site within the scope containing a pinhole aperture; the aperture & illuminated plane in the specimen are confocal

4.  Light rays from illuminated plane of specimen pass through aperture, while light rays emanating from above or below plane do not participate in image formation; out-of-focus points are invisible 

5.  Objects out of the plane of focus have little effect on image quality of each section, so parts of specimen above & below plane of focus do not degrade image

6.  If desired, images from separate optical sections can be stored in computer & used to reconstruct a 3D model of the entire object that can be viewed from any desired angle

Transmission Electron Microscopy: Principles

I.  Types of electron microscopy – transmission electron microscopy (TEM) & scanning electron microscopy (SEM)

A.  Transmission electron microscopes (TEMs) – form images using electrons that are transmitted (pass) through a specimen

B.  Scanning electron microscopes (SEMs) – utilize electrons that have bounced off the surface of the specimen (see next major section)

II. Background information on transmission electron microscopy (TEM)

A.  Provides much greater resolution than light microscopy – greater resolving power derives from the wave properties of electrons; electron wavelength is much lower than that of visible light

1.  The limit of resolution of a microscope is directly proportional to the wavelength of the illuminating light: the longer the wavelength, the poorer the resolution

2.  Electron wavelengths are not constant like those of light – electron wavelength is related to its speed of travel, which, in turn, depends on the accelerating voltage applied in scope (see equation below)

3.  Standard TEMs operate with voltage range from 10,000 - 100,000 V

4.  At 60,000 V, electron is ~0.05Å - amounts to resolution limit of ~0.03Å using N. A. attainable with light scope; this is shorter than typical distance between centers of atoms in molecule - ~1Å

C.  With standard TEM, resolution is actually about 2 orders of magnitude less than its theoretical limit

1.  This is due to serious EM lens spherical aberration; avoid it with very small N. A. (0.01 - 0.001)

2.  Thus, the practical resolution limit of standard TEMs is in range of 3 - 5Å

3.  The actual limit when observing cell structure is more typically in range of 10 - 15Å

III.  TEM structure - tall, hollow, cylindrical column within which electron beam confined and through which the beam passes & a control console

A.  At top of column is cathode (tungsten wire filament) - when heated, it serves as electron source

1.  High voltage applied between cathode & anode draws electrons from filament & accelerates them in fine beam down column

2.  Column is kept in vacuum (air is pumped out) or beam would not be maintained, since electrons would be prematurely scattered by collision with gas molecules 

3.  Electrons focused on specimen not by glass lenses but by electromagnetic lenses in TEM column wall 

4.  Strength of magnets is controlled by the current provided them, which is determined by the positions of the various dials on the console

5.  EM condenser lenses are placed between electron source & specimen; they focus beam on specimen

B.  Specimen is supported on small, thin metal grid (3 mm dia) that is inserted with tweezers into grid holder, which, in turn, is inserted into EM column & thus placed in the middle of electron beam

1.  Lens focal lengths & strength vary depending on current supplied to them; thus one objective lens provides the entire magnification range delivered by the instrument

2.  The image from the EM objective lens (like LM) serves as the object for an additional lens system

3.  Objective lens image is only magnified ~100 times, but unlike the light scope, there is sufficient detail present in the image to magnify it an additional 10,000 times

4.  By altering current to various lenses of scope —> magnification varies from 1,000 - 250,000 times; not empty magnification, since the resolution of the image is so high

C.  Electrons that pass through specimen are focused on phosphorescent screen at bottom of column

1.  Screen is coated by fluorescent crystals, which emit their own visible light when struck by electrons; light is perceived by eye as image

D.  Image formation in EM depends on differential scattering of electrons by parts of specimen

1.  If no specimen in column —> electrons pass through to screen making field uniformly bright

2.  If specimen is in beam path, its parts scatter electrons, which do not pass through very small aperture at back focal plane of objective lens & do not participate in image; screen just below scatter is dark

3.  Electron scattering by part of specimen is proportional to size of its atoms' nuclei

4.  Because of low atomic number of its atoms (carbon, oxygen, nitrogen, hydrogen), biological material has little intrinsic ability to scatter electrons

E.  To increase electron scattering & obtain required contrast, tissues are fixed & stained with solutions of heavy metals

1.  These metals penetrate tissue and the structure of cells & selectively complex with different parts of organelles (staining not uniform)

2.  Those parts of cell that bind the greatest number of metal atoms allow passage of fewest electrons

3.  The fewer electrons that are focused on screen at given spot, the darker is the spot

4.  If part of specimen has more metal atoms —> more scattering —> spot on screen below dark

5.  If part of specimen has fewer metal atoms —> less scattering —> spot on screen below bright

F.  Lift viewing screen out of way & allow electrons to strike photographic plate under screen —> image of specimen is recorded directly on film

1.  Photographic emulsion on plate is directly sensitive to electrons —> photograph

IV.  Specimen preparation for TEM – as in LM, tissues must be fixed, embedded & sectioned

A.  Fixation is much more critical than for light microscopy since resolution is so much better; it must stop life of cell without significantly altering its structure; relatively minor damage is very apparent

1.  Must use very small tissue pieces (<0.5 mm) to obtain most rapid fixation & least cellular damage

2.  Fixatives - chemicals that denature & precipitate cellular macromolecules (must not form artifact)

3.  Artifacts form if such chemicals cause coagulation or precipitation of materials that had no structure in living cell

4.  Best way to show that a structure is not an artifact is to show that it exists in cells fixed in lots of ways or not fixed at all

5.  Rapidly freeze rather than fix tissue; if cells look same with freezing & fixatives —> no artifacts

B.  Most common EM fixatives – glutaraldehyde & osmium tetroxide (also a stain since it is a heavy metal); often used sequentially to ensure optimal tissue fixation

1.  Glutaraldehyde – 5-carbon compound with an aldehyde group at each end of molecule; aldehydes react with amino groups in proteins, cross-linking the proteins into an insoluble network

2.  Osmium tetroxide – osmium is heavy metal & reacts mostly with fatty acids leading to cell membrane preservation; membrane-bound osmium scatters electrons & makes membranes visible in EM

C.  After fixation, specimen is dehydrated by treatment with alcohol solutions & tissue spaces are filled with a material that supports tissue sectioning

1.  Specimens must be very thin (< 0.1 µm; thickness of ~4 ribosomes) for EM 

2.  Wax cannot be sliced thin enough (rarely sliced thinner than 5 µm; Epon & Araldite (epoxy resins) - most widely used embedding materials; firmer than wax & can be sliced thinner 

D.  Specimen sectioned using ultramicrotome - use cut glass or finely polished diamond face (knife); sections are cut by slowly bringing the plastic block down across an extremely sharp cutting edge 

1.  Sections coming off knife edge are floated onto surface of water in trough contained just behind knife edge —> sections are then picked up with metal specimen grid & dried onto its surface

2.  Float specimen grid on drops of heavy metal solution (primarily uranyl acetate & lead citrate) —> they bind to macromolecules & provides atomic density to scatter electron beam

E.  Can also treat with metal-tagged ABs/other materials that react with specific molecules in tissue section

1.  Such studies are usually carried out on tissues embedded in acrylic resins (e.g., Lowicryl)

2.  Acrylic resins are more permeable to large molecules than are epoxy resins

V.  Cryofixation & the use of frozen specimens – cells & tissues do not have to be fixed with chemicals  & embedded in plastic resins in order to be observed with the EM 

A.  Specimens are often rapidly frozen (cryofixation); just as with fixatives, it stops metabolic processes & preserves biological structure

1.  Since it fixes the cell or tissue without altering the cell's macromolecules, it is less likely to lead to formation of fixation artifacts

B.  The major problem with cryofixation is the formation of ice crystals, which grow outward from sites where nucleation occurs

1.  As ice crystal grows, it destroys fragile contents of cell in which it develops

2.  Best way to avoid ice crystal formation is by freezing specimen so rapidly that crystals do not have time to grow large enough to do damage to cell structure

3.  Water that becomes frozen in this liquid-like state is said to be vitrified

C.  Techniques employed to achieve ultra-rapid freezing

1.  Smaller specimens can be plunged into liquids of very low temperature (liquid propane, boiling point of –42°C ) or placed against metal block cooled by liquid helium (boiling point of –273°C)

2.  Larger specimens (100 µm dia) - expose tissue to high hydrostatic pressure while spraying it with liquid N2 jets to freeze it; high pressure drops H2O freezing point, which lowers ice crystal growth rate

D.  How can frozen block of tissue be prepared & visualized?

1.  After suitable preparation, can section frozen block of tissue with special microtome (cryomicrotome)

2.  Cryosections (frozen sections) - very useful for enzyme studies, whose activities tend to be denatured by chemical fixatives

E.  Frozen sections can be prepared much more quickly than paraffin or plastic sections, thus they are often used by pathologists to examine light microscopic structure of tissues removed during surgery

1.  Can determine whether tumor is malignant while patient is still on operating table

F.  Frozen cells do not have to be sectioned to reveal internal structure – unlike standard electron micrographs, image of cells prepared this way has 3D quality, generated by computer rather than directly by camera

1.  The computer merges a large number of 2D digital images of the cell that are captured as the specimen is tilted at defined angles relative to the path of the electron beam

2.  Method is similar in principle to computerized axial tomography (CAT scans),  which uses multitude of X-ray images taken at different angles to body to generate 3-D image; machinery rotates, not body

3.  The 3D computerized reconstruction is called a tomogram & the technique is called cryoelectron tomography (cryo-ET)

4.  Cryo-ET has revolutionized the way in which nano-sized intracellular structures can be studied in unfixed, unstained, fully hydrated, flash-frozen cells

5.  In a real sense, Cryo-ET & other high resolution computational approaches can provide an important bridge between the cellular & molecular worlds

6.  Cryo-ET can also be used for generating 3-D structures of membrane proteins & purified macromolecules

Transmission Electron Microscopy: Techniques

I.  Negative staining - particularly well suited for revealing subunit organization within a particle

A.  EM is well suited for examining very small particulate materials, including high-MW aggregates like viruses, ribosomes, multisubunit enzymes, cytoskeletal elements, protein complexes

1.  Can resolve shapes of large molecules (nucleic acids, individual proteins) if there is sufficient contrast from their surroundings

2.  One of the best ways to make such substances visible is negative staining

B.  How is negative staining done? - general description

1.  Heavy metal deposits are collected everywhere on specimen grid, except where particles are present

2.  Structure of specimen stands out by its relative brightness on viewing screen

II.  Shadow casting - visualize very small isolated particles; use them as objects to cast shadows

A.  Procedure - place grids containing specimen in sealed chamber, then evacuate with vacuum pump

1.  Chamber contains filament composed of heavy metal (usually platinum) together with carbon 

2.  Heat filament to high temperature —> evaporates & coats accessible surfaces with metallic coat

3.  Surfaces facing filament are covered; opposite surfaces of particles & grid spaces in their shadow remain uncoated & unable to scatter electrons

B.  In EM, shadow areas are bright on viewing screen (do not scatter electrons); metal coated regions dark

1.  Relationship is reversed on photographic plate (negative of image) —> so usually print negative image

2.  Particle appears lit by bright, white light (coated surface); dark shadow is cast by particle

C.  Technique provides excellent contrast for isolated materials & produces 3-D effect

III.  Freeze-fracture replication & freeze etching - well suited for examining membrane interior

A. Once tissue is frozen, freeze-fracture replication is often used to view the ultrastructure of frozen cells

1.  Small pieces of tissue are placed on small metal disk & rapidly frozen as described previously

2.  The disk is then mounted on cooled stage within a vacuum chamber

3.  Knife edge is allowed to strike frozen tissue block —> a fracture plane spreads from contact point, splits tissue into 2 pieces, follows path of least resistance (often through center of membranes)

4.  Cell structures deviate fracture plane upward or downward so surfaces are irregular & fracture face has elevations, depressions, ridges that reflect contours of protoplasm traversed

5.  Fractured surface used to form replica (replication) by depositing heavy metal layer (still in fracture chamber); metal deposited at angle —> shadows accentuate local topography (shadow casting)

6.  Uniform carbon layer is deposited on top of metal layer from directly overhead (no shadow) —> cements patches of metal into solid layer —> metal-carbon replica is viewed in scope

7.  Tissue template is thawed, removed & discarded before viewing replica

B.  Variations in thickness of the metal in different parts of the replica produce the necessary contrast in the image by allowing differential electron penetration to the screen & the film

C.  Fracture planes take path of least resistance through block; often go through center of cell membrane; studies by Daniel Branton et al. played role in development of fluid mosaic model in early 1970s

1.  Technique very well suited to examine integral membrane protein distribution as they span bilayer

D.  Freeze etching enhances freeze-fracture information - delivers very high resolution; cell parts seen in deep relief; can reveal shapes of macromolecular complexes as they are presumed to exist in cell

1.  Frozen, fractured tissue, still in cold chamber, is exposed to vacuum at elevated temperature for one to a few minutes —> during this time, a layer of ice evaporates (sublimes) from surface

2.  Specimen surface is coated with heavy metal & carbon after some ice is gone —> view in scope

3.  Metallic replica thus produced reveals both the external surface & internal structure of cell membrane

E.  In deep-etching techniques, greater amounts of surface ice are removed & one gets a fascinating look at cell organelles; individual parts of cell stand out in deep relief against the background

1.  It delivers very high resolution & can be used to reveal the structure & distribution of macromolecular complexes, like the cytoskeleton, as they are presumed to exist within the living cell

Scanning Electron and Atomic Force Microscopy

I.  Scanning EM (SEM) – used to view surfaces of objects ranging in size from virus to an animal head

A.  Uses electrons that have bounced off of specimen surface

1.  Get view in great clarity & detail of the object surfaces from size of virus to animal head

2.  Goal of specimen preparation is to produce an object that has the same shape & surface properties as the living state, but it must be totally dehydrated (devoid of fluid)

3.  Since the specimen will be observed in a vacuum, it cannot have water, hence the dehydration

B.  Since water is a high percentage of the weight of a living cell & it is associated with virtually every cellular macromolecule, its removal can have a very destructive effect on cell structure

1.  When cells are simply air-dried, destruction results from surface tension at air-water interfaces

2.  SEM specimens are fixed, passed through a series of alcohols & dried by process of critical-point drying

II.  Preparation of specimen by critical-point drying - method of drying without damaging cell or specimen

A.  Takes advantage of the existence of a critical temperature & pressure for each solvent in a closed container at which vapor & liquid densities are equal —> no surface tension between gas & liquid

B.  The solvent of cells is replaced with a liquid transitional fluid (usually CO2) vaporized under pressure

C.  Cells are therefore not exposed to any surface tension that might distort their 3-D structure

D.  After drying, the specimen is coated with a thin layer of metal (usually gold or gold-palladium), which makes it a suitable target for an electron beam

III.  Image formation - SEM image formation is indirect

A.  In TEM, the electron beam is focused by condenser lenses to simultaneously illuminate the entire viewing field

B.  In SEM, electrons are accelerated as a fine beam (as small as 5 nm dia) that scans specimen

1.  Image is formed by electrons that are reflected back from specimen (back-scattered) or

2.  By secondary electrons given off by specimen after it is struck by the primary beam

C.  Reflected or secondary electrons strike a detector that is located near the surface of specimen so image formation in SEM is indirect

D.  Another moving electron beam simultaneously scans the face of cathode-ray tube, producing an image similar to that seen on a television screen

1.  Electrons bouncing off the specimen & striking detector are responsible for strength of the signal to the beam in cathode-ray tube

2.  The more electrons collected from the specimen at given spot —> the stronger the signal to the tube & the greater intensity of the beam on the screen at the corresponding spot

3.  Image reflects surface topology of specimen because topology (crevices, hills, pits) determines the electrons collected from various parts of surface

IV.  Notable properties of SEM

A.  Great range of magnification (from ~15 to 150,000 times for standard instrument)

B.  Resolving power is related to the diameter of electron beam – newer models deliver resolution of <5 nm, which can be used to localize gold-labeled ABs bound to a cell's surface

C.  Remarkable depth of focus (~500X that of light microscope at corresponding magnification)

1.  This gives SEM images their 3-D quality & allows visualization & appreciation of outer cell surface structure & the various processes, extensions & extracellular materials that interact with environment

V.  Atomic force microscope (AFM) – not an electron microscope; it is a high resolution scanning instrument that is becoming increasingly important in nanotechnology & molecular biology

A.  AFM operates by scanning a delicate probe over the surface of the specimen being observed

1.  Associated with the probe is a tiny moveable beam (or cantilever) that is displaced by variations in the topography of the specimen 

2.  The degree of displacement correlates with differences in the height of portions of the specimen

B.  The instrument generates a nanoscale map of the displacement, which is translated into a 3-dimensional topographic image of the surface of the specimen

C.  Unlike other methods of molecular structure determination, like X-ray crystallography & cryo-ET, which average structure of many individual molecules……

1.  AFM provides an image of each individual molecule as it is oriented in the field

The Use of Radioisotopes

I.  Tracer - substance that reveals its presence in one way or another & thus can be localized or monitored during an experiment

A.  Such labeled groups enable a molecule to be detected without affecting the specificity of its interactions

1.  Radioactive molecules participate in the same reactions as nonradioactive species, but their location can be followed & the amount present can be measured

B.  Types of tracers

1.  Electron-dense tracers

2.  Fluorescently-labeled tracers

3.  Spin-labeled tracers

4.  Density-labeled tracers

5.  Radioactively labeled tracers

II.  Isotopes – an atom's identity & its chemical properties are determined by number of positive protons in nucleus; all hydrogen atoms have 1 proton, helium atoms have 2 protons & lithium atoms 3 protons

A.  Isotopes are atoms with the same number of protons & varying number of neutrons – not all hydrogen, helium or lithium atoms, however, have the same number of neutrons 

1.  Example - H, the simplest element, has 3 isotopes: 1H-normal (0 neutrons), 2H-deuterium (heavy; 1 neutrons), 3H-tritium (radioactive; 2 neutrons)

B.  Isotopes are sometimes radioactive - when they contain an unstable combination of protons & neutrons

1.  Instability gives atom tendency to break apart or disintegrate to achieve a more stable configuration

2.  With this disintegration, energy is released as a particle or electromagnetic radiation that can be monitored by appropriate instruments

3.  Radioactive atoms occur throughout periodic table; radioactive isotopes can be made from nonradioactive elements in laboratory

4.  All kinds of biological molecules are available in a radioactive state with ≥1 radioactive atom in its structure

C.  Most important aspects of a radioactive isotope, other than the identity of the element are:

1.  Type of radiation emitted

2.  Energy of the radiation

3.  Half-life of the isotope

D.  Three main forms of radiation can be released by atoms during disintegration:

1.  Alpha particle - 2 protons + 2 neutrons; equivalent to helium atom nucleus (least energetic) and/or

2.  Beta particle - equivalent to an electron and/or

3.  Gamma radiation - electromagnetic radiation or photons (most energetic)

III.  Beta particles – the most commonly used isotopes; emitted by most biologically important isotopes; easiest to detect & quantitate accurately

A.  Beta emitters are monitored by either of 2 different methodologies: liquid scintillation spectrometry & autoradiography

1.  Liquid scintillation spectrometry – radioactive sample is mixed in vial with scintillation fluid, which contains compounds that emit light if struck by a beta particle

a.  Amount of light emitted in scintillation fluid is a measure of the amount of radioactivity in sample

2.  Autoradiography – used when one wants to know where a particular isotope is located in a cell, a polyacrylamide gel, or on nitrocellulose filter

B.  Half-life (t1/2) - measure of an isotope's instability (length of time needed for half of it to break down)

1.  The more unstable a particular isotope is —> the greater the likelihood that a given atom will disintegrate in a given amount of time

2.  With 1 Curie of tritium, half that amount of radioactive material will be left after ~12 years (tritium's half-life; a Curie is the amount of radioactivity required to yield 3.7 x 1010 disintegrations/sec

3.  In early metabolic pathway studies (photosynthesis), the only available carbon radioisotope was 11C; experiments using this isotope were carried out very quickly because of its short half-life (20 min)

4.  When longer half-life isotopes became available, like 14C in 1950s (half-life of 5700 years), research became somewhat easier

IV.  Autoradiography - use if want to know where given isotope is located (in cell, polyacrylamide gel, nitrocellulose filter); a broad-based technique 

A.  Its importance was demonstrated in early discoveries on the synthetic activities of cells, the pulse-chase experiments used to follow secretory proteins through cell

B.  Autoradiography procedure – takes advantage of the ability of a particle emitted from radioactive atom to activate photographic emulsion, much like light or X-rays activate emulsion on piece of film

1.  If photographic emulsion is brought into close contact with radioactive source, the particles emitted by the source leave tiny, black silver grains in the emulsion after photographic development

2.  Autoradiography is used to localize radioisotopes within tissue sections that have been immobilized on a slide or TEM grid

3.  Photographic emulsion is applied to the sections on slide or grid as a very thin overlying layer

4.  The specimen is put into lightproof container to allow emulsion to be exposed by emissions

5.  If emulsion is left in contact with isotope for longer time —> get more exposed silver grains

6.  Observe slide or grid in scope —> silver grains are located in emulsion above radiolabeled tissue/cells 

Cell Culture

I.  Why use cultured cells in research? – study cells & cell function in simplified, controlled, in vitro system; can remove cells from influences they are normally subject to within a complex multicellular organism

A.  Cultured cells can be obtained in large quantity

B.  Most cultures contain only a single cell type; a wide variety of different cells can be grown in culture

C.  Many different cellular activities can be studied in cell culture, including endocytosis, cell movement, cell division, membrane trafficking, macromolecular synthesis

D.  Cells can differentiate in culture (conversion of embryonic cells to highly specialized cells)

E.  Cultured cells respond to treatment with drugs, hormones, growth factors & other active substances

II.  First vertebrate cell culture (1907) - over time, optimal conditions were determined & contamination by microorganisms eliminated

A.  Early culture media included a great variety unknown substances; even today, most media contain serum

1.  Cell growth was accomplished by adding fluids from living systems to culture (lymph, blood serum, embryo homogenates)

2.  Cells were found to need a variety of nutrients, hormones, growth factors & cofactors to remain healthy & proliferate; even today, most culture media contain large amounts if serum

B.  Cell culturists are trying to develop defined, serum-free media that support cell growth

1.  They have tested combinations of various ingredients for ability to support cell growth & proliferation

2.  A growing number of cell types have been successfully cultured in artificial media that lack serum or other natural fluids

  3.  Composition of these chemically defined media is a relatively complex mix of nutrients, vitamins, a variety of purified proteins (insulin, epidermal growth factor, transferrin [provides cells with iron])

C.  Since they are so rich in nutrients, tissue culture media invite the growth of microorganisms

1.  Culturists must go to extreme lengths to maintain sterile conditions within their working space

2.  They use sterile gloves, sterilize all supplies & instruments, employ low antibiotic levels in media and conduct activities beneath a sterile hood

III.  Process of cell culture: secondary culture - get cells (often previously cultured cells frozen in liquid N2), thaw them & place them in medium; called secondary culture since cells are derived from a previous culture

IV. Process of cell culture: primary culture – get cells directly from organism, most animal cell primary cultures are from embryos

A.  Embryo tissues are more readily dissociated into single cells than those of adults

B.  Take tissue from embryos & treat with proteolytic enzyme, like trypsin - digests extracellular domains of proteins that mediate cell adhesion

C.  Wash tissue free of enzyme & usually suspend cells in saline solution lacking Ca2+ ions & containing a substance like ethylenediamine tetraacetate (EDTA), a chelator that binds (chelates) Ca2+ ions

1.  Ca2+ ion removal greatly facilitates cell separation since it plays a key role in cell-cell adhesion

D.  Once cells are in single-cell suspension, two kinds of primary cultures can be started: mass culture & clonal culture 

V.  Types of primary cell culture – mass & clonal culture

A.  Mass culture – a relatively large number of cells is added to culture dish

1.  Cells settle & attach to dish bottom & form relatively uniform cell layer; surviving cells grow & divide 

2.  After a number of generations the cells form cell monolayer that covers the bottom of dish

B.  Clonal culture – a relatively small number of cells is added to a dish so that each cell resides at some distance from its neighbors

1.  Under these conditions, each surviving cell proliferates to form a separate colony or clone whose members are all derived from the same original cell

VI.  Normal (nonmalignant) cells can only divide a limited number of times (~50-100) before they senesce & die

A.  Thus, many cells commonly used in tissue culture studies have undergone genetic modifications

1.  These modifications allow them to grow indefinitely (such cells are called a cell line)

2.  They typically can grow into malignant tumors, if injected into susceptible laboratory animals

B.  Frequency with which normal cells are transformed into cell line depends on organism from which they are derived —> mouse cells transform at relatively high frequency; human cells only rarely if ever

1.  Human cell lines (e.g., HeLa cells) are typically derived from human tumors or from cells treated with cancer-causing viruses or chemicals

VII.  Many different types of plant cells can also be grown in culture 

A.  In one approach, culturists can treat cells with cellulase, which digests away surrounding cell wall, releasing naked cell (protoplast)

1.  Grow protoplasts in special, chemically defined, medium that promotes their growth & division

2.  Under suitable conditions, cells can grow into an undifferentiated clump of cells (callus)

3.  Callus can be induced to develop shoots from which a new plant can regenerate

B.  In alternate approach, leaf tissue cells can be induced by hormone treatment to lose their differentiated properties & develop into callus material —> transfer callus to liquid media & start cell culture

The Fractionation of a Cell's Contents by Differential Centrifugation

I.  What is differential centrifugation? – a technique used to isolate a particular organelle in bulk so that its function can be studied or so that an enzyme can be isolated from it; it depends on the principle that:

A.  As long as organelles are more dense than the surrounding medium, they will move to the bottom of a centrifuge tube when placed in a centrifugal field but……..

B.  Since they have different sizes & shapes, they will move to the bottom of that tube at different rates 

II.  Procedure 

A.  First break cells open - mechanical disruption in isotonic buffer (often contains sucrose) with mechanical homogenizer; isotonic buffer prevents rupture of membrane vesicles due to osmosis

B.  Subject homogenate to series of sequential centrifugations at increasing centrifugal forces (often use ultracentrifuge)

1.  First, expose homogenate to low centrifugal force for short period of time —> only the largest cellular organelles (nuclei & any remaining whole cells) are sedimented into pellet

2.  Successively greater forces, bring down smaller organelles (mitochondria, chloroplasts, lysosomes, peroxisomes) —> microsomes (vacuolar & reticular cytosol membrane fragments)  —> ribosomes

3.  A speed of 75,000 rpm & a force of 500,000 times that of gravity in an ultracentrifuge, brings down ribosomes

4.  Postribosomal supernatant contains cell's soluble phase & those particles too small to be removed by sedimentation

C.  Initial differential centrifugation steps are crude —> further steps are required so one must purify more (sucrose density-gradient centrifugation)

1.  Centrifuge crude preparation through sucrose (or some other type of) density gradient

2.  Distributes the contents of the sample into various layers according to the components' densities

3.  Composition of various fractions can be determined by microscopic examination or by measuring the amounts of particular proteins known to be specific for particular organelles

D.  Organelles thus purified, keep very high level of normal activities (if conditions do not denature them)

1.  Isolated organelles can be used in cell-free systems to study a wide variety of cell activities

2.  Examples of activities that can be studied: synthesis of membrane-bound proteins, formation of coated vesicles, solute transport, ionic gradient development, oxidative phosphorylation, etc.

Isolation, Purification and Fractionation of Proteins: Background Information

I.  To study a protein's fine structure, it must be isolated in a relatively pure state – most cells contain thousands of proteins, some at low concentration, so this is a challenging task

II.  Purification is generally accomplished by the stepwise removal of contaminants

A.  Two proteins may be very similar in one property, like overall charge, but very different in others, like molecular size or shape

B.  Successive techniques that take advantage of different properties of proteins being separated are used

C.  Start with methods that work best with high protein concentrations, then move to more sensitive ones

III.  Measure purification as an increase in specific activity (ratio of amount of protein of interest to the total amount of protein present in the sample)

A.  Some identifiable feature of the protein allows determination of its relative amount in sample (assay)

1.  Catalytic activity may be used as an assay to monitor purification if it is an enzyme

2.  Also use assays of immunologic properties, electrophoretic properties, electron microscopic properties, binding activity, etc. and other criteria

B.  Total protein concentration is determined by a variety of tests for protein, including total nitrogen (can be very accurately measured & quite constant at ~16% of dry weight for all proteins)

Isolation, Purification and Fractionation of Proteins: Selective Precipitation

I.  First step in purification should be one that is good for highly impure preparations & able to yield a large specific activity increase

A.  Usually, the first purification step uses solubility differences by selectively precipitating desired protein

B.  Protein solubility properties are determined largely by the distribution of hydrophilic & hydrophobic side chains on its surface

C.  Protein solubility in a given solution is based on the relative balance between protein-solvent & protein-protein interactions

1.  Protein-solvent interactions —> keep proteins in solution

2.  Protein-protein interactions —> cause proteins to aggregate & precipitate from solution

II.  Salt most often used is ammonium sulfate (extremely soluble in water & has high ionic strength)

A.  Gradually add saturated ammonium sulfate solution to crude protein extract

B.  As [salt] increases, precipitation of contaminating proteins increases —> discard precipitate

C.  Eventually, protein being sought precipitates; indicated by loss of activity in assay of soluble fraction

D.  Once desired protein precipitates, contaminating proteins are left behind in solution; one can then redissolve the pellet containing the desired protein

Isolation, Purification and Fractionation of Proteins:  Liquid Column Chromatography

I.  Chromatography – term for a variety of techniques in which a mixture of dissolved components is fractionated as it moves through a porous matrix

A.  In liquid chromatography, components in mixture can associate with 1 of 2 alternative phases: 

1.  Mobile phase, consisting of a moving solvent; liquid chromatography is distinguished from gas chromatography in which the mobile phase is represented by an inert gas

2.  Immobile phase, consisting of the matrix through which solvent is moving (often packed into column)

B.  Proteins to be fractionated are dissolved in solvent, passed through column & interact with matrix to varying extents

1.  Materials making up immobile phase contain sites to which the proteins in solution can bind

2.  As individual proteins interact with matrix —> their progress through the column is retarded

3.  If a particular protein has a greater affinity for matrix —> its passage through column is slower 

4.  Different proteins attracted to matrix with different affinities, so they are retarded to different degrees & thus separated

C.  As solvent passes through column, it is collected as it drips out bottom into series of tubes (fractions)

1.  Proteins in mixture with least affinity for matrix appear in first fractions emerging from column

2.  Those with more affinity for matrix come out later or may bind more permanently to column

II.  High - Performance Liquid Chromatography (HPLC) - gives very high resolution, much more than traditional chromatographic procedures

A.  Long, narrow columns are used

B.  Mobile phase is forced through a tightly packed noncompressible matrix under high pressure

III.  Ion-exchange chromatography – it is unlikely that many proteins in a partially purified prep have the same overall charge, since they are large, polyvalent electrolytes; ionic charge is basis of purification

A.  Overall charge of a protein is the sum of the individual charges of its component amino acids

1.  Depends on pH of medium, since the charge of each amino acid R group depends on medium pH

2.  Lower pH & negatively charged groups are neutralized, positively charged groups get more abundant

3.  Increase pH & the opposite happens

B.  For each protein, there is a pH (isoelectric point) at which negative & positive charges are equal - most proteins have isoelectric point below 7; they are neutral at that pH

C.  In ion exchange chromatography, charged groups are covalently attached to inert matrix material (cellulose; tiny beads)

1.  Diethylaminoethyl (DEAE)-cellulose - positively charged; binds negatively charged molecules; an anion exchanger

2.  Carboxymethyl (CM)-cellulose - negatively charged; binds positively charged molecules; a cation exchanger

D.  Procedure - starts by applying protein solution to column packed with resin

1.  Allow protein solution to pass through column in buffer whose composition promotes binding of all or some proteins to the resin

2.  Proteins are bound to resin reversibly & can be displaced in stepwise manner by sequential addition of different buffers or a continuous gradient in pH or ionic strength

3.  Generally displace proteins by increasing ionic strength of buffer (adds small ions to compete with the charged groups of the macromolecules for sites on the resin) and/or changing its pH

4.  Proteins are eluted from column in order from least strongly bound to most strongly bound

IV.  Gel filtration chromatography - separates proteins (or nucleic acids) primarily on the basis of their effective size (hydrodynamic radius)

A.  Matrix made of tiny beads packed into column; the solution containing proteins passes through slowly

1.  Beads are composed of cross-linked polysaccharides (dextrans, agarose) of different porosity

2.  Varying porosity allows proteins to diffuse in & out of the beads differentially

B.  Example: protein being purified (with 125 kD molecular mass) is mixed with contaminating proteins (250 & 75 kD) of similar shape, one much larger and one much smaller

1.  Pass mix through column with beads that allow entry of globular proteins of <~200 kD into their interiors (Sephadex G-150)

2.  As it passes through column bed, 250 kD protein cannot enter beads, stays dissolved in moving solvent phase & elutes as soon as preexisting solvent in column (bed volume) has dripped out

3.  The other 2 proteins diffuse into interstices within beads & are retarded in passage through column

4.  As more & more solvent moves through column, the proteins move down its length & out bottom

5.  When they eventually move out of column, they do so at different rates —> smaller species retarded more than larger ones; 125 kD protein is pure since it elutes, while 75 kD protein still in column

V.  Affinity chromatography - takes advantage of unique structural properties of desired protein; contrasts with previous techniques, which used the bulk properties of a protein to effect purification or fractionation

A.  Desired protein can be specifically withdrawn from solution, while all others stay behind in solution

1.  Proteins interact with specific substances; enzymes interact with substrates, receptors with ligands, antibodies with antigens

2.  Covalently attach such an interacting molecule to column's inert, immobilized material (matrix) & pass protein mixture through column —> protein that binds column-linked molecule is retained

B.  Example: attach insulin to agarose beads, pack them into column & pass a crude insulin receptor prep through the column

1.  If conditions favor interaction, receptor binds to insulin on column; contaminants go through

2.  After contaminants have passed through the column & out the bottom, insulin receptor can then be displaced from matrix binding sites by changing ionic composition and/or pH of column solvent

C.  Can achieve near total purification of desired protein in a single step, unlike other procedures

Determining Protein-Protein Interactions

I.  One way to learn about a protein's function is to identify the proteins with which it interacts; several techniques have been used to do this, including affinity chromatography

II.  Another technique uses ABs – consider protein A, which has been identified & purified, that is part of a complex with 2 other proteins in the cytoplasm, proteins B & C

A.  After A is purified, prepare an AB against it & use it as a probe to bind to & remove A from solution

B.  If cell extract is prepared containing A-B-C complex & then incubated with anti-A AB, then binding of antibody to the A protein will usually result in coprecipitation of other proteins bound to A 

C.  In this case, proteins B & C should coprecipitate; they can then be identified by other means

III.  Yeast two-hybrid system – technique most widely used to search for protein-protein interactions; invented in 1989 by Stanley Fields & Ok-kyu Song (SUNY-Stony Brook)

A.  Depends on expression of reporter gene, like -galactosidase (lacZ), whose activity is readily monitored by a test that detects a color change when the enzyme is present in a yeast cell population

1.  lacZ gene expression in this system is activated by a particular protein, a TF) that contains 2 domains, a DNA-binding domain & an activation domain

2.  DNA-binding domain mediates binding to promoter & activation domain mediates interaction with other proteins involved in gene expression activation; both domains needed for transcription to occur

B.  2 different types of recombinant DNA molecules are prepared: 

1.  One has DNA segment encoding TF DNA-binding domain linked to DNA segment encoding a "bait" protein X (a protein already characterized & for which potential binding partners are sought)

2.  The other DNA has a DNA segment encoding TF activation domain linked to DNA encoding an unknown protein Y; assume Y is a protein capable of binding the "bait" protein

C.  Such DNAs (or cDNAs) are prepared from mRNAs by reverse transcriptase; both recombinant DNAs, if expressed in yeast cell, lead to production of hybrid proteins

D.  If produced in cell alone, neither the X- nor Y-containing hybrid proteins activate lacZ transcription

1.  If yeast cell is transfected with both of these recombinant DNAs, the X & Y proteins interact with one another to reconstitute a functional TF that transcribes the lacZ gene

2.  The transcription of lacZ that results can be detected by cell's ability to make -galactosidase

E.  Allows researchers to "fish" for proteins encoded by unknown genes that are able to interact with the "bait" protein

Isolation, Purification and Fractionation of Proteins: Polyacrylamide Gel Electrophoresis

I.  Electrophoresis - techniques depending on the ability of charged molecules to migrate if placed in electric field; a powerful technique widely used to fractionate proteins

A.  Electrophoretic separation of proteins is usually accomplished using polyacrylamide gel electrophoresis (PAGE)

1.  Proteins are driven by applied current through a gelated matrix made of polymers of small organic molecules (acrylamide)

2.  Polymers are cross-linked to form a molecular sieve (may be in the form of a thin slab formed between 2 glass plates or a cylinder formed in a glass tube)

B.  Once polymerized, the gel (slab or tube) is suspended between 2 compartments containing buffer into which opposing electrodes have been dipped

1.  In slab gel, concentrated, protein-containing samples are applied in slots at top of gel (prepared in glycerol or sucrose solution, whose density prevents mixing with buffer in upper compartment)

2.  Apply voltage between buffer compartments & current flows across slab —> proteins move toward electrode of opposite charge 

3.  Usually buffers are alkaline, so proteins have "-" charge & move toward "+" electrode, the anode, at the opposite end of the gel

4.  After electrophoresis, the slab is removed from the glass plates & stained

C.  Relative protein movement in gel depends on size, shape & charge density (charge/unit of mass)

1.  High charge density moves protein through gel more forcefully —> migrates at more rapid rate

2.  Polyacrylamide forms a cross-linked molecular sieve that entangles proteins passing through gel

3.  The larger the protein the more it is entangled in molecular sieve & held back (retarded; migrates more slowly)

4.  Shape is also a factor: compact globular proteins move faster than long fibrous proteins of comparable molecular mass

D.  The acrylamide & cross-linking agent concentrations are also important factors affecting movement 

1.  If acrylamide is less concentrated (down to ~2%) —> less cross-linking —> protein migrates faster

2.  Gel containing 5% acrylamide might be useful for separating proteins of 60 – 250 kDa, while a gel of 15% acrylamide might be useful for separating proteins of 10 – 50 kDa

E.  Follow electrophoresis progress by charged tracking dye migration moving just ahead of fastest proteins

1.  When tracking dye has moved to desired location —> current is turned off & gel removed from its container —> stained to see proteins (Coomassie Blue or silver stain)

2.  If proteins are radiolabeled, can locate them with autoradiography (press gel against X-ray film) or

3.  Slice gel up into fractions to isolate proteins, which retain their biological activity

F.  Alternatively, can transfer proteins with current to nitrocellulose filter placed against gel

1.  Proteins are absorbed onto membrane (filter) surface in same relative positions they occupied in gel

2.  Proteins in individual bands are identified by their interaction with specific antibodies (Western blot)

II.  SDS-PAGE - PAGE usually done in presence of "-" charged detergent, sodium dodecyl sulfate (SDS)

A.  SDS molecules bind in large numbers to all types of proteins —> denature proteins —> lose activity

1.  Electrostatic repulsion between SDS molecules unfolds proteins into similar rod shape; differences in shape are no longer a factor in separation

2.  # of protein-bound SDS molecules is roughly proportional to molecular mass (~1.4 g SDS/g protein)

3.  Thus, all proteins have same charge density & are driven through gel with same force, regardless of size

4.  Large proteins are retarded more by cross-linking than are smaller proteins

B.  Thus, proteins are separated by SDS-PAGE on the basis of a single property - molecular mass 

1.  SDS-PAGE can also be used to determine the molecular mass of various proteins

2.  Can compare band positions of proteins of unknown MW to those of known standards to find MW

III.  Two-dimensional gel electrophoresis (1975) - 2-D separation based on 2 different properties; developed by Patrick O'Farrell at Univ. of Calif., San Francisco; used to fractionate complex mixtures of proteins

A.  First separate proteins by isoelectric point with isoelectric focusing within tube gel

B.  Tube gel is removed & placed on top of SDS-saturated polyacrylamide slab gel & subjected to SDS-PAGE —> proteins move into slab & are separated according to molecular mass

C.  Once separated, individual proteins can be removed from gel & digested into peptide fragments —> fragments can be analyzed by mass spectrometry

D.  Resolution is sufficiently high that virtually all proteins in cell can be distinguished (several thousand)

E.  Due to resolving power, 2D-gel electrophoresis is ideally suited to detect changes in proteins found in cell under different conditions, at different developmental or cell cycle stages or in different organisms

1.  Not suitable for distinguishing among proteins that have high molecular mass (MW), that are highly hydrophobic, or that are present at very low copy numbers per cell

Determining the Structure of Proteins and Multisubunit Complexes:  Mass Spectrometry

I.  Proteomics depends heavily on analysis of proteins by mass spectrometry

A.  Mass spectrometers are analytical instruments used primarily to measure molecular masses, determine chemical formulas & molecular structure & identify unknown substances

B.  Mass spectrometers accomplish these goals as follows:

1.  They convert substances in a sample into positively charged, gaseous ions, which are accelerated through a curved tube toward a negatively charged plate

2.  As ions pass through tube, they are subjected to a magnetic field causing them to separate from one another according to molecular mass (or, more correctly, according to mass-to-charge [m/z] ratio)

3.  The ions strike an electronic detector located at the end of the tube

4.  Smaller ions travel faster & strike detector more rapidly than larger ions, which travel more slowly

5.  The input to the detector is converted into a series of peaks of ascending m/z ratio 

II.  Mass spectrometers have been a favorite of chemists for a while, but only in the last decade have biologists discovered their usefulness - protein biochemists can identify unknown proteins in a matter of hours

III. Steps - proteins usually trypsin-digested, fractionated by liquid chromatography & peptides introduced into mass spectrometer where they are gently ionized & made gaseous by either MALDI & ESI

A.  In matrix-assisted laser desorption ionization (MALDI), protein sample applied as part of crystalline matrix; it is irradiated by laser pulse; laser energy excites matrix & converts peptides into gaseous ions

B.  In electrospray ionization (ESI), an electric potential is applied to peptide solution, causing peptides to ionize & the liquid to spray as a fine mist of charged particles that enter the spectrometer

1.  Because it acts on molecules in solution, ESI is well suited to ionize peptides prepared from proteins fractionated by a widely employed liquid chromatography technique

IV.  Once molecular masses of peptides in sample are determined, the complete protein can be identified by a database search

A.  If protein is not identified unambiguously, one or more of peptides generated by tryptic digestion can be fragmented in a second step & subjected to another round of mass spectrometry

1.  Fragmentation is accomplished in mass spectrometer by collision of peptides with inert gas like argon

2.  The energy of collision breaks peptide bonds to produce a random collection of fragments of the original peptide

3.  Each fragment's amino acid sequence, & hence that of original peptide, can be determined by searching a database 

4.  The database contains the masses of theoretical fragments having every possible sequence of amino acids that can be formed from the proteins encoded by the genome

B.  This two-step procedure (called tandem MS or MS/MS) yields the amino acid sequence of the peptide(s) & hence the unmistakable identification of the protein

1.  MS/MS is so powerful that complex mixtures of hundreds of unknown proteins can be digested & subjected to mass spectrometry; the identity of each protein in mixture can be determined all at once

Determining the Structure of Proteins and Multisubunit Complexes:  X-Ray Diffraction Analysis

I.  Steps in X-ray diffraction (X-ray crystallography) – must get most purified preparation of protein possible

A.  Protein crystals are needed before X-ray crystallography can be used to characterize proteins

1.  Crystal is composed of a regularly repeating arrangement of a unit cell (a single protein)

2.  The ability to make a protein crystal is one of the best guarantees of molecular purity

B.  Protein crystal, once formed, is bombarded with a fine X-ray beam of monochromatic wavelength

1.  Radiation that is scattered (diffracted) by electrons of protein's atoms strikes an electron-sensitive detector placed behind the crystal 

2.  Diffraction pattern produced by the crystal is determined by the structure within protein; atoms in the crystal are in a repeating pattern, 

3.  The large number of molecules in crystal reinforces reflections —> it behaves like one large molecule

II.  Data obtained

A.  Reflection positions & intensities on plates are related mathematically to protein electron densities, since it is the electrons of the atoms that produced them; reflections appear as spots on plates

1.  Because of the way in which the crystal is bombarded, each plate represents a slice of molecule; one must analyze many plates to get a single diffraction pattern

2.  Resolution obtained by X-ray diffraction depends on the number of spots that are analyzed

B.  Spots closer to pattern center result from X-rays scattered at smaller angles from crystal; provide information about grosser aspects of protein (long spacings within molecule)

C.  Spots closer to periphery of photograph contain information about closely spaced aspects of molecules within crystal —> resolution depends on the number of spots analyzed

III.  Example: myoglobin, first protein whose structure was determined by X-ray diffraction; analyzed successively at resolutions of 6, 2 & 1.4 Å, with years elapsing between each completed determination

A.  Since covalent bonds are 1 - 1.5 Å in length & noncovalent bonds are between 2.8 Å & 4Å in length, information gathered on protein depends on resolution achieved

B.  In myoglobin, at 6 Å resolution —> showed manner in which myoglobin polypeptide chain is folded & the location of the heme moiety, but it is not sufficient to show structure within chain

1.  At 2 Å resolution —> groups of atoms could be separated from one another

2.  At 1.4 Å resolution —> positions of individual atoms were determined

C.  To date, the structures of several hundred proteins have been determined at atomic resolution (<1.2 Å) & a handful as low as 0.66 Å

IV.  Over time, technology has improved greatly & has been applied to analysis of larger & larger molecular structures (ex. – structure of ribosome); quite often the biggest challenge is getting usable crystals

A.  It took Max Perutz 22 years to solve hemoglobin structure a task that today might take a few weeks

B.  In most current studies:

1.  Intense, highly focused X-ray beams are generated by synchotrons, which are high-energy particle accelerators that produce X-rays as a byproduct, and

2.  Highly sensitive electronic detectors (charge-coupled devices or CCDs) that provide a digital readout of the diffraction data have replaced photographic plates

B.  Use of these instruments along with increasingly powerful computers allows researchers to collect & analyze sufficient data to determine the tertiary structure of most proteins in a matter of hours

C.  As a result of these advances, X-ray crystallography has been applied to the analysis of larger & larger molecular structures, like the determination of ribosomal structure 

1.  As in the ribosome study, the greatest challenge in this field is obtaining usable crystals

D.  X-ray crystallography is ideally suited for finding soluble protein structure (since they lend themselves to crystallization)

1.  It is less suitable for complex multisubunit structures (ribosomes, proteasomes) or for membrane proteins, where it is hard to get 3D crystals to analyze

V.  To study these structures an alternate technique is often used, which takes advantage of the great EM resolving power & computer-based image processing techniques

A.  There are 2 general approaches to the study of single particles with the electron microscope

1.  The particles are placed  on an electron microscope grid & negatively stained or

2.  Electron cryomicroscopy (cryo-EM), in which the particles are placed on a grid & rapidly frozen in liquid N2 without being fixed or stained

B.  In either case, the grids are placed in the column  of the microscope  & photographs of the particles are taken

1.  Each photo is a 2-D image of an individual particle in the orientation that it happened to assume as it rested on grid

2.  If 2-D images of 1,000s of different specimens in every conceivable orientation are subjected to high-powered computer analysis, 3-D image of particle can be reconstructed at resolution as low as 5 Å

3.  This method is also useful for capturing images of a structure (like a ribosome), at different stages during a dynamic process, like the elongation step of protein synthesis

4.  Using this approach, several of the major conformational changes that occur during each step of translation have been revealed

C.  In addition, atomic resolution structures determined by X-ray crystallography can be fitted into lower resolution EM reconstructions 

1.  This helps to show how the individual molecules that make up a multisubunit complex interact & how they might work together to carry out a particular activity

2.  Used to describe a model of workings of actin-cofilin filament; this reconstruction served as basis for proposed mechanism by which cofilin can induce severing/deploymerization of actin filament

D.  Cryo-EM is also well suited for study of membrane proteins, like the nicotinic acetylcholine receptor 

1.  Such proteins can be closely packed at very low temperatures (e.g., -195°C) into 2D crystalline arrays within the plane of the membrane

2.  The structures were determined from combined, high-resolution EM images of many different protein molecules taken at various angles – called electron crystallography

VI.  X-ray diffraction also important to DNA research (suggested double helix to Watson & Crick)

A.  Use fibers of oriented DNA molecules instead of crystals

B.  Resolution is not at as high a level as with proteins

Purification of Nucleic Acids

I.  DNA purification – very different from steps used to purify proteins due to differences in structure; first step is usually homogenization of cells & isolation of nuclei from which DNA is extracted

A.  Extraction medium is buffered salt solution + detergent, like SDS, that lyses nuclei, releasing DNA; solution viscosity rises as DNA is released; detergent also inhibits nuclease activity found in preparation

B.  The goal of the rest of the procedure is to separate DNA from contaminants, like RNA & protein

1.  Deproteinization usually accomplished by shaking extract with phenol or phenol/chloroform mix

2.  Phenol alone or the mix is an active protein denaturant —> proteins lose solubility & precipitate

3.  Phenol & buffered saline are immiscible, so one centrifuges to separate the phases: DNA (& RNA) are in solution's upper aqueous phase; protein present as precipitate at boundary between phases

4.  Aqueous phase is taken from tube & procedure repeated until no more protein is at phase boundary

5.  Precipitate DNA/RNA with cold ethanol layered on top of aqueous phase (spool DNA out at saline/ethanol interface; in contrast, RNA goes to vessel bottom, settling as flocculant precipitate)

6.  Redissolve DNA & RNase treat it to remove contaminating RNA; then destroy RNase with a protease

7.  Then deproteinize with phenol to remove protease & reprecipitate DNA with ethanol

II.  RNA can be isolated in a similar way, but treat with DNase, instead of RNase, in final purification steps; an alternative procedure for isolating RNA in a single step was published in 1987 (see below)

A.  Homogenize tissues in a solution containing 4 M guanidine thiocyanate

B.  Mix RNA extract with phenol & shake with chloroform (or bromochloropropane)

C.  Centrifuge the suspension, leaving RNA in the upper aqueous phase & both DNA & protein at the interface between the 2 phases

III.  An alternative approach to DNA purification involves the use of membrane or matrices to which DNA will  bind under specific conditions

A.  To do this, cells are lysed in a solution that facilitates selective binding of DNA to the matrix

1.  The lysate is applied to the matrix & contaminants are removed from the DNA by washing

2.  Finally, the DNA is rinsed from the matrix with an elution buffer

B.  These matrices are often set up in tiny columns inside centrifuge tubes, so that binding, washing & elution steps can be accomplished efficiently through the application of centrifugal force

IV.  Separation of DNAs by gel electrophoresis - separate nucleic acids by MW (nucleotide length)

A.  Small RNAs or DNAs (a few hundred nucleotides or less) can be separated by PAGE

B.  Larger nucleotide molecules are separated on agarose gels because of this gel's greater porosity; they have trouble making their way through the cross-linked polyacrylamide

1.  Agarose is a polysaccharide extracted from sea weed

2.  It is dissolved in hot buffer, poured into a mold & gelated by lowering temperature

3.  Lower agarose concentrations (as low as 0.3%) are used to separate larger DNA fragments

C.  Separation of DNA molecules > ~25 kb is generally done by technique of pulsed-field electrophoresis in which the direction of the electric field in the gel is periodically changed

1.  Change in field direction causes DNA molecules to reorient themselves during their migration

D.  Principles for DNA/RNA electrophoresis are similar to those for protein SDS-PAGE - all nucleic acid molecules have similar charge density (the number of negative charges per unit of mass)

1.  All nucleic acids, regardless of length, have an equivalent potential for migrating in electric field with agarose providing resistance

2.  The greater the molecular weight, the more slowly a DNA/RNA fragment travels through the gel

E.  After electrophoresis, the DNA fragments in the gel are visualized by soaking the gel in a solution of a stain like ethidium bromide

1.  Ethidium bromide intercalates into the double helix & causes the DNA bands to appear fluorescent when viewed under UV light

F.  Gel electrophoresis sensitivity is so great that DNA or RNA molecules varying by only a single nucleotide can be separated from one another, a feature that has led to an invaluable method for DNA sequencing

G.  Since rate of migration through a gel can also be affected by a molecule's shape, electrophoresis can be used to separate molecules with different conformations, such as circular & linear forms

Measurement of Protein and Nucleic Acid Concentration by Spectrophotometry

I.  Spectrophotometer - measures amount of light of a specific wavelength absorbed by a solution of protein or nucleic acid; it can be used to determine the amount protein or nucleic acid in a given solution

A.  To make measurement, you place solution in a special, flat-sided quartz container (quartz is used since, unlike glass, it does not absorb UV light; a cuvette) & place cuvette in light beam of spectrophotometer

1.  Amount of light passing through a solution unabsorbed (transmitted light) is measured by photocells on the other side of the cuvette

B.  Photons are absorbed totally so if more light is absorbed, there must be more of the molecule absorbing it; thus, absorption can be used as a sensitive measure of concentration

II.  Only tyrosine & phenylalanine of the 20 amino acids absorb UV light (with an absorbance maximum at ~280 nm); most proteins have a typical percentage of these amino acids

A.  If these amino acids are found at a typical percentage in proteins being studied —> absorbance of UV light by the solution containing the proteins at this wavelength is a measure of protein concentration

B.  But since most proteins are colorless, if one wants to use absorbance of visible light one must use one of a number of chemical assays (Lowry or Biuret) to convert protein solution from colorless to colored

1.  Protein in solution is engaged in a reaction that produces the colored product

2.  Concentration of colored product is proportional to protein concentration as is absorbance

III.  Nucleic acids absorb maximally at 260 nm; it is the wavelength of choice for DNA or RNA concentration measurement

Fractionation of Nucleic Acids

I.  Any systematic fractionation method must exploit differences between members of a mixture to effect separation

A.  Nucleic acid molecules can differ from one another in:

1.  Overall size

2.  Base composition

3.  Conformation

B.  Accordingly, fractionation methods for nucleic acids are based on these features

II.  Solution or suspension stability depends on its components; some things float, some sediment & some things stay in solution (remain stable) indefinitely - why do substances sediment through a liquid medium?

A.  Factors, which determine whether or not a given component will settle through a liquid medium

1.  Size, shape & density of component

2.  Density & viscosity of the medium

B.  Component in solution or suspension will sediment through centrifugal field if it has greater density than surrounding medium —> it is concentrated toward bottom of centrifuge tube

1.  Larger particles sediment more rapidly than smaller particles of similar shape & density

2.  Tendency of sedimentation process to concentrate molecules is counteracted by the effects of diffusion, which causes molecules to be redistributed in a more uniform or random arrangement

3.  Sedimentation depends on rate of diffusion compared to opposing centrifugal force applied

4.  Larger proteins & nucleic acids diffuse more slowly than smaller ones 

C.  With ultracentrifuges, it is now possible to generate centrifugal forces up to 500,000 x g, which is high enough to overcome the effects of diffusion & cause macromolecules to move to bottom of tube

1.  Centrifugation proceeds in a near vacuum to minimize frictional resistance

III.  DNA (& RNA) molecules are extensively analyzed by techniques utilizing the ultracentrifuge

A.  There are two most commonly used centrifugation techniques used to study nucleic acids

1.  Velocity sedimentation

2.  Equilibrium centrifugation 

B.  The rate at which a given molecule moves in response to centrifugal force in a centrifuge is known as its sedimentation velocity

1.  Since sedimentation velocity changes as centrifugal force changes, a given molecule is characterized by a sedimentation coefficient, which is its sedimentation velocity divided by the force

C.  Various macromolecules & their complexes are referred to as having a particular S value – unit S (or Svedberg, after the ultracentrifuge's inventor) is equivalent to a sedimentation coefficient of 10-13 sec

1.  S value alone does not provide molecular mass because the velocity at which a particle moves through a liquid column depends on a number of factors, including shape 

2.  It is a good measure of relative size as long as one deals with the same type of molecule: the 3 E. coli rRNAs (5S, 16S & 23S) have lengths of 120, 1600 & 3200 nucleotides, respectively

IV.  Sedimentation behavior of nucleic acids (Technique 1) – in velocity (or rate-zonal) sedimentation, nucleotide molecules are separated according to nucleotide length

A.  Procedure

1.  Sample layered over solution with increasing sucrose concentration (or some other substance)

2.  The preformed gradient increases in density & viscosity from top to bottom

3.  Molecules move through gradient at a rate determined by their sedimentation coefficient when subjected to high centrifugal forces 

4.  If sedimentation coefficient is larger, molecule moves farther in given time period of centrifugation

B.  Nucleic acids are denser than medium so they continue to sediment as long as centrifuge is on; (centrifugation never reaches equilibrium)

1.  The densest sucrose is ~1.2 g/ml density, while DNA has density of 1.7 g/ml

2.  After prescribed period, tube is removed from centrifuge & its contents are fractionated; the relative positions of the various molecules are determined

3.  Presence of viscous sucrose prevents mixing of tube contents (due to convection or handling) allowing molecules of the same S value to form a band & stay together

4.  Find S values by comparing studied molecule positions to standards with known S values (markers)

V.  Sedimentation behavior of nucleic acids (Technique 2) – in equilibrium (isopycnic) sedimentation, nucleic acids are separated on the basis of their buoyant density; procedure follows below:

A.  Mix highly concentrated heavy metal salt solution (CsCl or Cs2SO4) with DNA

B.  Centrifuge with high force (speed) for extended period (2 - 3 days) to make continuous gradient

1  Heavy cesium ions are slowly driven to bottom of tube, forming a continuous density gradient through the liquid column

2.  Eventually, Cs tendency to be concentrated toward the bottom of the tube is balanced by the opposing tendency of diffusion to redistribute the cesium ions —> the gradient is stabilized

C.  DNA molecules place themselves at the point in the gradient equaling their own buoyant density & they are then no longer subject to further movement

D.  Molecules of equivalent density form narrow bands in tube; very sensitive; can separate on basis of base composition or different isotopes of nitrogen (15N vs. 14N)

Nucleic Acid Hybridization

I.  Nucleic acid hybridization includes a variety of related techniques that are based on the observation that 2 single-stranded nucleic acid molecules of complementary base sequence can form a double-stranded hybrid

A.  Mix many DNAs, some of them are complementary —> complementary DNAs form hybrid; mix usually has 100s of DNA fragments of identical length & overall base composition; differ only in sequence

1.  Assume one DNA fragment among them was a portion of the -globin gene, while all the other fragments contained unrelated gene sequences

2.  There is only one way to distinguish between the -globin gene & the others —> hybridization

B.  For example, mix many single-stranded DNA fragments (the -globin gene & many unrelated genes) & excess -globin mRNAs —> form DNA-RNA hybrids; leaving other DNA fragments single-stranded

1.  Separate hybrids & single-stranded DNA by passing mix through hydroxylapatite under ionic conditions allowing hybrids to bind Ca phosphate salts in column

2.  Nonhybridized DNA molecules pass through unbound

3.  Release hybrids from column by increasing elution buffer concentration

C.  Hybridize complementary single-stranded nucleic acids under conditions (temperature, ionic strength, etc.) favoring formation of hybrids or double-stranded molecules

1.  Done with both hybridizing nucleic acids in solution or one immobilized by adsorption on filter (nylon or nitrocellulose) or as part of chromosome

II.  Typical procedure for hybridization - the Southern blot (named after its developer Edward Southern) 

A.  One population of single-stranded nucleic acids in hybridization experiment is often immobilized in electrophoretic gel – example: electrophorese mixture of genomic DNA restriction fragments down gel

B.  To carry out hybridization, the gel is first treated to render the DNA single-stranded & the single-stranded DNA is then transferred from gel to a nitrocellulose membrane (blotting)

1.  The DNA is fixed onto membrane by heating it to 80°C in vacuum

2.  Once the DNA is attached, the membrane is incubated with a labeled, single-stranded DNA (or RNA) probe capable of hybridizing to a complementary group of fragments

3.  Unbound probe is then washed away & bound probe location determined by autoradiography

C.  Method is called Southern blot; one or a few restriction fragments containing a particular sequence can be identified in Southern blot; this is true even if there are 1000s of unrelated fragments in gel

D.  RNA molecules separated by electrophoresis can also be identified with labeled DNA probe after being blotted onto a membrane – Northern blot

III.  Procedure for labeling DNA probes –  can be done in a variety of ways

A.  A radioactive probe incorporates a radioactive isotope (like 32P) at one or more locations in the molecule; the presence of the probe is detected by autoradiography

B.  Probes can also be labeled with fluorophores & detected by fluorescence

C.  Another commonly used label is biotin, a small organic molecule that can be covalently linked to the DNA backbone

1.  Biotin is detected by the protein avidin (or streptavidin), which binds tightly to it

2.  The avidin itself must be labeled for detection, such as with a fluorophore

IV.  Nucleic acid hybridization can also give a measure of the similarity in sequence between 2 DNA samples, for example, from two different organisms

A.  The more distant the evolutionary relationship between 2 species, the greater is the divergence of their DNA sequences

B.  Mix purified DNA from 2 different organisms, A & B, denature it & allow it to reanneal, a percentage of the DNA duplexes are formed by DNA strands from the 2 species

1.  Since they contain mismatched base pairs, such duplexes are less stable than those formed from DNA strands of same species; instability is reflected in lower temperature at which they melt

2.  When DNAs from different species are allowed to reanneal in different combinations, Tm or melting temperature of hybrid duplexes provides a measure of evolutionary distance between the organisms

C.  Two other types of nucleic acid hybridization protocols are discussed previously in text: in situ hybridization & hybridization to cDNA microarrays (discussed in earlier chapters)

Chemical Synthesis of DNA

I.  Hybridization analysis requires single-stranded nucleic acid molecules for use as probes

A.  Other fundamental techniques for DNA manipulation & analysis in lab also require short, single-stranded nucleic acid molecules or oligonucleotides

B.  Chemical synthesis of DNA & RNA is therefore a key supporting technology for many procedures

II.  The chemical reactions have been automated & oligonucleotide synthesis is now carried out by computer-controlled machines hooked to reservoirs of reagents

A.  The operator enters the desired nucleotide sequence into the computer & keeps the instrument supplied with materials

B.  The oligonucleotide is assembled one nucleotide at a time from the 3' end to the 5' end of the molecule, up to a total of ~100 nucleotides

C.  Modifications such as biotin & fluorophores can be incorporated into the molecules

D.  If a double-stranded molecule is needed, it is synthesized as 2 complementary single strands that can be hybridized together

E.  Longer synthetic molecules are made in segments which are joined together

III.  Development of chemical techniques devised to synthesize polynucleotides having a specific base sequence began in early 1960s as part of an attempt to decipher the genetic code by H. Gobind Khorana

A.  Khorana et al. (1970s) – synthesized a complete bacterial tyrosine tRNA gene (126 bp), including the nontranscribed promoter region

1.  The gene was put together from >20 segments: each of these segments was synthesized individually & later joined enzymatically

2.  The artificial gene was then introduced into bacteria having mutations for this tRNA —> replaced the previously deficient function

B.  Keiichi Itakura et al. (City of Hope Med. Ctr., 1977) – synthesized the gene for somatostatin (a small, 14 amino acid residue hypothalamic peptide hormone) 

1.  Inserted gene into specially constructed plasmid downstream from bacterial regulatory sequences

2.  Introduced it into E. coli, where it was transcribed & translated

C.  1981 – gene for the first average-sized protein, human interferon, was synthesized; required synthesis & assembly of 67 different fragments to produce a single duplex of 514 bp

1.  Made whole gene containing initiation & termination signals recognized by bacterial RNA polymerase

D.  Edward Wimmer & colleagues (SUNY-Stony Brook, 2002) – synthesized an infectious poliovirus from scratch

1.  Obtained 7741-nucleotide sequence of RNA virus from public databases & ordered a series of oligonucleotides & assembled them to create a DNA copy of the virus genome

2.  They introduced the DNA into cell extracts where it was transcribed & translated

3.  The viral RNA & proteins assembled into particles capable of infecting & killing mice

4.  Announcement was hailed as both a technical achievement & criticized as a possible guidepost for terrorists seeking to generate biological weapons

5.  Ensuing debate led the editors of scientific journals to agree to consider security implications of research before making it public

Recombinant DNA Technology: Cloning – Background Information

I.  What is recombinant DNA? - a piece of DNA containing sequences derived from >1 source

A.  Over last 30 years, it has allowed advances in analysis of eukaryotic genome

B.  Can be used in myriad ways

II.  Before discussing the uses of recombinant DNA, it is important to discuss the major tool that allowed the production of recombinant DNAs – restriction enzymes (type II restriction endonucleases)

A.  Restriction enzymes were discovered in 1970s in bacteria; they are nucleases that recognize short nucleotide sequences in duplex DNA & cleave DNA backbone at specific sites on both duplex strands

1.  They were called this because they function in bacteria to destroy viral DNAs that might enter the cell, thereby restricting the potential growth of the viruses

2.  The bacterium protects its own DNA from nucleolytic attack by methylating the bases at susceptible sites, a chemical modification that blocks the action of the enzyme

3.  Enzymes from several hundred different prokaryotic organisms have been isolated; together, they recognize >100 different nucleotide sequences

4.  Most restriction enzymes recognize restriction sites that are 4 - 6 nucleotides long & make cuts at specific sites on both duplex strands; they are characterized by particular type of internal symmetry

B.  A sequence recognized by the restriction enzyme EcoR1 is:

1.  Segment has twofold rotational symmetry; it can be rotated 180° without a change in base sequence; if one reads sequence in same direction (3' to 5' or 5' to 3') on either strand, base order is same

2.  A sequence with this type of symmetry is called a palindrome

3.  When EcoR1 attacks this palindrome, it breaks each strand at the same site in the sequence, indicated by arrows between the A & G residues

4.  The dots indicate the methylated bases in this in this sequence that protect the host DNA from enzymatic attack

5.  Some restriction enzymes cleave bonds directly opposite one another on the 2 strands producing blunt ends; others, like EcoR1, make staggered cuts

C.  Since a particular sequence of 4 to 6 nucleotides occurs quite frequently simply by chance, any type of DNA is susceptible to fragmentation by these enzymes

1.  The use of restriction enzymes allows DNA of human genome or that of any other organism to be dissected into a precisely defined set of specific fragments

2.  Once DNA from a particular individual is digested with a restriction enzyme, the fragments generated can be fractionated on the basis of length by gel electrophoresis

3.  Different enzymes cleave the same DNA prep into different sets of fragments and the sites within the genome that are cleaved by various enzymes can be identified & ordered into a restriction map

III.  Formation of recombinant DNAs – can be formed in a variety of ways

A.  Example method – DNA molecules from 2 different sources are treated with a restriction enzyme that makes staggered cuts in DNA duplex

1.  Staggered cuts leave short, single-stranded tails that act as sticky ends that can bind to a complementary single-stranded tail on another DNA molecule to restore a double-stranded molecule

2.  Often one DNA fragment is bacterial plasmid (a small, circular, double-stranded DNA molecule that is separate from main bacterial chromosome)

3.  The other DNA fragment comes from human cells after treatment with the same restriction enzyme used to open the plasmid

4.  When human DNA fragments & plasmid are incubated together + DNA ligase, the 2 types of DNAs are H bonded to one another by sticky ends & then ligated to form circular DNA recombinants

5.  Recombinant DNAs also formed from DNA fragments generated by restriction enzymes that make blunt ends; plasmid & restriction fragment ends are later altered to allow them to stick to one another

6.  First recombinant DNAs made by basic method above (1973) – Paul Berg, Herbert Boyer, Annie Chang & Stanley Cohen (Stanford & U. of Ca., San Fran,); birth of modern genetic engineering

C.  Once this is done, you have produced a large number of different recombinant molecules, each of which has a bacterial plasmid & a piece of human DNA incorporated into a DNA circle

1.  If you want to isolate a single human gene like the one that encodes insulin, you have to separate this specific fragment from all of the others

2.  This is done by DNA cloning

IV.  DNA cloning - technique for producing large quantities of a specific DNA sequence; one can make millions of copies of recombinant DNA in a short period of time from one or a few initial copies

A.  DNA to be cloned is first linked to vector DNA, a vehicle for carrying foreign DNA into a suitable host cell, like E. coli; the vector contains DNA sequences that allow it to replicate in host cell

B.  2 types of vectors are usually used to clone DNAs within bacterial hosts after incorporating the desired gene within them

1.  DNA segment to be cloned is joined to a plasmid; then investigators make bacteria pick up plasmid from medium; as bacteria reproduce so does plasmid containing the desired gene

2.  Foreign DNA is inserted into a portion of  (lambda) bacteriophage viral genome; altered virus is allowed to infect bacteria —> many viral progeny containing foreign DNA are produced

C.  Whichever type of vector is used, once the DNA segment is inside a bacterium, it is replicated along with the bacterial or viral DNA & partitioned into the daughter cells or progeny viral particles

1.  The number of recombinant DNA molecules increases in proportion to the number of bacterial cells or viral progeny formed

2.  From a single recombinant plasmid or viral genome inside a single bacterial cell, millions of copies of the DNA can be formed in a short period of time

D.  Once the amount of DNA has been sufficiently amplified, recombinant DNA can be purified & used in other procedures; thus, cloning can be used to amplify a particular DNA sequence & also to purify it

1.  One can isolate a pure form of any specific DNA fragment among a large, heterogeneous DNA molecule population

Recombinant DNA Technology: Cloning Eukaryotic DNAs in Bacterial Plasmids

I.  Foreign DNA is inserted into plasmid (recombinant DNA) & bacteria are transformed & grown in culture

A.  Plasmids used for cloning are modified versions of those found in bacterial cells; like natural counterparts they contain:

1.  An origin of replication

2.  One or more genes that make the recipient cell resistant to one or more antibiotics; antibiotic resistance allows researchers to select for those cells that contain a recombinant plasmid

B.  Recombinant DNAs with different foreign DNAs are added to bacterial culture where they can be taken up by bacterial transformation as demonstrated by Avery, MacLeod & McCarty

1.  In most common technique, they pretreat bacteria with Ca2+ ions, then briefly heat shock them —> after treatment, they are stimulated to take up DNA from surrounding medium

2.  Transformation - recombinant DNA gets into bacteria; usually, only a small percentage of bacterial cells is competent to pick up & retain one of the recombinant plasmids

C.  Once taken up, the plasmid replicates autonomously & is passed on to progeny during cell division

D.  Select for bacteria with plasmid by growing bacteria on antibiotic whose resistance gene is on plasmid

1.  This procedure eliminates bacteria that have not taken up plasmids

2.  Next, those bacteria with plasmids carrying the desired DNA fragment, if any, must be identified

II.  Identification of bacteria containing desired piece of DNA (e.g., insulin) in their plasmids

A.  Plate out plasmid-bearing cells at low density on petri dishes so that all of the progeny of each cell (a clone of cells) remain physically separate from the progeny of other cells

1.  Each cell contains a different piece of foreign DNA, since you started out with lots of recombinant plasmids; once they have grown into separate colonies, search for those with desired gene (insulin)

B.  Look for colonies or phage plaques with insulin gene using replica plating & then in situ hybridization

1.  Replica plating - make plates with representatives of all bacterial colonies in precisely the same position on each dish, one of which is then used to localize the DNA sequence being sought

2.  Replica plating requires that the cells be lysed & the DNA fixed onto the surface of a nylon or nitrocellulose membrane

3.  Once DNA fixed in place, it is denatured to prepare it for in situ hybridization

4.  The membrane is incubated with labeled, single-stranded DNA probe containing the sequence complementary to that being sought 

5.  After incubation, wash off unhybridized probe & locate hybrids by autoradiography

6.  Culture live representatives of identified clones from corresponding sites on original plates & grow into large colonies, which serves to amplify the recombinant DNA plasmid

7.  Extract DNA from cultured cells that have been harvested & readily separate recombinant plasmid from larger chromosome by various techniques, including equilibrium centrifugation 

8.  Treat isolated recombinant plasmids with same restriction enzyme used to form it —> cloned DNA segment is released from rest of vector DNA & isolated from the plasmid by centrifugation

C.  Not practical to search for single human gene since it would require cloning tens of thousands of DNA fragments & hundreds of separate petri dishes —> better to use viral vector for this

Recombinant DNA Technology: Cloning Eukaryotic DNAs in Phage Genomes

I.  Most often bacteriophage lambda () is used to clone eukaryotic DNAs

II.  Lambda genome - linear, double-stranded DNA molecule; ~50 kb in length

A.  Usually use a modified strain with 2 cleavage sites for EcoR1 (breaks genome into 3 large segments)

B.  All information needed for infectious viral growth (infection & cell lysis) is found in 2 outer segments

C.  Dispensable middle piece of DNA can be replaced with foreign DNA of up to ~25 kb

III.  Package recombinant DNA into phage heads in vitro & use them to infect host bacteria; phage DNAs lacking the insert are too short to be packaged

A.  Once inside bacteria, eukaryotic DNA replicates with viral genome & is packaged into new viruses

B.  Cells lyse & release viruses, which infect new cells, etc.; soon form clear spot (plaque) on bacterial lawn; each plaque has millions of phages; each carries 1 copy of same foreign eukaryotic DNA fragment

C.  Identify desired piece of DNA in plaques with replica plating & in situ hybridization, as described above

IV.  Advantages of phage lambda as cloning vector

A.  DNA is nicely packaged in form in which it can be readily stored & easily extracted

B.  Virtually every phage containing a recombinant DNA is capable of infecting a bacterial cell

C.  One petri dish can accommodate >100,000 different plaques & only several 100 bacterial colonies

Enzymatic Amplification of DNA by PCR:  Background

I.  Polymerase Chain Reaction (PCR) - developed by Kary Mullis (Cetus Corporation, 1983); widely used to amplify, cheaply & readily, specific DNA regions without the need for bacterial cells

A.  Many different PCR protocols have been developed for a multitude of different applications in which anywhere from one to a large population of related DNAs can be amplified

B.  PCR amplification is readily adapted to RNA templates by first converting them to complementary DNAs using reverse transcriptase

II.  Simplest protocol – employs a heat-stable DNA polymerase (Taq polymerase), originally isolated from Thermus aquaticus, a bacterium that lives in hot springs at temperatures >90°C (higher than normal)

A.  Mix DNA sample with 4 deoxyribonucleotides & an aliquot of Taq polymerase

1.  Also add a large excess of 2 short synthetic DNA fragments (oligonucleotides) that are complementary to DNA sequences at the 3' ends of the DNA region to be amplified

2.  These short oligonucleotides serve as primers to which nucleotides are added during the following replication steps

B.  Then heat mixture to ~95°C, hot enough to melt (denature) DNA in mix & separate DNA molecules into their 2 component strands

C.  The mix is then cooled to ~60°C —> allows primers to hybridize to the target DNA strands 

D.  Raise the temperature to ~72°C —> allows the thermophilic polymerase to add complementary nucleotides to the 3' end of the primers

1.  As the polymerase extends the primers, it selectively copies the target DNA

2.  Forms new complementary DNA strands

E.  Raise temperature again causing newly formed & original strands to separate from each other

  F.  The sample is then cooled to allow synthetic primers in mixture to bind once again to the target DNA, which is now present at twice the original amount

G.  Repeat cycle over & over again, doubling the amount of the specific DNA region flanked by the bound primers with each cycle

H.  Generates billions of copies of this one specific DNA region from minute amounts in just a few hours using a thermal cycler - used in criminal cases

1.  Thermal cycler automatically changes the temperature of reaction mixture, allowing each step in the cycle to take place 

Enzymatic Amplification of DNA by PCR:  Applications

I.  Amplifying DNA for cloning or analysis

A.  Since its invention, PCR has found many uses

1.  It can generate many copies of a specific DNA fragment prior to cloning the fragment

2.  This is an efficient approach if the target sequence is known in sufficient detail that the nucleotide sequence of 2 complementary primers can be specified

3.  This is particularly helpful in a case where the source DNA is very scarce, since PCR can generate large amounts of DNA from miniscule samples like that in a single cell

B.  PCR has been used in criminal investigations to generate DNA quantities from a spot of dried blood left on crime suspect's clothing or from DNA present as part of single hair follicle left at crime scene

C.  For this purpose, one selects regions of genome for amplification that are highly polymorphic (i.e., vary at high frequency within population) - thus, no 2 individuals will have same-sized DNA fragments

D.  This same procedure can be used to study DNA fragments from well-preserved fossil remains that may be millions of years old

E.  The activity of DNA polymerase in PCR is also employed in DNA sequencing

II.  Testing for the presence of specific DNA sequences

A.  The hybridization of primers that is intrinsic to the PCR reaction can also be used in a form of hybridization analysis; in this case, the PCR reaction itself serves as the detection system

B.  For example, suppose you wish to determine whether or not a tissue sample contains a particular virus – you could answer  this with Southern hybridization or PCR

1.  In PCR, nucleic acid is isolated from the sample & PCR primers complementary to viral DNA are added, along with the other PCR reagents —> the reaction is carried out

2.  If the virus genome is present in the sample, the PCR primers will hybridize to it & the PCR reaction will generate a product

3.  If the virus is not present, the PCR primers will not hybridize & no product will be generated

III.  Comparing DNA molecules

A.  If 2 DNA molecules have the same base sequence, they will yield the same PCR products in reactions with identical primers

B.  This is the premise for quick assays that compare the similarity of 2 DNA samples such as genomic DNA from bacterial isolates

1.  PCR is performed on the samples using several primers, which can be specifically designed or randomly generated

2.  The products are separated by gel electrophoresis and compared

3.  The more similar the sequences of the bacterial genomes, the more similar their PCR products will be

IV.   Quantifying DNA or RNA templates

A.  PCR can also be used to determine how much of a specific nucleotide sequence (DNA or RNA) is present in a mixed sample

B.  One approach to this quantitative PCR uses the binding of a dye specific for double-stranded DNA to quantify the amount of double-stranded product being generated

1.  The accumulation of product is proportional to the amount of template present in the sample

C.  Another approach uses what have been called molecular beacons 

1.  Molecular beacons are short reporter oligonucleotides with fluorochrome bound to one end & quencher molecule on other end that hybridize in the middle of target sequence to be amplified

2.  As long as the short oligonucleotide is intact, the fluorochrome & quencher are close enough in proximity that fluorescense is quenched

3.  When the DNA polymerase synthesizes a new strand of DNA complementary to the template, its exonuclease activity degrades the reporter oligonucleotide

4.  The fluorochrome is thus separated from the quencher & fluoresces

5.  The amount of fluorochrome liberated in a given PCR cycle is directly proportional to the number of template molecules being copied by the polymerase

DNA Sequencing

I.  By 1970, many protein sequences known, but not many DNA sequences; it took much longer to determine DNA sequences than protein sequences - why?

A.  Unlike DNA molecules, polypeptides come in defined & manageable lengths

B.  A given polypeptide species could be readily purified

C.  A variety of techniques were available to cleave the polypeptides at various sites to produce overlapping fragments

D.  The presence of 20 different amino acids having widely varying properties made separation & sequencing of small peptides a straightforward task

II.  Frederick Sanger, et al. (1977) – they determined the first polypeptide amino acid sequence (insulin) in 1950s (~25 years earlier) & now sequenced an entire viral genome (X174; 5375 nucleotides in length)

A.  In 1970, it might have taken >1 yr to sequence relatively short DNA fragments (a few dozen nucleotides)

1.  In those days, sequencing was typically done manually by an individual working at a lab bench who also personally read & interpreted the results of the reactions

B.  15 years later, comparable task could be done in single day – today sequencing usually performed in series of automated procedures at centralized facilities that sequence hundreds of samples every day

1.  The results are interpreted by computer & stored in databases that can be readily analyzed with commonly available software

2.  Not surprisingly, with these technological advances comes an avalanche of sequence data, 

3.  Among these data are compete genomic sequences for the human, dog (a female boxer), domestic cat, chicken, mouse, rat, several insects, fungi & many bacteria

III.  These advances in DNA sequencing were made possible by developments in several areas

A.  Molecular approaches to DNA sequencing

B.  Instrumentation that could be automated

C.  More powerful & widely available computers and

D.  Software for data analysis

IV.  The initial key was the development of approaches for determining the sequence of DNA fragments

A.  This advance itself became possible because of the discovery of  restriction enzymes & the development of cloning technologies

1.  Restriction enzymes - means to make population of small, defined DNA fragments (very important)

2.  DNA cloning technology 

B.  These methods provided the means necessary to prepare a defined  DNA fragment in sufficient quantity to carry out the necessary biochemical procedures

C.  In mid-1970s, 2 new sequencing techniques were developed almost at same time; Frederick Sanger & A. R. Coulson (Med. Research Counc., Cambridge, England) & Allan Maxam & Walter Gilbert (Harvard)

1.  The Sanger-Coulson method used an enzymatic approach – became the most widely used

2.  The Maxam-Gilbert method used a chemical approach

V.  After PCR came along, Sanger-Coulson sequencing scheme & PCR were merged into sequencing method combining biochemistry of Sanger & Coulson with the repetitive cycles & automation of PCR

A.  This so-called cycle sequencing is now the most commonly used method

B.  One begins with a population of identical template molecules, either a PCR product  or cloned DNA fragment

1.  The template DNA is mixed with a primer that is complementary to the 3' end of one strand of the region to be sequenced

2.  If the template is a PCR product, the sequencing primer can be one (but only one) of the PCR primers

3.  Reaction mix also contains heat-stable Taq DNA polymerase, all 4 deoxynucleoside triphosphate precursors (dNTPs) & low conc. of modified precursors dideoxynucleoside triphosphates (ddNTPs)

4.  Each ddNTP (ddATP, ddGTP, ddCTP & ddTTP) has been modified by the addition of a different-colored fluorescent dye to its 3' end

VI.  The steps in the cycle sequencing methods

A.  The sequencing reaction begins, like PCR, by heating the mixture to a temperature that causes the 2 template strands to denature (step 1)

B.  Next, the reaction is cooled so that the primer can hybridize to the template DNA (step 2)

1.  Note that in contrast to PCR, only one primer is present, so that only one of the 2 strands of template DNA can hybridize to a primer

C.  The Taq polymerase adds dNTPs to the end of the primer that are complementary to the template molecule, synthesizing a new complementary strand of DNA (step 3)

1.  Every now & then, the polymerase inserts a ddNTP instead of a dNTP

2.  Dideoxynucleotides lack a hydroxyl group at both their 2' & 3' positions

D.  When one of these nucleotides has been incorporated onto end of growing chain, the lack of 3' OH makes it impossible for polymerase to add another nucleotide, thus causing chain termination (step 4)

1.  Because the ddNTP is present at much lower concentration in the reaction mixture than the corresponding dNTP, the incorporation of the ddNTP is infrequent & random

2.  It may be incorporated near the beginning of one chain, near the middle of another, or not until the end of a third chain

3.  Regardless, when the ddNTP is incorporated, growth of the chain ceases at that point

E.  After the chain extension phase of the reaction is complete, the temperature is raised again to denature the new double-stranded DNA molecules

1.  Cycle of hybridization, synthesis & denaturation is repeated many times

2.  This generates a large population of daughter DNA strands that by now includes molecules in which a ddNTP has been incorporated at every position

3.  For every A on the template strand, for example, there will be daughter molecules that terminate in a ddTTP at that position

4.  When all cycles are complete, the reaction products are separated by electrophoresis on very thin capillary gels (step 5)

F.  High-resolution gel electrophoresis can separate  fragments that differ by only one  nucleotide in length

1.  If, for example, initial region to be sequenced contained 100 nucleotides, then each of the 100 labeled  daughter molecules would migrate to a different point in the gel

2.  The shortest daughter will migrate the furthest, followed by the shortest + 1 nucleotide, followed by the shortest + 2 nucleotides, etc.

3.  Each successive band in the gel would contain the daughter molecules  that are one nucleotide longer than those in the previous band

4.  Since each ddNTP was labeled with a unique fluorescent dye, the color of the band (viewed under UV light) reveals the identity of the terminal nucleotide on each daughter molecule

5.  The order of the colors in gel therefore corresponds to base sequence of the template molecule

VII.  The Sanger-Coulson reactions can be performed without the PCR adaptation, in which case there is only a single round of DNA synthesis

A.  Under these circumstances, it is necessary to use more template DNA

B.  The reactions can also be performed with a labeled primer rather than labeled ddNTPs

1.  This was the original method used by Sanger & Coulson, & the primers were labeled radioactively

2.  In this case, the sequencing must be set up as 4 independent reactions, in which only one of the 4 ddNTPs is added to each

3.  The 4 reaction products are run side by side in adjacent gel lanes  

4.  The template sequence is determined by looking across 4 lanes to determine which lane (A, G, C or T) contains the next daughter molecule

VIII.  Once nucleotide sequence is determined, various software tools can be employed to analyze it 

A.  The amino acid sequence encoded by the DNA can be determined & compared to other known amino acid sequences to provide information about the polypeptide's possible function

B.  The amino acid sequence also provides clues  as to the tertiary structure of the protein, particularly those parts of the polypeptide that may act as membrane-spanning segments of integral membrane proteins

C.  The nucleotide sequence itself can also be compared to other known nucleotide sequences; such comparisons can be used to: 

1.  Assess the evolutionary relatedness or history of the DNA sequences

2.  Identify the DNA fragment just sequenced or 

3.  Compare the genomic features of various organisms or individuals

Recombinant DNA Technology: Background Information and Formation of a DNA Library

I.  DNA libraries - collections of cloned DNA fragments; 2 basic types: genomic & cDNA libraries; DNA cloning is used to produce them

A.  Genomic library – made from total DNA extracted from nuclei; contain all DNA sequences of species

1.  Once the genomic library of a species is available, researchers can use the collection to isolate specific DNA sequences, like those containing the human insulin gene

B.  cDNA library – derived from DNA copies of an RNA population; typically made from mRNAs present in a particular cell type & thus correspond to genes that are active in that type of cell

II.  Production of a genomic library - treat genomic DNA at low enzyme concentration with 1 or 2 restriction enzymes that recognize very short sequences (HaeIII - recognizing GGCC; Sau3A - recognizing GATC)

A.  Low enzyme concentration prevents cleavage of all susceptible sites; only a small percentage cleaved

1.  A given tetranucleotide is expected to occur by chance with such a high frequency that any sizable DNA segment will be sensitive to fragmentation 

2.  DNA is randomly fragmented since the DNA is treated with enzymes under conditions in which most susceptible sites are not cleaved

B.  The partially digested genome is fractionated by gel electrophoresis or density gradient centrifugation

1.  Fragments of suitable size (e.g., 20 kb in length) are incorporated into lambda phage particles

2.  These phage are used to generate the million or so plaques needed to ensure that every single segment of the mammalian genome is represented

C.  Because the DNA is treated with enzymes under conditions in which most susceptible sites are not being cleaved, for all practical purposes, the DNA is randomly fragmented

D.  Once the phage recombinants are produced, they can be stored for later use; it constitutes a permanent collection of all of the DNA sequences in the genome of the species (library)

1.  If one wants a particular sequence, s/he grows phage on bacteria & screens for the presence of that sequence, using in situ hybridization

2.  Each plaque originates from the infection of a single recombinant phage

3.  Randomly cleaved DNA has advantage – it generates overlapping fragments; useful in chromosome walking (analysis of chromosome regions extending out in both directions from particular sequence)

E.  Chromosome walking - depends on such overlapping fragments 

1.  Isolate fragment containing globin gene coding region, label that fragment & use it as a probe to screen genomic library & isolate fragments with which it overlaps

2.  Repeat with new fragments used as labeled probes in successive screening steps, moving along DNA; one gradually isolates a longer & longer part of the original DNA molecule

3.  Using this approach, one can study organization of linked sequences in extended chromosome region

Recombinant DNA Technology: Cloning Larger DNA Fragments in Specialized Cloning Vectors

I.  YACs & BACs are specialized cloning vectors that are used to clone much larger DNA fragments; used if one must clone fragments >20-25 kb; for example, YAC accepts fragments as large as 1,000 kb

A.  Yeast artificial chromosome (YAC) - artificial versions of normal yeast chromosomes with all of the elements needed for it to be replicated during S phase & segregated to daughter cells during mitosis:

1.  One or more replication origins, telomeres at chromosome ends & centromere to which spindle fibers can attach during chromosome separation

2.  A gene whose encoded product allows cells containing the YAC to be selected from those lacking the element

3.  DNA fragment to be cloned

4.  Furthermore, yeast cells can also take up DNA from medium so that, once created, YACs can be easily introduced into yeast cells to make use of the above features

B.  Labs involved in sequencing genomes have relied heavily on an alternate cloning vector, the bacterial artificial chromosome (BAC); can also accept large foreign DNA fragments (up to ~500 kb)

1.  BACs are specialized bacterial plasmids (F factors) that contain a bacterial origin of replication & the genes required to regulate their own replication

C.  BACs have advantage over YACs in high-speed sequencing projects because they can be cloned in E. coli, which readily picks up exogenous DNA

1.  E. coli also has an extremely short generation time

2.  It can also be grown at high density in simple media

3.  Finally, it does not corrupt the cloned DNA through recombination

II.  To make such big fragments, like those used in YACs & BACs - treat with restriction enzymes that recognize particularly long nucleotide sequences (7 - 8 ) containing CG dinucleotides

A.  CG dinucleotides have special functions in the mammalian genome, so they presumably do not appear nearly as often as would be predicted by chance

1.  Thus, they have fewer cut sites & make fewer & longer fragments

2.  Example: Not I - recognizes the 8 nucleotide sequence (GCGGCCGC); it typically cleaves mammalian DNA into fragments several hundred thousand base pairs long

B.  Fragments of this length can then be incorporated into YACs & BACs & cloned within host yeast or bacterial cells

III.  Above discussion dealt with cloning DNA fragments isolated from extracted DNA (genomic fragments)

A.  When working with genomic DNA, one is usually seeking to isolate a particular gene or gene family from among hundreds of thousands of unrelated sequences

B.  In addition, genomic fragment isolation allows one to study a variety of topics:

1.  Regulatory sequences flanking the coding portion of a gene

2.  Noncoding intervening sequences

3.  Various members of a multigene family, which often lie close together in genome

4.  DNA sequence evolution (duplication, rearrangement) by comparing DNA of different species 

5.  Interspersion of transposable genetic elements

Recombinant DNA Technology: Formation of a cDNA Library

I.  Cloning of cDNAs has also been important in the analysis of gene structure & gene expression; allows you to do things that cloning the entire genome does not

A.  cDNA cloning allows identification of sequences active at given time in a particular cell type

II.  To produce a cDNA library:

A.  Isolate mRNA population & use reverse transcriptase to form a population of DNA-RNA hybrids —> then convert DNA-RNA hybrid to double-stranded cDNA population

1.  Nick the RNA of DNA-RNA hybrid with RNase H & replacing the RNA with DNA by using DNA polymerase I

B.  The double-stranded cDNA is then combined with the desired vector (e.g., a plasmid) & cloned

III.  mRNA populations typically contain thousands of different messages, but individual species may be present in markedly different numbers (they exhibit different abundance)

A.  Thus, a cDNA library has to contain a million or so different cDNA clones to be certain that all of the rarer mRNAs will be represented

B.  Also reverse transcriptase is not a very efficient enzyme; it tends to fall off its template mRNA before the copying job is completed; thus, it can be difficult to obtain a population of full-length cDNAs

C.  Clones must be screened to isolate one particular sequence from a heterogeneous population of recombinant molecules

IV.  Analysis of cloned cDNAs serves several functions – it is generally easier to study a diverse set of cDNAs than the corresponding population of mRNAs, since the cDNAs are more stable

A.  One can use the cDNAs to learn about the variety of mRNAs present in a cell

B.  One can determine the percentage of mRNAs shared by two different types of cells

C.  One can determine the number of copies of different mRNAs present in a cell

V.  A single, cloned & amplified DNA molecule is also useful

A.  The cDNA contains only that information present in the mRNA

B.  Thus, comparison between a cDNA & its corresponding genomic locus can provide information on the precise locations of the noncoding intervening sequences (introns) within the DNA

C.  The purified cDNA can be readily sequenced to determine the polypeptide amino acid sequence; a shortcut & very practical

D.  Labeled cDNAs can be used as probes to screen for complementary sequences among recombinant clones

E.  cDNAs also lack introns & thus have an advantage over genomic fragments when one is trying to synthesize eukaryotic proteins in bacterial cell cultures

Recombinant DNA Technology: DNA Transfer into Eukaryotic Cells & Mammalian Embryos

I.  Transduction – viral-mediated gene transfer; a very widely used strategy to get genes into eukaryotic cells, leading to transcription & translation

A.  Strategy is to incorporate desired nonviral DNA into genome of nonreplicating virus & allow that virus to infect cell; virus integrates its genome into cell genome along with nonviral gene

B.  Depending on the type of virus used, the gene of interest can be expressed transiently for a period of hours to days or it can be stably integrated into the genome of the host cell

1.  Stable integration is usually accomplished using modified retroviruses, which contain an RNA genome that is reverse transcribed into DNA inside the cell

2.  The DNA copy is then inserted into the DNA of the host chromosomes

C.  Retroviruses have been used in many of the recent attempts at gene therapy to transfer a gene into the cells of a patient lacking that gene

1.  Most clinical trials have not been successful due to low infection efficiency of current viral vectors

II.  Methods of transfection - introduction of naked DNA being studied (transgene) into cultured cells

A.  Most often, cells are treated with either calcium phosphate or DEAE-dextran, both of which form a complex with the added DNA that promotes its adherence to the cell surface

1.  Only about 1 cell in 105 takes up the DNA & incorporates it stably into the chromosomes

2.  It is unknown why this small percentage of cells is competent to be transfected, but those that are transfected may pick up several fragments

3.                                   One can select for those cells that have taken up foreign DNA by including a gene that allows transfected cells to grow in a particular medium in which nontransfected cells cannot survive 

4.  Since these cells typically pick up more than one DNA fragment, the selection gene can be on a different fragment from the transgene (the gene whose role is being studied)

B.  Electroporation - cells are incubated with DNA in special vials containing electrodes that deliver a brief electric pulse

1.  Membrane becomes transiently permeable to DNA when current is applied

2.  Some DNA gets into nucleus & is integrated into the chromosomes

C.  Lipofection – cells are treated with DNA that is bound to positively charged lipids (cationic liposomes) that are capable of fusing with the lipid bilayer of cell membrane & delivering the DNA to cytoplasm

III.  Microinjection - inject DNA directly into cell nucleus; oocyte/egg nuclei are particularly well-suited

A.  Xenopus oocytes used to study foreign gene expression; inject foreign DNA —> freely transcribed

1.  Oocyte nucleus contains all the machinery necessary for RNA synthesis; when foreign DNA is injected into nucleus, it is readily transcribed

2.  RNAs synthesized from injected templates are processed normally & transported to cytoplasm, 

3.  In cytoplasm, RNAs are translated into proteins (detected immunologically or by specific activity)

B.  Inject foreign genes into mouse embryo nuclei; goal is its integration into embryo chromosomes & its passage to all cells of embryo & subsequent adult, not to monitor gene expression in injected cell

1.  Genetically engineered animals whose chromosomes carry foreign genes called transgenic animals

2.  They provide means to monitor where & when in embryo particular genes are expressed

3.  Also reveal impact of extra copies of particular genes on the development & life of animal 

IV.  Transgenic animals 

A.  Ralph Brinster (Univ. of Penna.) & Richard Palmiter (Univ. of Wash.), 1981 - introduced gene for rat growth hormone (GH) into fertilized mouse eggs

1.  Injected DNA constructed to have rat GH gene coding region just downstream from mouse metallothionein gene promoter (a strong promoter)

2.  Metallothionein synthesis normally greatly enhanced by metals (cadmium, zinc) or glucocorticoids

3.  Treat mice with metals (cadmium, zinc) or glucocorticoids —> GH induced; mice got very big

B.  Transgenic animals provide mechanism for creating animal models (lab animals exhibiting a particular human disease they would normally not have) – example: mutant human amyloid precursor protein

1.  In transgenic mice carrying a gene encoding a mutant form of human amyloid precursor protein (APP), the mice develop neurological & behavioral symptoms reminiscent of Alzheimer's disease

2.  They are an important resource in the development of therapies for Alzheimer's disease

C.  Transgenic animals also being developed as part of agricultural biotechnology

1.  Incorporate foreign growth hormone genes into chromosomes of pigs —> grow much leaner than control animals lacking gene

2.  Excess growth hormone stimulates conversion of nutrients into protein rather than fat, explaining why they are leaner

V.  Transgenic plants - introduce gene into cultured plant cells —> grow into mature plants with foreign gene; derives from fact that whole plants can be grown from individual cultured plant cells

A.  Agrobacterium tumefaciens can get foreign DNA into plant cells – outside lab, it lives in symbiotic association with dicotyledonous plants & causes tumorous lumps (crown galls) - how does it do this?

1.  During infection, a section of bacterial Ti plasmid (the T-DNA region) is passed from the bacterium into the plant cell; this part of plasmid enters plant cell & incorporates into plant chromosomes

2.  The T-DNA region of the Ti plasmid induces the cell to proliferate & provide nutrients for bacteria 

B.  Ti plasmid can be isolated from bacteria & linked with foreign genes —> make recombinant plasmid

1.  Altered plasmid can be taken up in culture by undifferentiated dicotyledonous plant cells (like carrots, tobacco); called T-DNA transformation

2.  The technique has been used to transform plant cells with genes derived from bacteria that code for insect-killing toxins; they protect plants from insect predators

C.  Other techniques developed to get foreign genes into monocotyledonous plant cells - can serve as targets for microscopic, DNA-coated tungsten pellets fired by gene gun; already used for number of plant cells

VI.  The 2 most important plant genetic engineering goals - improve photosynthesis & N2-fixation; both are crucial bioenergetic functions

A.  Any significant improvement in photosynthetic efficiency —> great increases in crop production if we can engineer modified version of CO2-fixing enzyme that is less susceptible to photorespiration

B.  N2 fixation is carried out by certain genera of bacteria (e.g., Rhizobium); live in symbiotic relationship with certain plants (like soybean, peanut, clover, alfalfa, pea)

1.  Bacteria reside in swellings (leguminous nodules) located on roots, where they remove N2 from atmosphere, reduce it to ammonia & deliver the product to the cells of the plant

2.  Trying to isolate bacterial genes involved in this activity & introduce them into chromosomes of nonleguminous plants (now depend heavily on added fertilizer for reduced nitrogenous compounds)

3.  Identify genes responsible for N2 fixation in Rhizobium  —> deliver to plants —> less fertilizer

C.  Might be possible to alter genome of either plant or bacteria so that new types of symbiotic relationships can be developed

Determining Eukaryotic Gene Function by Gene Elimination:  In Vitro Mutagenesis & Site-Directed Mutagenesis

I.  Until fairly recently, investigators discovered new genes & their function by screening for mutants that exhibited abnormal phenotypes

A.  It was only through the process of random mutation that the existence of genes became apparent; this process of learning about genotypes by studying mutant phenotypes is called forward genetics

B.  Since the development of gene cloning & DNA sequencing techniques, researchers have been able to identify & study genes directly without knowing anything about the function of the encoded protein

C.  Over last 20 years, researchers have developed means to carry out reverse genetics, which is a process of determining phenotype (i.e., function) based on knowledge of genotype 

1.  The basic approach of reverse genetics is to eliminate the function of a specific gene, & then determine what effect the elimination of that function has on phenotype

II.  Isolation of naturally occurring mutants has played an enormously important role in determining the function of genes & their products

A.  But natural mutations are rare events & it is not feasible to use such mutations to study the role of particular amino acid residues in the function of a particular protein

B.  Rather than waiting for organism to appear with  unusual phenotype & identifying responsible mutation, …….

1.  Researchers can mutate gene (or its associated regulatory regions) in a desired way & observe the resulting phenotypic change

C.  These techniques are collectively termed in vitro mutagenesis; they require that the gene, or at least the gene segment, to be mutated has been cloned

III.  One procedure was developed by Michael Smith (Univ. of British Columbia) & is called site-directed mutagenesis (SDM); DNA-synthesis machines can alter the sequences of natural DNA

A.  It allows researchers to make very small, specific changes in a DNA sequence, such as:

1.  The substitution of one base for another or

2.  The deletion or insertion of a very small number of bases

B.  Usually accomplished by first synthesizing a DNA oligonucleotide containing the desired change

C.  Then allow the oligonucleotide to hybridize to a single-stranded preparation of the normal DNA & then use the synthesized oligonucleotide as a primer for DNA polymerase

D.  DNA polymerase elongates the primer by adding nucleotides complementary to the normal DNA

E.  Next, clone the modified DNA & determine the effect of the modification by introducing the modified DNA into an appropriate host cell

1.  If site is part of regulatory region —> it may affect gene expression rate, which can be monitored

2.  If site is part of coding region, it gives insight into site's role in overall protein structure & function

III.  Scientists typically use SDM to ask very targeted questions about the function of a gene or protein

A.  They might change one amino acid into another  to gain insight into the role of that specific site in the overall function of a protein

B.  Alternatively, they might introduce small changes in the regulatory region of a gene & determine the effect on gene expression

IV.  If the intent of SDM is simply to eliminate the function of a gene, less specific methods can be used

A.  For example, one can cut a gene sequence at a restriction site & use DNA polymerase to make the single-stranded regions of the sticky ends into double-stranded DNA 

1.  One can then ligate those ends back together, which can destroy the reading frame of a protein

B.  In other instances, an entire restriction fragment might be removed from the gene

C.  2 widely used techniques for eliminating gene function in vivo

1.  Knockout mice

2.  RNA interference

V.  Constructing a mutation in vitro is only part of reverse genetics

A.  To study the effect of an engineered mutation on phenotype, it is necessary to substitute the mutant allele for the normal gene in the organism in question

B.  Development of a technique for introducing mutations into the mouse genome opened the door for reverse genetic studies in mammals & literally revolutionized the study of mammalian gene function

Determining Eukaryotic Gene Function by Gene Elimination:  Knockout Mice

 I.  Knockout      mice -  scientists                          have produced mice that lack a functional copy of a particular gene to see the resultant phenotype & determine the significance of the missing gene product

A.  For example, p53 knockout mice (lacking a functional p53 gene) – invariably develop malignant tumors

B.  Such knockout mice can give unique insight into human disease's genetic basis & are also mechanism for studying the various cellular activities in which particular gene product might be engaged

II.  The various procedures used to generate knockout mice were developed in the late 1980s by Mario Capecchi (Univ. of Utah), Oliver Smithies (Univ. of Wisconsin) & Martin Evans (Cambridge Univ.)

A.  Isolate embryonic stem cells (ES; have virtually unlimited powers of differentiation) from mammalian blastocyst

1.  Mammalian blastocyst is early embryonic development stage comparable to blastula stage in other animals; it is composed of 2 distinct parts: trophectoderm & inner cell mass (ICM)

2.  Outer blastocyst layer is trophectoderm; it gives rise to most of the extraembryonic membranes characteristic of mammalian embryos

3.  Trophectoderm's inner surface contacts cell cluster (ICM), projecting into spacious cavity (blastocoel)

4.  ICM gives rise to cells that make up embryo & contains ES cells, which differentiate into all of the various tissues of which a mammal is composed 

B.  Isolate blastocyst ES cells & culture them in vitro under conditions encouraging growth & proliferation

C.  Transfect ES cells with DNA fragment containing nonfunctional, mutant allele of gene to be knocked out & antibiotic-resistance genes (used to select for cells that have incorporated altered DNA into genome)

1.  Of those cells taking up DNA, ~1 in 104 undergoes homologous recombination in which transfecting DNA replaces homologous DNA sequence (normal allele)

2.  Cells produced are heterozygous for this gene —> select for them on basis of their drug resistance

D.  Inject some donor ES cells into recipient mouse embryo blastocoel —> implant in hormonally prepared female mouse's oviduct to carry embryo to term;    recipient embryo is from a black strain

1.  As embryo develops in its surrogate mother, injected cells                                             join embryo's own ICM & contribute to formation of embryonic tissues                    , including germ cells           of gonads

2.  Chimeric mice recognized, since their coat has characteristics of both the donor & recipient strains

E.  Mate chimeric mice     to member of inbred black strain —> if their germ cells contain knockout mutation, all cells of their offspring will be heterozygous, which are distinguished by their brown coat coloration 

F.  These heterozygotes are then mated to one another —> some offspring are homozygous for mutant allele & thus lack functional copy of gene —> observe effect on organism

G.  Any gene within the genome, or any DNA sequence for that matter, can be altered in any desired manner using this experimental approach

IV.  Interpretation of observations in gene knockout experiments

A.  Sometimes deletion of particular gene can lead to absence of particular process —> provides convincing evidence that the gene is essential for that process

B.  Often, deletion of a gene thought to participate in essential process causes little or no alteration in animal's phenotype —> difficult to interpret: 

1.  If no phenotype change, it may be that gene is not involved in process being studied or, as is usually the case, absence of gene product is compensated for by product of entirely different gene

2.  Compensation by one gene for another can be verified by producing mice that lack both of genes in question (i.e., a double knockout)

C.  Sometimes, absence of gene leads to death of the mouse during early development, which also makes it difficult to determine the role of the gene in cellular function

1.  Researchers can get around this by using technique that allows particular gene to be knocked out only in one or more desired tissues, while the gene is expressed in the remainder of animal

2.  These conditional knockouts generally survive to adulthood & allow researchers to study the role of the gene in the development or function of affected tissue

Determining Eukaryotic Gene Function by Gene Elimination:  RNA Interference

I.  RNA interference – RNAi is a process in which a specific mRNA is degraded due to the presence of a small, double-stranded RNA (dsRNA) whose sequence is contained within the mRNA sequence

A.  Plant, nematode or fruit fly gene function studied by simply introducing a dsRNA into organism by various means & examining organism phenotypes resulting from depletion of corresponding mRNA

1.  Using this approach, information about the functions of large numbers of genes can be gathered in a relatively short period of time

2.  Less laborious & costly than generating knockout animals & yields essentially same information

B.  RNAi can be used to study mammalian cell gene function by incubating the cells with small dsRNAs encapsulated in lipids or by genetically engineering the cells to produce the dsRNAs themselves

1.  Once inside the cells, the dsRNA leads to degradation of target mRNA, leaving the cell unable to produce additional protein encoded by that gene

2.  Any deficiencies in the cell phenotype can be attributed to a marked reduction in the level of the protein being investigated

C.  Demonstrated by effect on chromosome segregation following transfection of cells with a dsRNA that targets Aurora B kinase

1.  Chromosomes in such a cell lie adjacent to the mitotic spindle, suggesting the absence of kinetochore-microtubule interactions

II.  Libraries containing thousands of small dsRNAs, or vectors containing DNA encoding these RNAs, are also available for the study of human gene function

A.  Researchers using these libraries can study the effects on any given process that results from the elimination of expression of virtually any gene in the genome

B.  These studies have led to new insights into the role of numerous genes whose functions were previously unidentified

The Use of Antibodies: Background

I.  What are antibodies (ABs; also known as immunoglobulins)? - proteins made by lymphoid tissue after antigen (Ag; foreign materials) exposure; they exhibit a high degree of specificity (makes them useful)

A.  AB preparation binds only those select molecules in cell having a small part that fits into Ag-binding site of AB; can select a few proteins out of the thousands in a cell

B.  Antibodies can distinguish between two polypeptides that differ by as little as one amino acid

C.  There are basically 2 approaches to the preparation of ABs that interact with a given Ag – traditional & monoclonal AB approaches

II.  Traditional approach for the production of ABs

A.  Repeatedly inject animal (typically a rabbit or goat) with Ag

1.  After a period of several weeks, blood is drawn that contains the desired ABs

2.  Whole blood is treated to remove cells & clotting factors —> produces an antiserum

3.  Test antiserum for AB titer & purify immunoglobulins (Igs) from it

B.  Traditional approach still used, but it has certain inherent disadvantages:

1.  Due to AB synthesis mechanism, animal invariably makes a variety of different species of Igs (with different V regions in their polypeptide chains; polyclonal) even if the Ag was highly purified

2.  Cannot get a pure preparation of a single AB species with this technique, since the ABs are too similar to be fractionated; the ABs are polyvalent

The Use of Antibodies: Monoclonal Antibodies

I.  Preliminary discoveries leading to production of monoclonal (univalent) ABs

A.  ABs made by a clone of AB-producing cells (derived from single B lymphocyte) have identical Ag-combining sites

1.  Polyvalent AB heterogeneity against a single purified Ag is due to activation of many B cells

2.  Each of the B cells has membrane-bound ABs with an affinity for a different part of the Ag

B.  Theoretically, one could obtain a pure preparation of a single monoclonal AB if 1 AB-producing plasma cell were isolated by a procedure something like the one that follows:

1.  Inject an animal with a purified Ag & wait a period of weeks for ABs to be produced

2.  Then remove the spleen or other lymphoid organs and prepare a suspension of single cells

3.  Isolate those cells producing the desired ABs from the suspension & grow these particular cells as separate colonies so as to obtain large quantities of this particular Ig

C.  This procedure should yield an AB molecule prep made by a single colony or clone of cells, a monoclonal antibody, but……. antibody-producing cells do not grow & divide in culture

1.  To make monoclonal ABs in this way, an additional manipulation is required

D.  Malignant myeloma cells are a type of cancer cell that grows rapidly in culture & makes lots of AB

1.  ABs made by myeloma cells were valuable in AB structure study, but of little use as analytic tools since the ABs produced by them are not formed in response to a specific Ag

2.  Instead, myeloma cells develop from random conversion of a normal lymphocyte to a malignant state; they make AB that was being made by the particular lymphocyte before it became malignant

II.  Monoclonal antibody production - Cesar Milstein & Georges Köhler (Med. Research Counc., Cambridge, Engl., 1975) – did experiments leading to development of univalent AB preps directed against specific Ags

  A.  They combined the properties of normal AB-producing lymphocytes & the immortal myeloma cell

B.  They fused malignant myeloma cells & normal lymphocytes to make hybrid cells —> hybridomas

1.  They grow & proliferate indefinitely, producing large amounts of a single, monoclonal AB

2.  The AB made is the one being made by normal lymphocyte before fusion with myeloma cell

C.  Procedure – do not even need purified Ag; can use soluble Ag or one that is part of cell; it may even be minor component of entire mixture

1.  Ag was injected into mouse to cause proliferation of specific AB-producing cells

2.  After several weeks, spleen is removed & dissociated into single cells 

3.  AB-producing lymphocytes were fused with a population of malignant myeloma cells, making the hybrids immortal (capable of unlimited cell division)

4.  Hybrids were selected from the unfused cells by their ability to grow in a medium in which only they can survive

5.  Hybridomas are then grown clonally in separate wells & individually screened for production of AB against Ag being studied

6.  Clone hybrid cells containing appropriate AB in culture (in vitro) or as tumor cells in recipient animal (in vivo) —> get lots of monoclonal AB, in essentially unlimited amounts

7.  Once produced, they can be stored indefinitely in frozen state & aliquots can be made available to researchers around the world

D.  Do not need to begin procedure with purified Ag; 

E.  Monoclonal ABs are useful in research, and are also useful in diagnostic medicine to determine concentration of specific proteins in blood or urine

1.  Monoclonal ABs form the basis of certain home-pregnancy tests that monitor the presence of a protein (chorionic gonadotrophin) that appears in the urine a few days after conception

III.  No matter how they are obtained, one can use ABs as highly specific probes in a variety of analytic techniques, e.g., localizing proteins in cells

A.  ABs can be used as highly specific probes in protein purification - add purified AB to crude protein mixture  —> specific protein being sought selectively combines with AB & precipitates from solution

B.  ABs can also be used along with various types of fractionation procedures to identify a particular protein (Ag) among a mixture of proteins

1.  In Western blot, a mixture of proteins is first fractionated by 2D gel electrophoresis

2.  The fractionated proteins are then transferred to a sheet of nitrocellulose filter

3.  Filter is then incubated with an AB prep that has been labeled either radioactively or fluorescently

4.  The location on the filter of a specific protein bound by the AB can be determined from the location of the bound radioactivity or fluorescence

C.  Monoclonal ABs have also been useful as therapeutic agents in humans – however, efforts to develop human hybridomas that produce human ABs have been unsuccessful, so………

1.  Mice have been genetically engineered so that ABs they produce are increasingly human in amino acid sequence

2.  Some of these humanized monoclonal ABs have been approved for the treatment of several diseases 

3.  More recently, mice have been engineered so that their immune system is essentially human in nature —>  they produce ABs that are fully human in structure

D.  First fully human antibody (Humira; approved for rheumatoid arthritis treatment); made by very different method using bacteriophage rather than hybridomas to make monoclonal antibodies

1.  The technique is known as phage display

2.  Billions of different phage particles are generated in which each phage carries a gene encoding a human antibody molecule that possesses a unique variable region

3.  Different phage within this vast library encode different antibodies, that is, antibodies with different variable regions

4.  In each case, antibody gene is fused to gene encoding one of viral coat proteins so that when phage is assembled within host cell, the antibody molecule is displayed on the surface of the viral particle

E.  Suppose that you have a protein (antigen) that you believe would be a good target for a particular therapeutic antibody

1.  The antigen is purified & allowed to interact with a sample of each of the billions of phage particles that make up the phage library

2.  Those phage that bind to the antigen with high affinity can be identified & allowed to multiply within an appropriate host cell

3.  Once it has been amplified in this way, the DNA encoding the antibody gene can be isolated & used to transfect an appropriate mammalian cell

4.  The genetically engineered cells can then be grown in large cultures to produce therapeutic quantities of the antibody

5.  Production of antibodies in cultured mammalian cells is an expensive venture & alternative "living factories" are being pursued

6.  Among the alternatives being considered for this purpose are goats, rabbits & tobacco cells

F.  Immunolocalization in cells – visualizing specific molecules in cells using labeled antibodies (see below)

IV.  Immunolocalization in cells depends on the use of ABs made specifically against a particular protein; the ABs are then conjugated to a substance that makes them visible under the microscope (LM or EM)

A.  The binding of the substance making the AB visible does not interfere with the specificity of AB-Ag interactions

1.  Often label ABs with small fluorescent molecules (fluorescein, rhodamine) for light microscopes —> forms derivatives that are incubated with cells or sections of cells

2.  Visualize the AB-binding sites in the fluorescence microscope (direct immunofluorescence)

B.  Types of immunolocalization – direct or indirect immunofluorescence

1.  Direct immunofluorescence - labeled AB is attached directly to Ag in cell

2.  Indirect immunofluorescence – incubate cells first with unlabeled AB (complexes with corresponding Ag); then, expose cell to fluorescent AB directed against AB used in first step; often preferable

C.  Advantages of indirect immunofluorescence

1.  Brighter image than direct method because numerous secondary ABs can bind to single primary AB

2.  Also has practical advantage – conjugated (fluorescent) secondary AB is readily bought from vendors

D.  Immunofluorescence (direct or indirect) provides remarkable clarity since only the proteins bound by the AB are revealed to the eye; all of the unlabeled materials remain invisible

E.  For EM, tag ABs with electron dense materials (iron-containing protein ferritin or gold particles)

Lecture Hints

When I first started teaching Cell Biology, I would discuss each technique when I got to an experiment in which it had played a role or a concept its use had helped to develop.  This resulted in disjointed lectures and frequent bouts of going off on tangents.  Students would often lose the train of thought and would complain about this to me directly or in course evaluations.  I have since chosen to cover techniques (Chapter 18) right after the series of biological chemistry lectures.  This has a number of advantages.  First, the biological chemistry, which plays such an important role in most, if not all, of the methodologies, is fresh in the students' minds.  Furthermore, by covering the techniques together, students have a better opportunity to compare and contrast the information obtained from each technique and they get a sense of how such methodologies may be used in sequence.  It also serves to remove the coverage of techniques as a disruptive influence on later lectures, while, at the same time, grouping them together into a cohesive collection of presentations early in the semester.

The Light Microscope

I usually cover the material on light microscopy and the material that follows it, electron microscopy, in the introduction to my first three laboratory exercises rather than in a traditional lecture setting.  I concentrate on the principles of both light and electron microscopy and some of the specialized techniques employed in both types of microscopy.  The material integrates well with the laboratory exercises we carry out in the microscopy teaching lab and some of the demonstrations I use are more easily and conveniently done there.  Obviously, the material can be easily handled in a traditional lecture.

I begin my discussion of microscopy by describing the components of the light microscope and the basic principles behind light microscopy.  I recommend that you tell your students how to calculate the magnification of a particular image.  Of course, one must simply find the product of the powers of the objective and ocular lenses.  Ask your students their opinion of the most important thing that a microscope does.  The vast majority of students will answer magnification.  Rarely will they give the correct answer - resolution.

Define resolution for the class - the ability to distinguish two objects located near each other as separate objects.  Then describe how the minimum resolving distance is determined (see formula in outline).  Also, emphasize the relative effects of wavelength and numerical aperture (effectively the light gathering ability of the lens) on resolution distance.  Emphasize that it is actually better if this distance is as small as possible, since that would mean that objects closer together can be distinguished.  

Give your class a list of conditions - numerical apertures and colors of light - and ask if they can tell you which conditions would yield the best resolution.  I often ask the class how they would build the best (short wavelength blue light, high N. A.) and worst microscopes (high wavelength red light, low N. A.) possible using these conditions.  Introduce the class to the concept of empty magnification - the idea that you can magnify an image more than you can resolve it and that the extra magnification is essentially useless.  Pointing out that items in the microscope field that are too close to be resolved become distorted, I ask my students what they get if they magnify such a distorted image.  The answer, of course, is that you get a larger distorted image.  I emphasize that our microscopes in the teaching lab magnify 1000-fold at their highest power, while their resolution is only about 500 times better than the naked eye.  I then explain how oil immersion works to boost resolution by effectively increasing N. A.  If oil is placed between the slide and the objective lens as it is in oil immersion, light between the specimen and the objective is not bent away from the lens as much as it is when no oil is present.  The result is more light gathered by the objective lens, a higher N. A. and better resolution.  

Illustration

Numerical Aperture and Resolution

I suggest some experiments that students can do on their own that illustrate the effect of numerical aperture on resolution.  Some rooms have their lights on a dimmer switch.  If the students have a room with such controls, they can try to brush their hair or some other similar activity at different light levels.  If they are like most people, low light levels will make it more difficult to see what they are doing in whatever task they have chosen.  Point out to them that as light levels are raised, the numerical aperture of the lens system in their eyes also increases.  Consequently, more light reaches the eyes and their ability to resolve images also increases.  A particularly good example of this is attempting to read in a darkened room.  If the light level is low enough, a reader can tell that the page contains lines of print, but the fine details on each letter that allow them to be recognized are not resolvable.  Thus, the text can be seen, but not easily read.  As light levels are increased, the fine structural details of the letters that allow their recognition can be resolved and the text can be read accurately.

Demonstration

Immersion Oil

I show my students how immersion oil works by demonstrating that the index of refraction of glass and the oil are nearly the same.  The caps on most bottles of immersion oil contain glass applicator sticks, which, when placed in the oil, become virtually invisible.  It is the refraction of light that makes objects placed in water visible.  Since the glass and oil have the same indices of refraction, there is no bending of light and the applicator stick is not visible unless one looks very closely and carefully.  Another demonstration that I have used often was shown to me by a colleague, Dr. Larry Reinking.  He wrote the letter "A" on the back of the frosted end of a frosted slide and placed it on the overhead projector.  The letter "A" is relatively difficult to see; light does not pass freely through the frosted part of the slide, thus largely obscuring the letter.  The frosting results from an irregular rather than smooth surface on that end of the slide.  If the irregular surface is filled in with immersion oil, the slide acts as if it were not frosted, since the indices of refraction of the oil and glass are the same.  The letter "A" is now clearly visible.  Finally, I tell the students about the plot of the movie The Presidio, which deals with the smuggling into the United States of diamonds secreted in bottles of spring water.  Diamonds have the same index of refraction as water; when submerged in the liquid, they become effectively invisible.

Also, address the issue of the visibility of the specimen and the reason for using stains.  We talk about the different stains for specific macromolecules (Feulgen stain for DNA, acetocarmine and periodic acid-Schiff (PAS) stain for carbohydrates, osmium tetroxide for lipids, methyl green - pyronin for RNA).  I emphasize the importance of counterstains and the use of degradative enzymes, like deoxyribonuclease, as a control for localizing the macromolecules being studied.  I also cover immunocytochemistry and the localization of specific macromolecules, like peptide hormones (glucagon, insulin, somatostatin), using enzymes (e.g., alkaline phosphatase) linked to antibodies.

Once I have adequately presented the principles of microscopy, I emphasize some of the specialized techniques used to learn about specimens: phase contrast microscopy (both traditional and inverted), fluorescence, polarization, confocal laser scanning, video, dark and bright field microscopy.  I describe each technique and the principles involved.  I also emphasize the advantages and disadvantages of each technique and what is gained and lost by altering the usual way of viewing specimens.  There are constant trade-offs.  Once I have described these techniques, I show my students micrographs taken with each method.  I ask if they can identify the method used.  I have gotten pictures of confocal scanning microscopy images from companies that sell such devices; such pictures are found in the available brochures these companies supply to their customers.  Bio-Rad has been particularly generous; they occasionally send me their newest confocal laser scanning brochure (although I believe they may have sold this part of their business).  There is also a microscope on the market that gives a three-dimensional image (the High Definition Real-Time 3D Microscope) made by the Edge Scientific Instrument Corporation (Santa Monica, CA; 310-396-9333).  Edge has literature that shows stereo images taken with this scope.  You can pass these around to the class along with the viewing glasses that allow the three-dimensional image to be seen.  It seems to capture the students' imaginations.

Transmission Electron Microscopy

Going back to the equation for resolution distance, ask your class why the resolution with electron microscopes is so much better than that with light microscopes.  The answer, of course, is that the wavelength of electrons is significantly lower than that for visible light.  The numerical apertures for light and electron microscopes are not all that different.  Consequently, the lower wavelength of electrons translates into the lower resolution distance and better resolution of electron microscopes.  I talk about the structure of electron microscopes and their function and discuss the preparation of samples and their staining.  Ask the students the physical differences between light and electron microscopy stains (colored vs. electron dense) and the functional reasons for the differences.  Describe the different ways in which electron microscope specimens may be prepared.  I ask my students what kind of knife is used to slice sections for electron microscopes.  I inform them, if they do not already know, that the specimens must be embedded in plastic (e.g., Epon), rather than wax, because of the required thinness of the section and the inability of wax to withstand an electron beam.  By asking leading questions, I can usually get them to answer glass or diamond knives.  I ask them the advantages and disadvantages of each.  Glass knives are cheaper, but do not last as long.  Diamond knives are expensive, but more durable.  See if they can figure out why glass knives must be made just before use, without being stored for a significant length of time.  The answer is that glass is fluid.  Once the edge is made, it will only remain sharp enough for a short while.  Eventually, glass in the edge will flow out of the edge, dulling it.

Briefly outline different electron microscope techniques: negative and positive staining, shadow casting, freeze fracture - freeze etch, scanning EM, atomic force or scanning tunneling microscopy, high voltage transmission electron microscopy, etc.  Mention the advantages and disadvantages of each and the kinds of images that are obtained in the corresponding electron micrographs.  Emphasize the differences between transmission and electron microscopes, their structure and function.  Compare and contrast the process of image formation in the two types of microscope.  I recommend that you show the class overheads or slides depicting the various techniques discussed so that the students can get a feel for the different quality of each picture.  This will aid them greatly in their attempts to understand and interpret the electron micrographs that they will encounter in this and other courses.

Another presentation that grabs students' attention is a discussion of the potential dangers of microscopy.  I tell them about mishaps in microscopy labs that I have witnessed or been told about.  Microscopy, on the surface, seems like such a safe pursuit, but I have friends and acquaintances, who have experienced some accidents.  For instance, there is the grad student, who dropped a microtome blade, injuring his shoe but fortunately not his foot, the cuts I experienced courtesy of a glass blade and a lack of manual dexterity in my electron microscopy course, etc.  Draw on similar stories from your own experience.  I also point out that many of the chemicals used in microscopy, especially EM, are toxic, radioactive and/or carcinogenic.

The Use of Radioisotopes

At times, it seems that students can be woefully underinformed about radioisotopes and their use in biology, despite their widespread application.  I recommend that you briefly explain what makes an isotope radioactive - an imbalance between protons and neutrons in the atomic nucleus - and the nature of the radiation emitted by such isotopes - alpha, beta and gamma.  Define the term half-life as well.  Describe the two major methods that biologists employ to quantify and/or locate radioisotopes in a particular specimen: liquid scintillation spectrometry and autoradiography.  Once again, emphasize the advantages and disadvantages of both of these techniques.  

Cell Culture

Briefly describe the process of cell and tissue culture.  Stress its importance in much of the research that has helped us to comprehend cellular physiology and behavior.  Outline the factors that must be addressed in order to make data obtained from cultured cells believable - the prevention of culture contamination, the maintenance of healthy cells by ensuring that appropriate factors are in the medium, maintenance of proper nutrient levels, etc.

The Fractionation of a Cell's Contents by Differential Centrifugation

Outline the philosophy behind differential centrifugation and how it can be used to isolate a particular cellular organelle.  I usually take my class through a procedure like that described in the Karp text book.  The transparency of the corresponding figure in Karp's text is useful for the class to follow.  Ask the students why an isotonic medium is employed and why the temperature is kept low.  Make sure they understand the significance of using longer spin times and/or higher velocities to isolate increasingly smaller organelles and structures. Mention the different types of centrifuge rotors - fixed angle and swinging bucket - and ask the students where the pellets will typically be found.  Tubes from swinging bucket rotors, of course, have a pellet formed on the bottom of the tube, while the pellet in tubes from a fixed angle rotor is found farther up the side of the tube.  Also distinguish between straight centrifugation and the use of a sucrose density gradient.  Describe the use of a gradient for separating organelles that sediment together during centrifugation without a gradient because of their similarity in size, despite differences in their densities, and how gradients can effect separation on the basis of density differences.  The figure in Karp (Figure 18.23) is excellent for this purpose and I recommend using it as a transparency, slide or part of a PowerPoint presentation in class.  

Isolation, Purification and Fractionation of Proteins

I emphasize in these lectures techniques that are used predominantly to isolate and purify proteins and generally leave the DNA techniques to be covered in the Genetics and Molecular Biology courses.  I do not want to steal the thunder of the professors in those courses.  

Differentiate between preparative and analytical techniques, but tell your students about the overlap between the two.  Emphasize that certain techniques work better if used early in a sequence of procedures, while others are more effective after the protein has already been purified to some degree.  Emphasize that the goal of all purification techniques is to retain the protein of interest while getting rid of the other proteins in the preparation.  Explain that a technique that decreases the amount of the protein to be purified significantly can be eliminated.  One that maintains levels of the protein to be isolated, while decreasing contaminant levels, is useful.  To make these kinds of judgments, one must have a way of measuring the amount of the desired protein present (usually by a specific assay of its enzyme activity or binding ability) as well as the amount of total protein present (Lowry or Biuret protein assays).  Knowing these values at each step allows the calculation of specific activity (enzyme or binding activity/total amount of protein).  With each purification step, the specific activity will get larger as long as some degree of purification is effected.  It will decrease if a particular step has been ineffective.  This affords an investigator the ability to assess at each step the effectiveness of his/her procedures.  If specific activity after a certain step is divided by the specific activity prior to its application, the number obtained describes the amount of purification obtained during that purification step (e.g., 10-fold or 23-fold purification).  This can also be used to estimate the overall purification after the application of a number of steps.  It would not be unusual nowadays to be able to purify a protein to homogeneity (in excess of 250,000-fold).  Once you have presented this concept, proceed to a description of various techniques.

Selective Precipitation

Ask students how proteins stay in solution.  Use leading questions if they are needed.  Of course, interactions of R groups with water are responsible for protein solubility.  Added saturated salt solutions compete with R groups for interactions with water.  Low salt concentrations initially salt proteins into solution, actually enhancing solubility.  Then proteins begin to drop out of solution as their interactions with water are stolen away by the salt, forcing the proteins to interact with each other.  More soluble proteins will generally stay in solution longer, but they too will eventually precipitate.  Point out the ways in which this technique may be used for preliminary purification.  You may wish to pose a hypothetical experiment illustrating this technique to see if your class understands.  I have used this approach and it is effective.  

The protein of interest on this chart obviously precipitates at 40% saturated ammonium sulfate.  Good purification can be achieved by removing the 30% pellet, raising the remaining supernatant to 40% saturation and then isolating the 40% pellet.  That way you eliminate proteins that precipitated out of solution at 30% saturation with ammonium sulfate and below and those that would precipitate out above 40% saturation.  With this procedure, you have isolated those proteins that precipitate out between 30 and 40% saturation with ammonium sulfate.

Chromatography

Describe the common features of all chromatography techniques: the mobile and immobile phases, for example.  Then, describe a number of different chromatography techniques, the principles that make them work, and the molecular traits by which they achieve their separations.  I usually cover gel filtration, ion-exchange chromatography and affinity chromatography, making clear that there are other techniques.  I also show students what the data output from each technique looks like and how to interpret the data.

In gel filtration, the data obtained are really in terms of Stokes' radius (the hydrodynamic radius), the longest radius of the molecule.  A spherical molecule has essentially one Stokes' radius.  A cigar-shaped molecule has a number of radii (essentially infinite).  The longest of these determines which pores the proteins can penetrate.  Thus, cigar-shaped molecules actually travel through gel filtration columns as if they are bigger than they really are because their movement is determined by their longest radius.  An estimate of their molecular weight would be too high, as demonstrated in the drawing above.  Therefore, the molecular weights of cigar-shaped molecules cannot be determined directly with gel filtration.  However, the molecular weights of globular (spherical) proteins can be accurately determined with gel filtration when compared against globular standards of known molecular weight.

Analogy

The Window Screen Analogy

I tell the class that gel filtration media are composed of inert cross-linked polymers (polyacrylamide or a polysaccharide) and that the amount of cross-linking can be controlled.  I tell the class to imagine a stack of window screen cut into a sphere (a very small sphere).  Furthermore, I tell them that this is unusual window screen in that the openings in the screen are not uniform.  Some of the openings are very big, some are middle-sized and some are small.  The average size of the openings can be controlled by the number of cross-links.  A larger number of cross-links results in a smaller average pore size throughout the sphere of window screening.  A smaller number of cross-links results in a larger average pore size.  I propose to the students that there are in a mixture of proteins a large molecule, a small molecule and a middle-sized molecule.  I then ask the class what would happen if each of these proteins was presented to a single bead of the medium.  They usually realize that the big molecule will move past the bead without penetrating it, the middle-sized protein will penetrate part of the way into the bead and the small molecule will penetrate the bead completely.  The degree to which a protein is able to penetrate a bead is directly related to the extent of movement retardation through the column that it experiences.  I then ask them to imagine what would happen, if the effect just described were to be multiplied over a column filled with millions of these tiny beads.  Students generally pick up that the larger protein will exit the column first, while the smallest will exit last.  The middle-sized protein will elute between the other two.  

Ion exchange chromatography seems to be the type of chromatography that most students seem to be able to grasp most easily.  If they've picked up anything by the time they get through the biochemistry section of my course, it's that positive charges attract negative charges and vice versa and that like charges repel.  I explain that two of the most prominent ion exchange media are Diethylaminoethyl (DEAE) - cellulose and Carboxymethyl (CM) - cellulose.  I ask them what they think the charges are on the two types of media.  Usually, they make the connection between amino groups and positive charges and carboxyl groups and negative charges.  When I ask them the overall charge of molecules that stick to DEAE-cellulose, they seem to figure it out.  The same goes for CM-cellulose.  

I then ask them how they would elute these molecules from the column.  Sometimes they get it right away, sometimes some leading questions are required.  The passage of salt solutions (NaCl, KCl) of increasing ionic strength through the column releases proteins in an essentially sequential manner.  Those proteins that adhere to the column less tightly (lower opposing charge) are eluted at lower salt concentrations, while proteins that are more tightly adherent (higher opposing charge)  elute at higher salt concentrations.  I point out that this means that the salt concentration at which a protein washes off the column is at least a rough indicator of its overall molecular charge.  I ask them to explain to me how, for example, KCl elutes proteins from a DEAE-cellulose column.  Often, somebody picks up that the K+ ions compete with the positively charged DEAE column for the negative charges on the protein, while the Cl- ions compete with the negatively charged R groups of the protein for interaction with the column's DEAE groups.  I then ask them to apply the same logic to CM-cellulose.  

Finally, I show them schematic drawings of the data collected in such an experiment or sometimes actual results from one of my older publications and demonstrate how information can be extracted from the column profiles.  To see if they've been following along, I ask them what they can tell me about any likely charge differences between molecules appearing at different parts of the profile, e.g., which of the two peaks on the profile represents the more positive (or negative) protein?  I try to bring in a modern fraction collector, so they can get an idea of how the samples are collected and tell them stories about my primitive fraction collector in graduate school that required me to come to the lab at 3 AM to make sure that the drops were not missing the collecting vials as they exited the column.  I also point out the sad truth that if any drops missed the vials, they were invariably the drops that contained the protein I was studying.  I have, on occasion, placed a bonus question on my exams containing a graph of an ion exchange elution profile with the salt gradient shown.  I ask questions like: At what KCl concentration does peak II elute?  Which protein is likely to have a lower negative charge - the one in peak I or peak II and why do you think so?

Affinity chromatography sometimes seems to give the students trouble.  Maybe it's their ability to comprehend the meaning of affinity or that it's too much chemistry.  I use the same example Karp does, isolation of the insulin receptor by attaching insulin to the inert matrix of the affinity column.  We also isolate horse serum albumin in lab with Affi-gel Blue (Bio-Rad) and then run an SDS-PAGE gel to demonstrate both techniques.  I ask the students how they would elute the bound receptor off of the column after it had bound.  I usually get the right answers - salt again, pH gradient and the passage of insulin through the column.  I point out that, if possible, you want to use the affinity column again since many of the media are expensive and/or time-consuming to produce.  Thus, it is a good idea to use a relatively gentle procedure for eluting your protein.  I also emphasize the high degree of purification that can be attained in this one step (2,000 - 3,000 fold; one or, at most, a few proteins separated from hundreds or more) and that affinity chromatography is usually employed after at least some preliminary purification has occurred.  I also mention that this technique is used successfully to purify mRNAs from total cellular RNA.  After reminding them that most mRNAs contain a poly(A) tail, the students can usually figure out that attaching oligo (dT) to the column matrix will do the trick. Sometimes they guess that oligo (U) might work.

Polyacrylamide Gel Electrophoresis

I describe the principles involved in separation of molecules by electrophoresis and the differences in isolation by nondenaturing PAGE, SDS-PAGE, isoelectric focusing and 2-D gel electrophoresis.  I also describe what the gels look like and what type of information can be extracted from them.  I extend the gel filtration analogy to cover gel electrophoresis.

Make sure that the students understand which properties each electrophoresis technique employs to effect a separation.  To do this effectively, it is important that the students understand the concept of charge density and its relationship to the movement of proteins through the gel.  Also, clarify the effect of treating proteins with SDS, including its influence on the charge density of the protein molecules.  It is also useful to outline the purpose of the molecules found in the sample buffer: glycerol or sucrose, a tracking dye, sodium dodecyl sulfate, mercaptoethanol, etc.  Ask them leading questions to direct them to the correct understanding of all of these concepts.  

Analogy

The One-Big-Bead Analogy

For all intents and purposes, an electrophoresis gel, whether in a tube or a slab, is very much like one giant bead of a gel filtration column.  Electrophoresis gels and chromatography beads are made of virtually identical materials as well.  When proteins are applied to the top of the gel and current applied, most of the molecules in the sample are attracted to the positive pole, but their movement is impeded.  Smaller molecules can travel farther into this giant bead than can larger molecules.  Consequently, despite the fact that what is happening can be explained in terms of the description of gel filtration, the results are just the opposite.  Smaller molecules move faster through the electrophoresis gel than the larger proteins, while, in gel filtration, the smaller molecules move through the column more slowly.  Ask the students if they can figure out why this is not a contradiction.  It is clear that on exams they get the two confused, so you may want to mention it a couple of times.

Determination of Protein Structure by X-ray Diffraction Analysis

I cover X-Ray diffraction only briefly, but I always show the class the Rosalind Franklin X-ray diffraction picture used by Watson and Crick when they figured out the double helical structure of DNA.  I also make sure that the students understand that this technique is useful for determining the fine structure of proteins.  I point out that this technique was used to demonstrate that enzymes and other proteins change shape when they bind to substrates and other ligands.

Purification and Fractionation of Nucleic Acids

I rarely have the opportunity to cover the purification and fractionation of nucleic acids.  That is another topic I leave for the Genetics and Molecular Biology courses.   I do, however, mention that many of the techniques that have been described for use with proteins can be adapted for use with nucleic acids.

Measurements of Protein and Nucleic Acid Concentration by Spectrophotometry

Students in my courses get extensive experience with the use of spectrophotometers and the theory and practice of spectrophotometry.  The first time they are to use spectrophotometry in lab, I go over the Beer-Lambert Law and explain absorption spectra, transmittance (transmission) spectra and the use of a standard curve.  They then carry out a lab exercise during which they collect data and graph a number of absorption spectra and construct a protein standard curve using the Biuret test.  They also are given some protein samples of unknown concentration for which they must determine the concentration.  In later labs, they make further extensive use of these concepts.

Demonstration

Fun With Colors

A demonstration that is very successful makes use of colored pieces of cellophane (colored theatrical gels used on spotlights are even better).  I set up a slide projector and place in front of it a prism obtained from Carolina Biological until I get the spectrum projected on the wall or a movie screen.  Before I do anything, I ask students why they see an object as having a particular color. They usually come up with the idea that a blue shirt is blue because the pigments in the shirt predominantly absorb light from the red end of the spectrum, while reflecting mostly the blue wavelengths, which then travel to their eyes (among other places), where they can be detected and perceived.  I point out that translucent materials like the colored gels I am about to use not only reflect certain wavelengths of light but also transmit them.  I then place one of the colored gels into the light path from the projector to the screen.  Nothing demonstrates the concept of absorbance better than this.  If you place a pure red filter in the light path, all colors but red drop out of the spectrum on the movie screen.  A blue gel absorbs red light and transmits blue.  The most interesting colors are magenta/purple gels (transmitting red and blue light), aqua gels (transmitting blue, green and some yellow wavelengths) and especially yellow gels, which transmit yellow light as expected, but also green and red light (usually totally unexpected by the class).  The first two make sense to most people, but yellow gels blow them away until I explain that a mix of equal intensities of red and green light are perceived as yellow.  You can mix colors on the screen using colored slides and three projectors if you want.  You can also bring the pixels in a color TV screen into the story as well.  Tell your students to look at a yellow area (or any other color) on a TV screen or computer monitor and, using a magnifying glass, observe which pixels are lit. If you want to make it more advanced, bring in the red, yellow and blue cones and color vision.  See if students can figure out what is going on with colorblindness.  I have used variations of this demonstration for students from elementary schools up to seniors in college. It is always successful.

Ultracentrifugation

Explain the principles involved in ultracentrifugation, if you have not already done so.  Remind students of those properties of the components being separated (size, shape, and density) and the medium (density and viscosity) that influence the movement of molecules during the centrifugation.  Much of this material can be covered during the discussion of differential centrifugation.  I usually reserve this part of the lecture for a discussion of the power of ultracentrifuges, a discussion that invariably develops a healthy amount of respect for them.  I tell two stories, one out of my experience and one that was told to me by one of my colleagues.  The first I can assure you is not embellished.  It happened as described.  The second may have been altered to some degree before it got to me, but I tell it as it was related to me.

Illustration

Fun With Ultracentrifuges

When I was in graduate school, I was working in a room that housed an ultracentrifuge.  I was involved with an enzyme assay at a work bench across the room from the ultracentrifuge that was running at the time.  My back was to the machine.  A fellow graduate student was running samples in a titanium swinging bucket rotor at about 25,000 rpm.  While pipetting, I suddenly heard behind me what sounded like a loud popping noise, followed by a sound like one of the ghosts dragging chains in A Christmas Carol.  My reaction time was still good enough in the mid-1970s that when I reflexively turned toward the noise, I saw the machine slam back to the floor.  It had been lifted off the floor by the event.  The machine's rotor, of course, immediately began to slow down.  After a second or two, I realized what had happened and went to get the other graduate student.  After we were sure the machine had come to a complete stop, we opened the top.  A plume of white smoke wafted out of the chamber.  We thought a new Pope had been elected.  The swinging buckets were stuck in the up position that they had been in when the incident happened.  Each of the four titanium buckets was split down the middle by the force of what had happened.  The bowl of the centrifuge was also pitted as pieces of metal that had broken free at high speed had acted like shrapnel.  The drive stem was bent at about a 45° angle.  The rotor had derated itself in the middle of the student's centrifuge run.  Her samples, the product of three weeks' work, had evaporated, the tubes in which they had been contained reduced to shards and powder.  The second story concerns essentially the same type of event but in a somewhat older machine with a bit less armor plating.  I was told that the rotor again derated itself but that this time it was a fixed angle rotor traveling at a somewhat higher speed.  The rotor came out of the side of the machine, partially disrobing a student in the room at the time when it grazed her somewhat baggy blouse (she was not hurt).  It then slammed into a cabinet and into a wall leaving a significant impression on both.  The hole in the side of the ultracentrifuge and the wooden door of the cabinet resembled Roadrunner cartoons in which Wile E. Coyote went through a wall leaving a clear outline of his body behind.  There was a hole in both places that clearly resembled a fixed angle rotor.  Students are amused and impressed by these stories and they manage to comprehend the power of these machines.  I know this because the next time they are in lab when the floor model centrifuge has been turned on, the students nearest the machine keep looking over their shoulder.

Nucleic Acid Hybridization, Recombinant DNA Technology, etc.

You may wish to summarize to varying degrees the techniques that relate to DNA.  I generally do not cover these techniques in great detail.  As mentioned earlier, this is by agreement with my colleagues who teach Genetics and Molecular Biology.  There are occasions where I mention these techniques without telling how they work.  I simply tell the students what they accomplish and how they relate to the topics we are discussing at the moment.

The Use of Antibodies

I am surprised at the number of sophomore to junior biology students who do not know what antibodies are and what they do.  Interestingly, the students have heard the words "antibody" and "antigen" and may even have an idea that the molecules these words describe are involved in fighting off infections, but they do not have more detailed knowledge of what they do.  They also often get "antibodies" and "antigens" confused..  Give them some working definitions that explain the roles of antibodies and antigens in at least a rudimentary fashion so that the students can understand that antibodies will seek out and bind specifically to antigens.  Their use in localizing specific molecules in the cell via fluorescence microscopy or radiolabeling of antibodies can then be described.  If you have discussed this previously under light microscopy techniques, of course, you need not do it again.

CRITICAL THINKING QUESTIONS

1.  Which of the following microscopes would exhibit the worst resolution?

a.  a microscope with an N. A. of 1.1 and using blue illumination

b.  a microscope with an N. A. of 1.1 and using orange illumination

c.  a microscope with an N. A. of 0.9 and using red illumination

d.  a microscope with an N. A. of 0.9 and using blue illumination

e.  a microscope with an N. A. of 1.1 and using red illumination

2.  Under which set of parameters above in question #1 would the best resolution be obtained?  a.

3.  What is the magnification of the image in a microscope using a 20X ocular and a 100X objective?  2,000X.

4.  Which is more important to microscopy and why?

a.  magnification, because it makes things easier to see

b.  magnification, because it allows you to distinguish separate objects

c.  resolution, because it allows you to distinguish separate objects

d.  resolution, because it makes things large enough to distinguish

e.  neither one is very important

5.  What is the major disadvantage of using most traditional staining techniques on a microscopy specimen? They kill specimens.

6.  Why is staining so important to light and electron microscopy?  It provides contrast for a specimen that might otherwise lack it, making that specimen easier to see in the light or electron microscope.

7.  What electron or light microscopy technique would you use to observe the following? 

a.  the fine detail of the outer surface of hair cells in the inner ear.  Scanning electron microscopy.

b.  the interior and exterior of surfaces of the plasma membrane to support the Fluid-Mosaic Model of membrane structure.  Freeze - fracture, freeze- etch method.

c.  very thin cross sections made in Epon in order to visualize the fine structure of cellular organelles and cell architecture in the cell interior.  Positive staining, transmission electron microscopy.

d.  particulate matter (like large multisubunit enzyme complexes or viruses) so as to exhibit the subunit structure.  Negative staining.

e.  specifically localize a molecule like tubulin in a cell using a light microscope.  Fluorescence microscopy.

f.  the highly organized structure at the molecular level of the spindle of a dividing cell using the light microscope.   Polarization microscopy.

g.  technique using lasers to optically section large specimens; the images may be computer-enhanced.  Confocal laser scanning microscopy.

h.  living cells without killing them by adding stain.  Phase-contrast microscopy.

i.   a three-dimensional view (almost bas-relief view) of chromosomes in a dividing cell.  Differential interference contrast (DIC) microscopy (Nomarski optics).

j.  living cells with high resolution and with extraordinary contrast at extremely low light levels to keep the cells alive during the observation.  Video microscopy.

k.  viewing poorly stained specimens by focusing light on the specimen alone; the specimen stands out brightly against a darker background.  Dark field microscopy.

8.   Why is resolution in electron microscopes so much better than that in light microscopes?  The wavelengths of electrons are much lower than those of visible light.  Thus, the resolution distance d is thus proportionally smaller.

9.   Why does oil immersion increase resolution so effectively?  Immersion oil has the same RI (refractive index) as the glass of a slide.  Thus, light does not refract away from the objective.  It increases the amount of light a lens can collect.  By increasing the amount of light collected by the lens, it increases the effective N. A.  This decreases d, the minimum resolution distance, and thus improves resolution.

10.  Why are cells homogenized in isotonic media before differential centrifugation?  If cells were homogenized in hypotonic media, osmotically active organelles like mitochondria and chloroplasts would burst.  Isolation by differential centrifugation would then be useless.

11.  What property of a protein can be determined directly with gel filtration chromatography if the protein is spherical?  Stokes' radius (hydrodynamic radius) can be determined directly for any protein with gel filtration.  Molecular weight can be determined directly if the molecule is spherical (globular).

12.  What property of a protein can be determined directly with gel filtration chromatography if the protein is an oblate spheroid (cigar-shaped)?  Stokes' radius (hydrodynamic radius) can be determined directly for any protein with gel filtration.

13.  If a solution containing the following tripeptides were passed through a column packed with DEAE-cellulose at pH 7, which of them would bind most effectively to the DEAE-cellulose column? 

a.  N - arginine - lysine - proline - C                         

b.  N - aspartate - glutamate - aspartate - C 

c.  N - phenylalanine - aspartate - glycine – C

d.  N - leucine - alanine - glycine – C

e.  N - aspartate - phenylalanine - leucine - C

14. If a solution containing the following tripeptides were passed through a column packed with CM-cellulose at pH 7, which of them would bind most effectively to the CM - cellulose column? 

a.  N - arginine - lysine - proline - C                         

b.  N - aspartate - glutamate - aspartate - C               

c.  N - phenylalanine - aspartate - glycine – C

d.  N - leucine - alanine - glycine – C

e.  N - aspartate - phenylalanine - leucine - C

15.  What technique would be used to determine the arrangement of atoms within a protein and has been used to detect and describe the arrangement in space of the atoms of DNA?  X-ray diffraction or X-ray crystallography.                                                   

16.  A protein you wish to study is located in the mitochondria of rat liver cells and is readily extractable from partially purified mitochondria.  It has a highly positive charge at pH 7.  You propose to use a series of techniques to purify the protein.  In what order would you use the techniques after you had homogenized the tissue?

a.  sucrose density gradient centrifugation (SDGF) - extraction - isoelectric focusing (IEF) - 

precipitation by ammonium sulfate

b.  differential centrifugation - SDGF - extraction - ammonium sulfate precipitation - CM (carboxymethyl)- cellulose chromatography - affinity chromatography

c.  ammonium sulfate precipitation - SDGF - extraction - CM cellulose chromatography - IEF - SDGF

d.  IEF - affinity chromatography - differential centrifugation - extraction

e.  differential centrifugation- SDGF - extraction - ammonium sulfate precipitation - DEAE 

(diethylaminoethyl) cellulose chromatography - affinity chromatography

17.  Which one of the following molecules will elute first from a gel filtration column?  (Assume that all amino acids have the same molecular weight.)

a.  a globular protein composed of 325 amino acids       

b.  a chromate ion                                                            

c.  a cigar-shaped protein composed of 326 amino acids

d.  a globular protein composed 147 amino acids

e.  a protein consisting of 200 amino acids

18.  You wish to carry out a preliminary purification of proteins that is rapid and based on their solubility.  What is the name of the technique you would be most likely to use?  Selective (ammonium sulfate) precipitation.

19.  What technique is best for accurately determining the amount of radioactive isotope in a cell fraction?

a.  scanning EM 

b.  scanning tunneling EM              

c.  liquid scintillation spectrometry 

d.  autoradiography 

e.  ultracentrifugation

20.  What technique is best for determining the location of radioactive isotope in a tissue or cell?

a.  scanning EM 

b.  scanning tunneling EM 

c.  liquid scintillation spectrometry

d.  autoradiography

e.  ultracentrifugation

21.  What techniques are most often involved in two-dimensional gel electrophoresis?  

a.  none of the other answers                  

b.  gel filtration, SDS-PAGE 

c.  isoelectric focusing, centrifugation

d.  isoelectric focusing, non-denaturing gel electrophoresis

e.  isoelectric focusing, SDS-PAGE

22.  Which technique below would be likely to result in the highest degree of purification of a preparation of a hormone receptor?

a.  ion exchange chromatography

b.  ammonium sulfate precipitation           

c.  gel filtration

d.  affinity chromatography

e.  homogenization

23.  What advantage does SDS-PAGE have over nondenaturing PAGE?  SDS-PAGE allows the accurate and direct determination of a polypeptide's molecular weight.  What advantage does nondenaturing PAGE have over SDS-PAGE?  Nondenaturing PAGE does not allow the direct determination of molecular weight, but does allow a protein to maintain its shape and thus its activity, thus allowing it to be easily located in the gel.  These properties are usually lost in SDS-PAGE.

24.  A protein sample is placed on a gel that contains a gradient of H+ ion concentration.  Current is applied and the individual proteins travel a certain distance into the gel until they stop.  Which protein purification technique was being used?  Isoelectric focusing.  Why do the proteins stop moving?  As they travel through the H+ ion gradient, the overall charges on the proteins change, becoming progressively more positive.  Eventually, all of the proteins will reach a pH at which their overall charge is neutral.  When their charges become neutral, the proteins are attracted to neither of the electrodes.  As a result, they stop moving and become concentrated in very tight, thin bands.

25.   You treat a chicken cell with antibodies against chicken membrane proteins and a mouse cell with antibodies against mouse membrane proteins.  The anti-chicken antibodies are labeled with fluorescein (green) and the anti-mouse antibodies are labeled with rhodamine (red).  You fuse labeled cells from the two different species (chicken and mouse) together.  Right after fusion is complete you place the resultant heterokaryon in a special microscope and shine UV-light on it.  One half of the cell glows red, the other half green.  What technique does this describe?  Fluorescence microscopy.

26.  Identify the techniques described below. 

a.  protein separation based on molecular charge and molecular weight   Nondenaturing PAGE.

b.  separation of proteins on the basis of molecular weight alone   SDS-PAGE.

c.  localization of radioactive molecules taken into a cell preparation   Autoradiography.

27.  Which technique effectively separates proteins when they travel through a charged medium, to which some of the proteins adhere, while others do not adhere and pass through the column?  Ion-exchange chromatography.

28.  You are purifying an enzyme.  Initially, before purification, the protein's specific activity is 25 units/mg protein.  After 3 successive purification procedures, the specific activity is 800,000 units/mg protein.  How much have you purified the protein?  32,000 times.

29.  You are using a variety of purification techniques to purify a protein and monitoring the specific activity after each step.  What circumstance would lead you to drop a particular step in the purification scheme?  You should drop the step if it results in a decrease in specific activity.

30.  You treat a nucleic acid preparation with phenol and buffered saline.  After a few such treatments, the aqueous phase is incubated with RNase.  What nucleic acid polymer is found in the aqueous phase?  DNA.

31.  DNA is digested with a restriction enzyme and applied to an agarose gel.  Four fragments are seen and they are found to be 10 kb, 8.5 kb, 5 kb and 2.2 kb long.  Which fragment is closest to the top of the gel?  The 10 kb fragment.  Which fragment is closest to the bottom?  The 2.2 kb fragment.

32.  DNA from organism A is hybridized separately to DNA of two other species, C and D.  The A-C hybrids melt at a temperature of 96°C; the A-D hybrids melt at a temperature of 91.5°C.  Which DNA, C or D, is likely to be most closely related to A?   A and C are most closely related, since they pair more stably.  This is demonstrated by the higher temperature required to separate these two chains of DNA after hybridization.

33. Below is a standard curve.   If an unknown protein sample has an optical density of 0.5, what is its concentration?   ~95 mg/ml.

34.  Why do you treat a bacterial plasmid and the eukaryotic DNA you wish to insert within it with the same restriction enzyme?  A particular restriction enzyme cuts DNA only at one specific sequence.  If the result of the cut leaves complementary "sticky ends" (single-stranded overhangs of DNA) on the end of the DNA fragments, the ends of any fragments cut by this enzyme will be able to base pair and be closed up by DNA ligase.  If the two pieces of DNA thus joined are a plasmid and eukaryotic DNA, a recombinant DNA has been constructed.  If you had treated the two DNAs with different restriction enzymes, the sticky ends of each would not match and the two fragments would not join.

35.  Why do you treat DNA to be inserted into YACs with restriction enzymes that have longer recognition sites?  YACs are used for cloning longer DNA fragments.  If the DNA to be cloned is treated with such a restriction enzyme, the probability of finding the restriction site decreases since longer sites are rarer.  This would result in fewer cuts in the DNA to be cloned and, thus, longer DNA fragments, which can be cloned efficiently in YACs.

36.  What is the difference between a cDNA library and genomic library?  A genomic library is a collection of all DNA sequences in an organism's genome, including both expressed and nonexpressed genes.  A cDNA library contains only those sequences transcribed in an organism, tissue, or cell at a particular time.  Nonexpressed sequences are not included.

37.  You have managed to incorporate stably into the genome of a mouse a gene for the growth hormone of a guinea pig.  What is such an animal called?  A transgenic animal.

38.  Two mice received the same mutant form of a gene when embryonic stem cells containing the mutant gene were injected into their early embryos.  When these two phenotypically normal mice are mated, some of their offspring show severe developmental anomalies.  How does this happen?

The mutant form of the gene has gotten into the gametes of both mice.  When gametes carrying the mutant gene fuse, they are homozygous for the mutant allele.  It is likely that the gene thus "knocked out" is important for normal development.  Thus, about one fourth of the offspring of these two mice should exhibit the anomalies.

39.  Why is the DNA polymerase used in PCR best if it comes from bacteria that live in hot springs?

Bacteria that live in hot springs are constantly exposed to temperatures significantly higher than most bacteria.  In order to survive, they have evolved a DNA polymerase that is much more stable at higher temperatures than that of bacteria that normally live under more temperate conditions.  Such a DNA polymerase can withstand, for longer periods of time, the high temperatures needed to melt DNA duplexes during PCR.

40.  Why must you fuse a myeloma cell to a lymphocyte in order to make monoclonal antibodies?  Normal lymphocytes are difficult to culture and proliferate; malignant myeloma cells, on the other hand, can proliferate quite well in culture.  By fusing a myeloma cell with a lymphocyte, you obtain a cell that can be used to create a clone of cells that all make one antibody molecule (the one made by the lymphocyte that originally fused with the myeloma cell).  The fused cell retains the ability of the myeloma cell to proliferate and the ability of the lymphocyte to make large amounts of a single antibody.  Thus, large amounts of a single (monoclonal) antibody can be produced.

41.  You have made an antibody to human actin by injecting a rabbit with appropriate doses of purified human actin.  If you wish to use indirect immunofluorescence to localize human actin in a cell, what would your next step be?  You would inject another animal, like a goat, with rabbit antibodies, allowing the goat to make antibodies against rabbit antibodies (goat anti-rabbit antibodies).  You would then label these secondary antibodies with a fluorescent molecule like fluorescein or rhodamine.  After exposing the human tissue in which you wish to visualize actin to rabbit anti-human actin antibody, you would expose the tissue to the labeled goat anti-rabbit secondary antibody.  The binding of the secondary antibody would boost the fluorescence signal and make it easier to localize the human actin.  The signal is boosted because more secondary antibodies can bind to the primary antibodies than the number of antibodies that can bind to actin in the same region

ART QUESTIONS

1.  In Figure 18.1, which lens is closest to the specimen and which is closest to the viewer's eye?  The objective lens is closest to the specimen, the ocular lens closest to the viewer's eye.

2.  According to Figure 18.2, which image is focused closest to the objective lens: that of the specimen or that of the light source?  That of the light source.

3.  How can you tell that the cell shown in Figure 18.5 is in metaphase?  The chromosomes are lined up in the center of the cell.

4.  Which of the microscopy techniques shown in Figure 18.6 gives a three-dimensional quality to the image?  Differential interference contrast (DIC) or Nomarski optics.

5.  The lens systems of light and electron microscopes are shown in Figure 18.12.  Of what are the lenses of an electron microscope composed?  Electromagnets.

6.  According to Figure 18.13, what is the first fixative used in the procedure used to prepare specimens for electron microscopy?  Glutaraldehyde.  What is the second fixative used in the procedure?  Osmium tetroxide.  What is the name of the device that is used to slice sections for the electron microscope and usually how thick are the specimens?  The sections are cut on an ultramicrotome and they are usually approximately 100 nm thick.  On what are the sections floated after they have been cut and before they are picked up with the grid?   Water.

7.  What is the most obvious difference between the two images of the tobacco rattle virus in Figure 18.14?  The negatively stained image appears flat, while the shadow cast image has a 3-D appearance.

8.  According to Figure 18.15, what modification to the shadow casting procedure facilitates the visualization of DNA and RNA molecules?  The modification is called rotary shadowing; the metal is evaporated at a very low angle, while the specimen is rotated.

9.  Look at the schematic drawing of the replica making process in Figure 18.16.  From which direction did the heavy metal come in forming the replica?  The upper left.

10.  In Figure 18.21a, 3H-uridine was used to label RNA.  Why was radiolabeled uridine used to label RNA instead of another nucleotide?  Since uridine is found only in RNA, labeled uridine will be incorporated only into RNA.  If labeled adenosine, cytosine or guanidine is used, they might be incorporated into both DNA and RNA, muddying the results.  If thymidine were radiolabeled, only DNA would be labeled and RNA would not be labeled at all.

11.  Assuming that the cells shown in Figure 18.22a are not cancer cells, they will form a single layer of cells on the culture dish.  What will this be called?  A monolayer.  A monolayer forms because cells surrounded on all sides tend to stop growing.  What is this phenomenon called?  Contact inhibition.

12.  According to Figures 18.23, what force is required to sediment mitochondria through a sucrose density gradient and separate them from lysosomes and peroxisomes and for how long must it be applied?  65,000 x g for 2 hours.  Which are the densest organelles - lysosomes or peroxisomes and how much denser are they?  Peroxisomes, with a density of 1.23 grams/ml, 0.05 grams/ml denser than mitochondria and 0.11 grams/ml denser than lysosomes. 

13.  If you were conducting an experiment like the one in Figure 18.24 and you wanted to isolate a basic protein, what ion exchange medium would you use?  Since basic proteins have a predominance of positively charged R groups, you would probably use carboxymethyl cellulose.

14.  You are separating three globular molecules by gel filtration of the following molecular weights: 125,000 daltons, 56,000 daltons and 12,500 daltons in a manner similar to that shown in Figure 18.25.  Which one would elute from the gel filtration column first?  The 125,000 dalton protein.

15.  In Figure 18.27, how would the binding of a "bait" protein to a "fish" protein be detected?  If the "fish" and "bait" proteins bind to each other, they will bring together the two parts of the transcription factor controlling the synthesis of a reporter gene, in this case, the lacZ gene, encoding -galactosidase.  If -galactosidase is synthesized, its presence can be easily detected by changes in the appearance of the yeast cells on the culture dish, specifically their color.

16.  In polyacrylamide gel electrophoresis, like that shown in Figure 18.28, what is the charge on the proteins migrating through the gel?  Negative.

17.  Are most of the proteins shown on the two-dimensional electrophoresis gel in Figure 18.29 acidic or basic proteins?  Since most of the proteins have isoelectric points below 7.0, most of the proteins on the 2-D gel are acidic proteins.

18.  Figure 18.34a shows the DNA moving toward the positive electrode.  Why does this happen and what part of the molecule is responsible?  DNA has a negative charge and is thus attracted to the positive electrode.  The negative charge is imparted to DNA by the phosphate groups of its backbone.  What stain was used to stain the DNA in Figure 18.34b?  Ethidium bromide.  What color is the stained DNA?  Fluorescent orange.

19.  Treatment with a restriction endonuclease cuts DNA into many different fragments of different sizes.  If you stained a gel on which you had separated the restriction endonuclease-treated DNA as in Figure 18.34, what would you see?  There would likely be no visible bands.  The DNA would be visible as a smear of DNAs of many different sizes.  How would you distinguish a piece of DNA containing a specific DNA sequence from among the wide variety of DNA fragments?  You would blot the gel onto a piece of nitrocellulose paper and expose the blot to a radiolabeled DNA probe containing a DNA sequence complementary to the desired DNA sequence.  After autoradiography, the position of the specifically bound radiolabeled DNA probe can be seen, usually as a narrow band.

20.  In Figure 18.35b, why is DNA quantified at 260 nm?  Nucleic acids absorb UV light maximally at 260 nm.  Therefore, that is the best wavelength at which to measure absorbance.  Why is G-C rich DNA denser than A-T rich DNA as depicted in Figure 18.35?  G-C base pairs possess 3 H bonds (while A-T base pairs have only 2 H bonds) among other differences and are denser than A-T base pairs.

21.  According to Figure 18.39, what is used to join covalently the sticky ends of DNA fragments to the free sticky ends of a bacterial plasmid?  DNA ligase.

22.  According to Figure 18.40, what is the reason for lysing bacterial cells and denaturing their DNA  after making a replica of the culture dish on a nitrocellulose filter.  Denaturing double-stranded DNA converts it to single strands, which adhere to the filter.

23.  Look at the outline of the procedure for Polymerase Chain Reaction in Figure 18.43.  Why are primers needed for this procedure?  The enzyme used in the procedure is a DNA polymerase.  All DNA polymerases require a primer to carry out DNA synthesis.  Why is the temperature of the reaction mixture lowered after the newly synthesized double-stranded DNAs from the previous step have been separated?  The cooling of the reaction mixture allows primers to hybridize to complementary sequences in the target DNA.  Why must the DNA polymerase be heat-resistant?  It must be heat-resistant to withstand the high temperatures required for DNA denaturation.

24.  In Figure 18.44a, every DNA fragment in the third lane of the gel from the left has what nucleotide on its 3' end?  A dideoxynucleotide containing a cytosine base.

25.  In the schematic drawing in Figure 18.45, what enzyme is used to synthesize the second strand of a cDNA molecule?  DNA polymerase I.  How is the nick made in the RNA chain of the RNA-DNA hybrid?  RNase H is used to make the nick in the RNA chain of the RNA-DNA hybrid.  After the nick is made in the RNA chain, how is the rest of the RNA digested?  The nick allows the DNA polymerase I enzyme to attach to the RNA at the free 3' hydroxyl at the nick.  The 5'—> 3' exonuclease activity of DNA polymerase I digests the RNA and replaces it with DNA as it uses the 3'-hydroxyl group that was exposed at the nick as a primer.  According to the text, what is the name of this process?  Nick translation.

26.  According to the procedure shown in Figure 18.49, how can you tell that you have a chimeric mouse?  Chimeric mice will be easily determined since the host (recipient) part of the embryo carries the black allele for coat color, while the donated cells also containing the mutant gene carry the allele for a different coat color.  If a mouse embryo has successfully picked up the donor cells, it will have a coat with the characteristics of both the donor and recipient strains.

27.  According to Figure 18.51, why do only the fused cells survive?  The mouse myeloma cells are mutants that cannot grow on HAT medium.  If they fuse with lymphocytes from the mouse spleen, which contain the normal form of the enzyme required for growth on HAT medium (hypoxanthine-guanine phosphoribosyl transferase), the fused cells will be able to survive on the HAT medium.  Some of these cells will make the desired antibody in large quantities.