The modern, detailed understanding of cell architecture is based on several types of
microscopy. Because there is no one “correct” view of a cell, it
is essential to understand the characteristics of the key cell-viewing techniques,
the types of images they produce, and their limitations.
Figure 5-1
.
Views of the epithelial cells lining the small intestine, produced by
three different microscopic techniques
(a) Scanning electron micrograph of the intestinal wall. The lumen, or
cavity, of the intestine, is lined by a sheet of epithelial cells that
rests on a fiber-filled material called the basal
lamina. Abundant fingerlike microvilli extend from the
lumen-facing surface of each cell. The three-dimensional appearance of
the cell surface is characteristic of images obtained by this technique.
(b) Transmission electron micrograph through two intestinal epithelial
cells. Clearly visible are the microvilli, often called the
brush border, two nuclei (N), and other organelles.
Parts of the basal lamina and a capillary (a type of small blood vessel)
that courses through the basal lamina are visible at the bottom.
Nutrients absorbed by the cells from the lumen find their way into
adjacent capillaries, which also provide hormonal signals to the cells.
(c) Stained section of the rat intestinal wall viewed in a fluorescence
microscope. The tissue section was stained with Evans blue, which
generates a nonspecific red fluorescence, and with a
yellow-green – fluorescing antibody
specific for GLUT2, a glucose transport protein. This technique
localizes GLUT2 to the basal and lateral sides of the intestinal cells
and shows that it is absent from the brush border. Capillaries run
through the lamina propria, a loose connective tissue beneath the
epithelial layer. [Part (a) from R. Kessel and R. Kardon, 1979,
Tissues and Organs: A Text-Atlas of Scanning Electron
Microscopy,W. H. Freeman and Company, p. 176; part (b) from
P.A. Cross and K.L. Mercer, 1993 Cell and Tissue Ultrastructure,
A Functional Perspective,W. H. Freeman and Companym, p.
293; part(c) see B. Thorens et al., 1990,Am. J.
Physiol.
259:C279, courtesy of B. Thorens.]
Schleiden and Schwann, using a primitive light microscope, first described individual
cells as the fundamental unit of life, and light microscopy has continued to play a
major role in biological research. The
development of electron microscopes greatly
extended the ability to resolve subcellular particles and has yielded much new
information on the organization of plant and animal tissues. The nature of the
images depends on the type of light or electron microscope employed and on the way
in which the cell or tissue has been prepared. Each technique is designed to
emphasize particular structural features of the cell. shows how a typical cell, the epithelial cell
lining the small intestine, appears when viewed by three different microscopic
techniques.
In this section, we focus on the most common application of light and electron
microscopy — to visualize fixed, killed cells.
Although this approach reveals much information, a critical question about such
results is how true to life is the image of a biological specimen that has been
fixed, stained, and dehydrated before examination? Thus we also consider some of the
refinements that allow microscopy of unaltered or less altered specimens.
Light Microscopy Can Distinguish Objects Separated by 0.2 μm or
More
Figure 5-2
.
The optical pathway in a modern compound optical
microscope
(a) The specimen is usually mounted on a transparent glass slide and
positioned on the movable specimen stage of the microscope. Light
from a bright source is focused by the condenser lenses onto the
specimen. The objective lenses pick up the light transmitted by the
specimen and focus it on the focal plane of the objective lens,
creating a magnified image of the specimen. Usually the image on the
objective focal plane is magnified by the ocular lens, or eyepiece,
which is focused on this objective focal plane; it picks up the
light emanating from the already magnified image of the specimen and
projects it onto the plane of the human eye or onto a piece of
photographic film or a video camera. The lamp field stop and other
apertures restrict the amount of light entering or leaving a lens.
(b) The half-angle, α, of the cone of light entering the
objective lens from the specimen is one parameter that determines
the resolution of a microscope: the larger the value of α,
the finer the resolution the objective lens can provide.
The
compound microscope, the most common microscope in use
today, contains several lenses that magnify the image of a specimen under study
(). The total
magnification is a product of the magnification of the individual lenses: if the
objective lens magnifies 100-fold (a 100X lens, the maximum
usually employed) and the
eyepiece magnifies 10-fold, the final
magnification recorded by the human eye or on film will be 1000-fold.
However, the most important property of any microscope is not its magnification
but its resolving power, or resolution — its ability to
distinguish between two very closely positioned objects. Merely enlarging the
image of a specimen accomplishes nothing if the image is blurry. The resolution
of a microscope lens is numerically equivalent to D, the
minimum distance between two distinguishable objects; the smaller the value of
D, the better the resolution. D depends on
three parameters, all of which must be considered in order to achieve the best
possible resolution: the angular aperture, α, or
half-angle of the cone of light entering the objective lens from the specimen;
the refractive index, N, of the air or fluid medium between the
specimen and the objective lens; and the wavelength,
λ, of incident light:
D = (0.61λ) ÷
(N × sin α).
Decreasing the value of λ or increasing either N or
α will decrease the value of D and thus improve the
resolution. Note that the magnification is not part of this equation.
The angular aperture, α, depends on the width of the objective lens and
its distance from the specimen (). Moving the objective lens closer to the specimen increases the
angle α and thus sin α, and therefore reduces
D (i.e., increases the
resolution). Intuitively, one can
recognize that increasing α allows a greater fraction of the light
emanating from the specimen to enter the objective lens. The refractive index
N is a measure of the degree to which a medium bends a
light ray that passes through it; the refractive index of air is defined as 1.0.
Use of immersion oil, which has a refractive index of 1.5, is a simple way to
reduce
D by 33 percent. An intuitive explanation for this
improvement is that a medium with a higher refractive index than air, if placed
between the specimen and the objective lens, will “bend”
more of the light emanating from the specimen such that it goes into the lens.
Finally, the shorter the wavelength of incident light, the lower will be the
value of
D and the better the
resolution.
Due to limitations on the values of α, λ, and
N, the limit of resolution of a light
microscope using visible light is about 0.2 μm (200 nm). No matter how
many times the image is magnified, the microscope can never resolve objects that
are less than ≈0.2 μm apart or reveal details smaller than
≈0.2 μm in size. This is true because the maximum angular
aperture for the best objective lenses is 70° (sin
70° = 0.94). With the visible light of
shortest wavelength (blue, λ = 450
nm) and with an immersion oil (N
= 1.5) above the sample, then
or about 0.2 μm.
Despite this limit of resolution, the light microscope can be used to track the
location of a small bead of known size to a precision of only a few nanometers!
If we know the precise size and shape of an
object — say, a 5-nm sphere of
gold — and if we use a video camera to record
the microscopic image as a digital image, then a computer can calculate the
position of the center of the object to within a few
nanometers. This technique has been used, to nanometer resolution, for tracking
the movement of gold particles attached via antibodies to specific proteins on
the surface of living cells.
Samples for Light Microscopy Usually Are Fixed, Sectioned, and
Stained
Figure 5-3
.
Preparation of tissues for light microscopy
A piece of fixed tissue is dehydrated by soaking it in alcohol-water
solutions, then in pure alcohol, and finally in a solvent such as
xylene. The specimen is next placed in warm liquid paraffin, which
is allowed to harden. A piece of the specimen is mounted on the arm
of a microtome. The arm moves up and down over a metal or glass
blade, cutting specimen sections a few micrometers (microns)
thick.
Specimens for light microscopy are commonly fixed with a solution containing
alcohol or formaldehyde, compounds that denature most
proteins and nucleic
acids. Formaldehyde also cross-links amino groups on adjacent molecules; these
covalent bonds stabilize
protein-
protein and
protein –
nucleic acid interactions and render
the molecules insoluble and stable for subsequent procedures. Usually the sample
is then embedded in paraffin or plastic and cut into thin sections of one or a
few micrometers thick ().
Alternatively, the sample can be frozen without prior fixation and then
sectioned; this avoids the
denaturation of
enzymes by fixatives such as
formaldehyde.
Since the resolution of the light microscope is ≈0.2 μm and
mitochondria and chloroplasts are ≈1 μm long (about the size
of bacteria), theoretically one should be able to see these organelles. However,
most cellular constituents are not colored and absorb about the same degree of
visible light, so that they are hard to distinguish under a light microscope
unless the specimen is stained. Thus the final step in preparing a specimen for
light microscopy is to stain it, in order to visualize the main structural
features of the cell or tissue. Many chemical stains bind to molecules that have
specific features. For example, hematoxylin binds to basic
amino acids (lysine and arginine) on many different kinds of proteins, whereas
eosin binds to acidic molecules (such as DNA, and aspartate
and glutamate side chains). Because of their different binding properties, these
dyes stain various cell types sufficiently differently that they are
distinguishable visually. Two other common dyes are benzidine,
which binds to heme-containing proteins and nucleic acids, and
fuchsin, which binds to DNA and is used in Fuelgen
staining.
Figure 5-4
.
Cytochemical staining
Light micrograph of a cross section of human skeletal muscle stained
for succinate dehydrogenase, an enzyme found only in mitochondria.
At this low magnification the stained mitochondria appear as purple
dots; in skeletal muscle there are several different types of cells
differing in the number of mitochondria. [From P. R. Wheater, H. G.
Burkitt, and V. C. Daniels, 1987, Functional Histology; A
Text and Colour Atlas,2d ed., Churchill Livingstone,
Flg. 1.23b, p.25. Photo Reserchers, Inc.]
If an
enzyme catalyzes a reaction that produces a colored or otherwise visible
precipitate from a colorless precursor, the
enzyme may be detected in cell
sections by their colored reaction products. This technique is called
cytochemical staining ().
Fluorescence Microscopy Can Localize and Quantify Specific Molecules in
Cells
Figure 5-5
.
The optical pathway in an epi-fluorescence microscope
Light from a multiwavelength source moves through an excitation
filter, which allows only the desired wavelength of exciting
radiation to pass. This radiation is reflected downward by the
dichroic mirror and focused by the objective lens onto the sample.
Fluorescent molecules in the sample are then excited to emit light
(fluoresce) at a specific and longer wavelength. This emitted light
is focused by the objective lens; most of it passes upward through
the dichroic mirror and is not reflected. A final barrier filter
blocks any residual light of wavelengths not corresponding to that
of the fluorescent substance used to stain the specimen.
Perhaps the most versatile and powerful technique for localizing
proteins within
a cell by light microscopy is
fluorescent
staining of cells and observation in the
fluorescence
microscope. A chemical is said to be
fluorescent
if it absorbs light at one wavelength (the
excitation
wavelength) and emits light (fluoresces) at a specific and longer
wavelength. Most fluorescent dyes emit visible light, but some (such as Cy5 and
Cy7) emit infrared light. In modern fluorescence microscopes, only fluorescent
light emitted by the sample is used to form an image; light of the exciting
wavelength induces the fluorescence but is then not allowed to pass the filters
placed between the objective lens and the eye or camera ().
Revealing Specific Proteins in Fixed Cells
Figure 5-6
.
Fluorescence micrograph showing the distribution of long
actin fibers in a cultured fibroblast cell
A fixed human skin fibroblast was permeabilized with a detergent
and stained with a fluorescent anti-actin antibody before
viewing. [Courtesy of E. Lazarides.]
Four very useful dyes for
fluorescent staining are rhodamine and Texas red,
which emit red light; Cy3, which emits orange light; and
fluorescein, which emits green light. These dyes have a
low, nonspecific affinity for biological molecules, but they can be
chemically coupled to purified antibodies specific for almost any desired
macromolecule. When a fluorescent
dye –
antibody complex is added to a
permeabilized cell or tissue section, the complex will bind to the
corresponding
antigens, which then light up when illuminated by the exciting
wavelength, a technique called
immunofluorescence
microscopy (). By staining a specimen with two or three dyes that fluoresce at
different wavelengths, multiple
proteins can be localized within a cell, as
illustrated in the chapter opening figure and in .
Revealing Specific Proteins in Living Cells
Fluorescence microscopy can also be applied to live cells. For example,
purified actin may be chemically linked to a fluorescent dye. Careful
biochemical studies have established that this “tagged”
molecule is indistinguishable in function from its normal counterpart. If
the tagged protein is microinjected into a cultured cell,
the endogenous cellular and injected tagged actin monomers copolymerize into
normal long actin fibers. This technique can also be used to study
individual microtubules within a cell.
Another technique for detecting specific proteins within living cells takes
advantage of green fluorescent protein (GFP), a naturally
fluorescent protein found in the jellyfish Aequorea
victoria. The bioluminescence of this organism, which radiates
a green fluorescence, is due to GFP. This 238-aa protein contains serine,
tyrosine, and glycine residues whose side chains have spontaneously reacted
with one another to form a fluorescent chromophore. By recombinant DNA
techniques discussed in Chapter
7, the GFP gene can be introduced into living cultured cells or
into specific cells of an entire animal. Because the introduced gene will
express GFP, the cells will emit a green fluorescence when irradiated; this
GFP fluorescence can be used to localize the cells within a tissue.
Figure 5-7
.
Use of green fluorescent protein (GFP) to localize GLUT4,
a glucose transport protein, within living fat cells
Cells were engineered to express a chimeric protein whose
N-terminal end corresponded to the GLUT4 sequence, followed
by the entirety of the GFP sequence. When a cell is exposed
to light of the exciting wavelength, GFP fluoresces
yellow-green, indicating the position of GLUT4 within the
cell. In resting cells (a), GLUT4 is in internal membranes
that are not connected to the plasma membrane. Successive
images of the same cell after treatment with insulin for
2.5, 5, and 10 minutes (panels b, c, and d, respectively)
show that, with time, increasing numbers of these
GLUT4-containing membranes fuse with the plasma membrane,
thereby moving GLUT4 to the cell surface and enabling it to
transport glucose from the blood into the cell. As detailed
in Chapter 20,
this is the principal mechanism by which insulin controls
the level of glucose in the blood. [Courtesy of J.
Bogan.]
View Movie: Reporter Constructs
Alternatively, the
gene for GFP can be fused to the
gene for another
protein
of interest, producing a
recombinant DNA encoding one long chimeric
protein
that contains the entirety of both
proteins. Cells in which this recombinant
DNA has been introduced will synthesize this chimeric
protein, whose green
fluorescence will reveal the subcellular localization of the
protein. illustrates how this
technique can demonstrate changes in the localization of a
protein within a
living cell following treatment with a particular
hormone.
Determining the Intracellular Concentration of Ca2+
and H+ Ions
Changes in the cytosolic concentration of Ca2+ ions or
pH frequently signal changes in cellular metabolism. The
Ca2+ concentration in the cytosol of resting cells,
for instance, is about 10−7 M. Many hormones or other
stimuli cause a rise in cytosolic Ca2+ to
10−6 M; this, in turn, causes changes in cellular
metabolism, such as contraction of muscle (Chapter 18).
Figure 5-8
.
Changes in the local concentration of
Ca2+in a sea urchin egg following
fertilization
The Ca2+ throughout the cell was monitored at
different times after fertilization using a fluorescence
microscope and fura-2, a Ca2+-binding dye
whose fluorescence is proportional to the
Ca2+ concentration. For graphic
purposes, the Ca2+ concentrations are
expressed in a calibrated color scale (right)
in units of micromolar Ca2+. When the sperm
penetrates the egg, the level of Ca2+ rises
initially at the point of sperm entry in the lower left part of
the cell and then gradually increases throughout the egg. This
spreading increase in cytosolic Ca2+
triggers the fusion of small vesicles with the plasma membrane,
causing changes in the cell surface that prevent penetration by
additional sperm. Eventually, the Ca2+
concentration becomes uniformly high and then falls uniformly to
the resting state. [See R. Y. Tsien and M. Poenie, 1986,
Trends Biochem. Sci.
11:450; courtesy of J. Alderton, M. Poenie, R. A.
Steinhardt, and R.Y.Tsien.]
The fluorescent properties of certain dyes, such as
fura-2,
facilitate measurement of the concentration of free Ca
2+
in the
cytosol. This dye contains five carboxylate groups that form ester
linkages with ethanol. The resulting fura-2 ester is
lipophilic and can
diffuse from the medium across the
plasma membrane into cells. Within the
cytosol, esterases hydrolyze fura-2 ester yielding fura-2, whose free
carboxylate groups render the molecule nonlipophilic, so it cannot cross
cellular
membranes and remains in the
cytosol. Each fura-2 molecule can bind
a single Ca
2+ ion but no other cellular cation, and the
amount of fura-2 bound to Ca
2+ is proportional, over a
certain range, to the Ca
2+ concentration. The
fluorescence of fura-2 at one particular wavelength is enhanced when
Ca
2+ is bound, and the fluorescence is proportional
to the Ca
2+ concentration. At another wavelength the
fluorescence of fura-2 is the same whether or not Ca
2+
is bound and provides a measure of the total amount of fura-2 in the segment
of the cell. By examining cells continuously in the fluorescence microscope
and measuring rapid changes in the ratio of fura-2 fluorescence at these two
wavelengths, one can quantify rapid changes in the fraction of fura-2 that
has a bound Ca
2+ ion and thus in the concentration of
cytosolic Ca
2+ ().
The fluorescence of other dyes is sensitive to the H+
concentration and can be used in a similar way to monitor the cytosolic pH
of living cells.
Confocal Scanning and Deconvolution Microscopy Provide Sharper Images of
Three-Dimensional Objects
Immunofluorescence microscopy has its limitations. The fixatives employed to
preserve cell architecture often destroy the
antigenicity of a
protein, that is, its ability to bind to its specific
antibody. Also, the method
generally gives poor results with thin cell sections, because embedding media
often fluoresce themselves, obscuring the specific signal from the
antibody.
Moreover, in microscopy of whole cells, the fluorescent light comes from
molecules above and below the plane of focus; thus the observer sees a
superposition of fluorescent images from molecules at many depths in the cell,
making it difficult to determine the actual three-dimensional molecular
arrangement (see ).
Figure 5-9
.
The advantage of confocal fluorescence microscopy
A mitotic fertilized egg from a sea urchin
(Psammechinus) was lysed with a detergent,
exposed to an anti-tubulin antibody, and then exposed to a
fluorescein-tagged antibody that binds to the first antibody. (a)
When viewed by conventional fluorescence microscopy, the mitotic
spindle is blurred owing to the background glow of fluorescence from
tubulin above and below the plane of focus. (b) The confocal
microscopic image is sharp, particularly in the center of the
mitotic spindle; fluorescence is detected only from molecules in the
focal plane. [From J. G. White, W. Amos, and M. Fordham, 1987,
J. Cell Biol.
104:41
The
confocal scanning microscope avoids the last problem by
permitting the observer to visualize fluorescent molecules in a single plane of
focus, thereby creating a vastly sharper cross-sectional image (). At any instant during
confocal imaging, only a single small part of a sample is illuminated with
exciting light from a focused laser beam, which rapidly moves to different spots
in the sample focal plane. Images from these spots are recorded by a video
camera and stored in a computer, and the composite image is displayed on a
computer screen.
Deconvolution microscopy is similar to confocal microscopy in
that a cross-sectional image is obtained, but the two techniques differ in the
details of how this image is generated. In both cases, the objective lens
collects light that originates from above and below the focal plane as well as
that which originates from within the focal plane. Confocal microscopes use a
pinhole to exclude the out-of-focus light. In contrast, deconvolution
microscopes collect all the light from several focal planes, and then
mathematically reassign the out-of-focus light to its correct focal plane with
the aid of a high-speed computer, a mathematical operation called
deconvolution.
To understand how a deconvolution microscope works, consider an infinitely small
fluorescent source of light, which can be approximated by a fluorescent bead
smaller than the resolution of the light microscope (i.e., <0.2 μm
in diameter). The emitted light radiates in all directions, and when the source
is in the focal plane of the objective, it appearsas a bright point of light.
When the point source is outside the focal plane of the objective, some of the
light is still collected by the objective lens, and the point source appears as
a halo. As the focal plane is moved farther away from the plane containing the
point source, the halo becomes larger and more diffuse. Knowing exactly how the
light emitted by an infinitely small fluorescent source is collected and
distorted by the optics of the sample and microscope, it is possible to
reconstruct an individual cross-sectional image (containing only light that
originated in the focal plane of interest) from a set of images taken as the
objective focal plane is moved through the plane of interest.
Figure 5-10
.
Optical sectioning of a developing Drosophila
egg chamber obtained with deconvolution fluorescence
microscopy
An egg chamber was labeled with the dye DAPI, which binds to DNA and
generates a blue fluorescence from the nuclei. Actin filaments were
labeled with the actin-binding chemical phalloidin coupled to
red-fluorescing rhodamine.
(Nuc = nucleus,
RC = ring cell,
NC = nurse cell, and
O = oocyte.) (a) A single optical
plane of an egg chamber. (b) A three-dimensional reconstruction of a
portion of the egg chamber shown in (a), consisting of stacked
serial optical sections obtained as above. Note the ring canals
(surrounded by actin filaments) that connect the nurse cells to one
another and to the developing oocyte. [Courtesy of D. Marcey.]
Cross-sectional images obtained with a deconvolution microscope may have even
greater detail than those obtained with a confocal microscope. Additionally, the
fluorescent labeling of the sample does not need to be as intense for
deconvolution microscopy as it does for confocal microscopy, since all the light
produced by a fluorescent sample is collected and analyzed by the microscope.
Three-dimensional images can be obtained by a refinement known as
optical sectioning. In this method, a computer records
individual fluorescent images of planes at different depths of the
sample — in effect, serial
sections — and combines the stack of images into
one three-dimensional image ().
Phase-Contrast and Nomarski Interference Microscopy Visualize Unstained
Living Cells
Figure 5-11
.
Light passing through a specimen can be redirected by refraction
and diffraction
(a) Refraction: Because light moves at different speeds in different
materials (more slowly in a medium of higher refractive index), a
beam of light is bent (refracted) as it passes from air into a
transparent object and bent again when it departs. Consequently, the
part of an incident light wave that passes through a specimen will
be refracted and will be out of phase (out of synchrony) with the
part of the wave that does not pass through the specimen. The
magnitude of the phase difference depends on the difference in
refractive index along the two paths and on the thickness of the
specimen. If the two parts of the light wave are recombined, the
resultant light will be brighter if they are in phase and less
bright if they are out of phase. (b) Diffraction: Light waves
impinging on a pinhole in an opaque object spread out in all
directions. Overlapping waves emanating from different sides of the
hole will reinforce one another in the directions (red arrows) where
the waves are in the same phase; to an observer in one of those
directions, the pinhole will seem bright. In other directions, where
the waves are out of phase, peaks of some light waves fall on
troughs of others and cancel one another out, producing dark areas.
These phenomena are called constructive and
destructive interference, respectively, and
explain the resulting diffraction patterns. Similarly, when light
impinges on an opaque object, the edges diffract the light waves,
producing an image that contains bright areas (white bands) when
viewed in some directions and dark areas (gray bands) in other
directions.
Detailed views of transparent, live, unstained cells and tissues are obtainable
with
phase-contrast microscopy and
Nomarski
interference microscopy. Both techniques take advantage of the
phenomena of refraction and diffraction of light waves (). As a result, small differences in
refractive index and thickness between parts of the specimen (say, between the
nucleus and
cytosol) or between the specimen and the surrounding medium can be
converted into differences of light and dark in the final image.
Figure 5-12
.
The optical pathway of the phase-contrast microscope
Incident light passes through an annular diaphragm, and the condenser
lens focuses a circular annulus (ring) of light on the sample. Light
that passes unobstructed through the specimen is focused by the
objective lens onto the thick gray ring of the phase plate, which
absorbs some of the direct light and alters its phase by one-quarter
of a wavelength. If a specimen refracts or diffracts the light, the
phase of some light waves is altered and the light waves are
redirected through the thin, clear region of the phase plate. The
refracted and unrefracted light are recombined at the image plane to
form the image.
Figure 5-13
.
Phase contrast and Nomarski optics
(a) Several live, cultured fat cells (adipocytes) viewed by
bright-field microscopy (left) and phase-contrast
microscopy (right). (b) Another specimen of
adipocytes viewed with Nomarski interference (differential
interference) microscopy. Thick black lines trace the surface
membrane of two cells. (c) A newly hatched larva of the nematode
Caenorhabditis elegans viewed with Nomarski
optics. The individual nuclei of many of the organism’s
959 cells are visible. [Part (a) courtesy of J. Bogan; part (b)
courtesy of P. Matsudaira and J. Bogan; part (c) from J. E. Sulston
and H. R. Horvitz, 1977, Devel. Biol.
56:110.]
The phase-contrast microscope generates an image in which the degree of darkness
or brightness of a region of the sample depends on the refractive index of that
region (). The improved
definition of subcellular structures in live, unstained cells obtained by
phase-contrast microscopy compared with standard bright-field microscopy is
illustrated in .
Nomarski, or differential, interference microscopy generates an image that looks
as if the specimen is casting a shadow to one side (): the “shadow” primarily
represents a difference in refractive index and thickness of a specimen rather
than its topography. In this technique, a prism splits an incident beam of
plane-polarized light so that one part of the beam passes through one region of
a specimen and the other part passes through a closely adjacent region; a second
prism then reassembles the two beams. Minute differences in thickness or in the
refractive index between adjacent parts of a sample are converted into a bright
image (if the two beams are in phase when they recombine) or a dark one (if they
are out of phase).
Phase-contrast microscopy is especially useful in examining the structure and
movement of larger
organelles, such as the
nucleus and mitochondria, in live
cultured cells. The greatest disadvantage of this technique is that it is
suitable for observing only single cells or thin cell layers. Nomarski
interference microscopy, in contrast, defines only the outlines of large
organelles, such as the
nucleus and vacuole. However, thick objects, such as the
nuclei in a worm, can be observed by combining this technique with optical
sectioning ().
Figure 5-14
.
Time-lapse micrographs show the movement of a cultured fibroblast
cell along a glass surface
A bit of debris on the substratum serves as a reference point. The
first image, at 0 min, was obtained by phase-contrast microscopy.
Successive images of the same cell, obtained by Nomarski optics,
show the lamella at the right of the cell retracting (R) and the
lamellipodia at the leading edge of the cell extending (E). In the
frame taken at 8 min, the leading edge has moved forward about 9
μm, and the lamellipodia there form a thin flat sheet. By 28
min, the broad leading edge has spread and separated into two
lamellae; the thin trailing edge of the cell has begun to retract
into the cell body. By 34 min, retraction of the trailing edge is
almost complete; only a thin thread of cytoplasm from the trail is
left behind, anchored to the substratum. [From W.-T. Chen, 1981,
J. Cell Sci.
49:1.]
Both phase-contrast and Nomarski interference microscopy can be used in
time-lapse microscopy, in which the same cell is
photographed at regular intervals over periods of several hours. This procedure
allows the observer to study cell movement, provided the microscope’s
stage can control the temperature of the specimen and the gas environment ().
Transmission Electron Microscopy Has a Limit of Resolution of 0.1 nm
The fundamental principles of electron microscopy are similar to those of light
microscopy; the major difference is that electromagnetic lenses, not optical
lenses, focus a high velocity electron beam instead of visible light. Because
electrons are absorbed by atoms in air, the entire tube between the electron
source and the viewing screen is maintained under an ultrahigh vacuum.
Figure 5-15
.
The optical path in a transmission electron microscope
A beam of electrons emanating from a heated tungsten filament is
focused onto the specimen plane by the magnetic condenser lens. The
electrons passing through the specimen are focused by a series of
magnetic objective and projector lenses to form a magnified image of
the specimen on a fluorescent viewing screen or a piece of
photographic film. The entire column, from the electron generator to
the screen, is maintained at a very high vacuum.
The
transmission electron microscope (TEM) directs a beam of
electrons through a specimen. Electrons are emitted by a tungsten cathode when
it is electrically heated. The electric potential of the cathode is kept at
50,000 – 100,000 volts; that of the anode, near
the top of the tube, is zero. This drop in voltage causes the electrons to
accelerate as they move toward the anode. A condenser lens focuses the electron
beam onto the sample; objective and projector lenses focus the electrons that
pass through the specimen and project them onto a viewing screen or a piece of
photographic film ().
In typical electron microscopes, electrons have the properties of a wave with a
wavelength of only 0.005 nm. Recall that the minimum distance D
at which two objects can be distinguished is proportional to the wavelength
λ of the light that illuminates the objects. Thus the limit of
resolution for the electron microscope is theoretically 0.005 nm (less than the
diameter of a single atom), or 40,000 times better than the resolution of the
light microscope and 2 million times better than that of the unaided human eye.
However, the effective resolution of the electron microscope in the study of
biological systems is considerably less than this ideal. Under optimal
conditions, a resolution of 0.10 nm can be obtained with transmission electron
microscopes, about 2000 times better than the best resolution of light
microscopes.
Preparation of Fixed, Stained Samples for TEM
Figure 5-16
.
Preparation of a sample of tissue for transmission electron
microscopy
The tissue is dissected, cut into small cubes, and plunged into a
fixing solution that cross-links and immobilizes proteins.
(Glutaraldehyde is frequently used; osmium tetroxide, another
fixing substance, also stains intracellular membranes and
certain macromolecules.) The sample is dehydrated by placing it
in successively more concentrated solutions of alcohol or
acetone; it is then immersed in a solution of plastic embedding
medium and put in an oven. Heat causes the solution to
polymerize into a hard plastic block, which is trimmed; sections
less than 0.1 μm thick are then cut with an
ultramicrotome, a fine-slicing instrument with a diamond blade
(a). The sections are floated off the blade edge onto the
surface of water in a trough. A copper grid coated with carbon
or some other material is used to pick up the sections, which
are then dried (b).
Like the light microscope, the transmission electron microscope is used to
view thin sections of a specimen, but the fixed sections must be much
thinner for electron microscopy (only
50 – 100 nm, about 0.2 percent of the
thickness of a single cell). Clearly, only a small portion of a cell can be
observed in any one section. depicts the preparation of a sample for transmission
electron microscopy. Generation of the image depends on differential
scattering of the incident electrons by molecules in the preparation.
Without staining, the beam of electrons passes through a cell or tissue
sample uniformly, so the entire sample appears uniformly bright with little
differentiation of components. Staining techniques are therefore used to
reveal the location and distribution of specific materials.
Figure 5-17
.
Localization of a specific protein by electron microscopy of
antibody-stained samples
(a) First the antibody is allowed to interact with its specific
antigen (e.g., catalase) in a section of fixed tissue. Then the
section is treated with a complex of protein A from the
bacterium Staphylococcus aureus and
electron-dense 5 – 7 nm diameter gold particles.
Binding of this complex to the common Fc domain of antibody
molecules makes the target protein, catalase in this case,
visible in the electron microscope. (b) A slice of liver tissue
was fixed with glutaraldehyde, sectioned, and then treated as
described in (a) to localize catalase. The gold particles (black
dots) indicating the presence of catalase are located
exclusively in peroxisomes. [From H. J. Geuze et al., 1981,
J. Cell Biol.
89:653. Reproduced from the Journal of Cell
Biologyby copyright permission of The Rockefeller
University Press.]
Heavy metals, such as gold or osmium, appear dark on a micrograph because
they scatter (diffract) most of the incident electrons; scattered electrons
are not focused by the electromagnetic lenses and do not form the image.
Osmium tetrox-ide preferentially stains certain cellular components, such as
membranes, which appear black in micrographs. Specific
proteins can be
detected in thin sections by use of electron-dense gold particles coated
with
protein A, a bacterial
protein that binds
antibody molecules
nonspecifically ().
Figure 5-18
.
Metal shadowing, a technique that makes surface details on
very small particles visible in the electron microscope
The sample is spread on a mica surface and then dried in a vacuum
evaporator. A filament of a heavy metal, such as platinum or
gold, is heated electrically so that the metal evaporates and
some of it falls over the sample grid in a very thin film. In
order to stabilize the replica, the specimen is then coated with
a carbon film evaporated from an overhead electrode. The
biological material is then dissolved by acid, so that the
observer views only the metal replica of the sample. In electron
micrographs of such preparations, the image is usually reversed:
carbon-coated areas appear light and platinum-shadowed areas
appear dark.
Electron microscopy also is used to obtain information about the shapes of
purified
viruses, fibers,
enzymes, and other subcellular particles. In one
technique, called
metal shadowing, a thin layer of
evaporated metal, such as platinum, is laid at an angle on a biological
sample (). An
acid bath
dissolves the biological material, leaving a metal replica of its surface,
which can then be examined in the transmission electron microscope.
Variations in the angle and thickness of the deposited metal allow an image
to be formed because some incident electrons will be scattered in various
directions rather than pass through the preparation. If the metal is
deposited mainly on one side of the sample, for instance, the image seems to
have “shadows,” where the metal appears dark and the
shadows appear light.
Cryoelectron Microscopy
Figure 5-19
.
Cryoelectron micrograph of unstained rotavirus
particles
A thin suspension of virus particles in water is applied to an
electron microscopy grid and frozen. It is then visualized in a
transmission electron microscope equipped with a sample stage
cooled with liquid nitrogen. The low temperature prevents the
ice surrounding the particles from evaporating in the vacuum.
Because many biological specimens scatter more electrons than
water does, the investigator can observe a very thin specimen
without fixing, staining, or dehydrating it. Note the minute
spikes (arrows) visible on some particles. [From B. V.
Venkataram Prasad et al., 1988, J. Mol. Biol.
199:269.]
Standard electron microscopy cannot be used to study live cells because they
are generally too vulnerable to the required conditions and preparatory
techniques. In particular, the absence of water causes
macromolecules to
become denatured and nonfunctional. However, the technique of
cryoelectron microscopy allows examination of hydrated,
unfixed, and unstained biological specimens directly in the transmission
electron microscope. In this technique, an aqueous suspension of a sample is
applied in an extremely thin film to a grid. After it has been frozen in
liquid nitrogen and maintained in this state by means of a special mount, it
is observed in the electron microscope. The very low temperature
(−196 °C) keeps the water from evaporating, even in a
vacuum, and the sample can be observed in detail in its native, hydrated
state without shadowing or fixing it (). By computer-based averaging of images of hundreds
of particles, a three-dimensional model almost to atomic
resolution can be
generated.
Scanning Electron Microscopy Visualizes Details on the Surfaces of Cells and
Particles
Figure 5-20
.
Scanning electron micrograph of a tendon located within the
shoulder of a 29-year-old male
The sample was rapidly frozen and metal-shadowed before viewing in a
scanning electron microscope. These long collagen fibrils are of
variable diameter and are oriented parallel to one another; similar
fibrils also form the major structural element of bone and similar
tissues. The side-by-side arrangement of the linear collagen
molecules comprising a fibril gives rise to a characteristic 64-nm
repeated pattern visible as parallel striations along the length of
each fibril. [Courtesy of D. Keene.]
The
scanning electron microscope allows the investigator to view
the surfaces of unsectioned specimens. These cannot be visualized with
transmission equipment because the electrons pass through the entire specimen.
The sample is fixed, dried, and coated with a thin layer of a heavy metal, such
as platinum, by evaporation in a vacuum (see ); in this case, the sample is rotated so that the
platinum is deposited uniformly on the surface. An intense electron beam inside
the microscope scans rapidly over the sample. Molecules in the specimen are
excited and release secondary electrons that are focused onto a scintillation
detector; the resulting signal is displayed on a cathode-ray tube. Because the
number of secondary electrons produced by any one point on the sample depends on
the angle of the electron beam in relation to the surface, the scanning electron
micrograph has a threedimensional appearance (; see also ). The resolving power of scanning electron microscopes, which is
limited by the thickness of the metal coating, is only about 10 nm, much less
than that of transmission instruments.
SUMMARY
-
Various microscopic techniques generate
different views of the cell and have different resolutions. The limit of
resolution of a light microscope is about 0.2 μm; of a
transmission electron microscope, about 0.1 nm; and of a scanning
electron microscope, about 10 nm.
-
Standard (bright-field) light microscopy is
best for stained or colored cells or tissue sections.
-
Fluorescence microscopy allows specific
proteins and organelles to be detected in fixed cells stained with a
fluorescent dye or fluorescent-labeled antibodies (immunofluorescence
microscopy). The movements of microinjected or expressed recombinant
fluorescent proteins also can be followed in living cells.
-
By use of dyes whose fluorescence is
proportional to the concentration of Ca2+ or
H+ ions, fluorescence microscopy can measure the
local concentration of Ca2+ ions and intracellular
pH in living cells.
-
Confocal imaging, which allows the observer
to view fluorescent molecules in a single plane of a specimen, permits
optical sectioning of the sample and produces very sharp images.
-
Phase-contrast and Nomarski optics enable
scientists to view the details of live, unstained cells and to monitor
cell movement.
-
Specimens for electron microscopy generally
must be fixed, sectioned, and dehydrated, and then stained with
electron-dense heavy metals.
-
Surface details of particles such as
viruses and collagen fibers can be revealed by electron microscopy of
metal-shadowed specimens (see ). -
Unfixed, unstained specimens can be viewed
in the electron microscope if they are frozen in hydrated form, a
technique called cryoelectron microscopy.
-
The scanning electron microscope can be
used to view unsectioned cells or tissues; it produces images that
appear to be three-dimensional.
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