As each of the electrons of the primary beam strike the specimen they
are deflected and slowed through interactions with the
atoms of the sample. In order to calculate a hypothetical trajectory
of a primary beam electron within a specimen a "Monte
Carlo" simulation is performed. Using values for mean free path, angle
of deflection, change in energy, and likelihood of a given
type of collision event for a primary electron, the trajectory can
be approximated using a random number factor (hence the
name Monte Carlo) to predict the type of collision.
[Fig 3.5a Gold]
By performing this simulation for a number (100 or greater) of primary
electrons of a given energy striking a specimen of
known composition, the geometry of the region of primary electron interaction
can be approximated.
[Fig. 3.5b Gold]
The size and shape of the region of primary excitation is dependent
upon several factors, the most important of which is the
composition of the specimen and the energy with which the primary electrons
strike the sample. A primary electron beam with
a high accelerating voltage will penetrate much more deeply into the
sample than will a beam of lower energy.
[Illustrate]
Region of Primary Excitation Cont'd:
Likewise, the shape of the primary excitation zone will vary depending
on the atomic weight of the specimen. Materials that
have a higher atomic number are significantly more likely to collide
with the primary electron beam than those of a low atomic
weight. This will cause the electron to undergo more interactions (shorter
mean free path), of a different nature (greater change
in angle and loss of energy) than would the same electron in a specimen
of lower atomic number. A beam interacting with such
a sample would therefore not penetrate as deeply as it would into a
specimen of a lower atomic weight.
[Illustration]
Another factor that affects the geometry of the primary excitation zone
is the incoming angle of the incident beam. Because the
tendency of the electrons to undergo forward scattering causes them
to propagate closer to the surface than a head on beam
the resulting signal comes from a slightly smaller area. This is another
reason for tilting the sample slightly towards the detector.
Finally, the dimensions of the tear-drop zone are dependent on the diameter
of the incoming spot. The smaller the initial spot,
the smaller will be the region of primary excitation. Because the tear-drop
zone is always larger than the diameter of the
primary beam spot this explains why the resolution of an SEM is not
equivalent to the smallest beam spot but is proportional to
it.
Of the various types of signals produced from interactions of the primary
beam with the specimen, each has a different amount
of energy associated with it. Because of this and because different
signals are more or less permeable to the sample, different
signals are emitted from different regions of the region of primary
excitation. At the top of the tear drop near the very surface of
the specimen is the region from which Auger electrons are emitted.
Because they have such a low energy, Auger electrons
cannot escape from very deep in the sample even though they may be
created there by primary or even backscattered
electrons. This narrow escape depth explains why Auger electron spectroscopy
is only useful for resolving elements located in
the first monolayer of a specimen and why their resolution is nearly
the same as size of the primary electron beam. Beneath the
region from which Auger electrons are emitted is the region of secondary
electron emission. Because they have a higher energy
and therefore a greater escape velocity the region of secondary electron
emission is not only deeper into the specimen but
broader in diameter than the zone of Auger electron emission. The two
regions are not mutually exclusive and secondary
electrons are emitted from the uppermost elements of the sample as
well.
Region of Primary Excitation Cont'd:
[Figure 3-5 here]
Backscattered electrons have an even greater energy than either secondary
or Auger electrons. Consequently they are capable
of escaping from even greater depths within the sample. For this reason
the depth and diameter of the region from which
backscattered electrons are emitted is greater than that for secondary
electrons and the resulting resolution from a backscatter
image is that much less. The deepest usable signal caused by penetration
of the primary beam comes in the form of
characteristic X-rays. Because the final size of such an X-ray emission
zone is so large that the resolution that can be obtained
is usually quite poor. Despite this however, characteristic X-rays
can provide valuable information about the chemical
composition of a specimen even in cases where a thin layer of some
other material (i.e. gold-palladium) may be deposited on
top. One other signal, the "white X-rays" or "X-ray continuum" is also
produced when the nucleus of an atom scatters electrons
(primary or backscattered) and releases excess energy. Because it is
not characteristic of the element that formed it, the X-ray
continuum is merely a form of background signal that must be accounted
for in measuring characteristic X-rays.
[Text pages 47-49]
Ultimately image formation in an SEM is dependent on the acquisition
of signals produced from the interaction of the specimen
and the electron beam. These interactions can be broken down into two
major categories 1) those that result in elastic
collisions of the electron beam on the sample [where instantaneous
energy Ei = Eo or initial energy] and 2) those that result in
inelastic collisions [where Ei < Eo]. In addition to those signals
that are utilized to form an image, a number of other signals are
also produced when an electron beam strikes a sample. We will discuss
a number of these different types of beam-specimen
interactions and how they are utilized. But first we need to examine
what actually happens when a specimen is exposed to the
beam.
[fig. 3.1 Goldstein]
To begin with we refer to the illumination beam as the "primary electron
beam". The electrons that comprise this beam are thus
referred to as being primary electrons. Upon contacting the specimen
surface a number of changes are induced by the
interaction of the primary electrons with the molecules contained in
the sample. Upon contacting the surface of the specimen
most of the beam is not immediately bounced off in the way that light
photons might be bounced off in a light dissecting
microscope. Rather the energized electrons penetrate into the sample
for some distance before they encounter an atomic
particle with which they collide. In doing so the primary electron
beam produces what is known as a region of primary
excitation. Because of its shape this region is also known as the "tear-drop"
zone. A variety of signals are produced from this
zone, and it is the size and shape of this zone that ultimately determines
the maximum resolution of a given SEM working with a
particular specimen.
[Insert generalized tear-drop diagram here]
The various types of signals produced from the interaction of the primary
beam with the specimen include Secondary electron
emission, backscatter electrons, Auger electron, characteristic X-rays,
and cathodluminescence. We will discuss each of these
in turn.
The most widely utilized signal produced by the interaction of the primary
electron beam with the sample is the secondary
electron emission signal. A secondary electron is produced when an
electron from the primary beam collides with an electron
from a specimen atom and loses energy to it. This will serve to ionize
the atom and in order to re-establish the proper charge
ratio following this ionization event an electron may be emitted. Such
electrons are referred to as "secondary" electrons.
Secondary electrons are characterized from other electrons by having
an energy of less than 50 eV.
[Diagram of Atom and collision of electrons in an outer shell]
Secondary Electrons Cont'd:
This is by far the most common type of image produced by modern SEMs.
It is most useful for examining surface structure and
gives the best resolution image of any of the scanning signals. Depending
on the initial size of the primary beam and various
other conditions (composition of sample, accelerating voltage, position
of specimen relative to the detector) a secondary
electron signal can resolve surface structures down to the order of
10 nm or better. The topographical image is dependent on
how many of the secondary electrons actually reach the detector. Although
an equivalent number of secondary electrons might
be produced as a result of collisions between the primary electron
beam and the specimen, secondary electrons that are
prevented from reaching the detector will not contribute to the final
image and these areas will appear as shadows or darker in
contrast than those regions that have a clear electron path to the
detector.
[diagram 2-24 here]
One of the major reasons for sputter coating a non-conductive specimen
is to increase the number of secondary electrons that
are emitted from the sample.
Secondary Electron Detector:
In order to detect the secondary electrons that are emitted from the
specimen a specialized detector is required. This is
accomplished by a complex device that first converts the energy of
the secondary electrons into photons. It is referred to as a
scintillator-photomultiplier detector or "Everhart- Thornley" detector.
The principle component that achieves this is the
scintillator. The scintillator is composed of a thin plastic disk that
is coated or doped with a special phosphor layer that is highly
efficient at converting the energy contained in the electrons into
photons. When this happens the photons that are produced
travel down a Plexiglas or polished quartz light pipe and out through
the specimen chamber wall. The outer layer of the
scintillator is coated with a thin layer [10-50 nm] of aluminum. This
aluminum layer is positively biased at approximately 10 KV
and helps to accelerate the secondary electrons towards the scintillator.
The aluminum layer also acts as a mirror to reflect the
photons produced in the phosphor layer down the light pipe. The photons
that then travel down the light pipe are amplified into
an electronic signal by way of a photocathode and photomultiplier.
The signal thus produced can now be used to control the
intensity of brightness on the CRT screen in proportion to the number
of photons originally produced.
In-Lens Detector:
The ability to image a specimen in the SEM is often limited not so much
by the specimen or the signal it produces but the ability
of the detector to collect this signal. This becomes a critical issue
at very short working distances (5 nm or less) which are
necessary for very high resolution work. A secondary electron detector
positioned to the side of the specimen is sometimes
blocked from receiving signal by the specimen and stage itself. This
is similar to the situation with a specimen that has a deep
cavity from which signal cannot escape despite the fact that it is
producing a significant amount of signal.
One attempt to overcome this limitation in signal collection is to place
a secondary electron detector within the final lens of the
SEM. In this way the detector is on nearly the same optical axis as
the primary beam itself thus the position of the detector
relative to the source of the signal is not the limiting factor in
signal detection. Because the secondary electron detector does not
need to be positioned between the specimen and the final lens very
short working distances can be used and very high
resolution obtained. The secondary electrons of the signal can be distinguished
from the electrons of the primary beam by both
their significantly lower energy and their directional vector (i.e.
opposite in direction to those of the primary beam. The
secondary electrons produced by the specimen do not interfere with
the primary beam electrons, the situation being analogous
to shooting a water pistol into the air during a driving rainstorm.
The chances of water droplets in from the water pistol actually
hitting the individual raindrops is vanishingly small despite the greater
numbers and significantly higher energy of the rainstorm.
Like the electrons of the primary beam, the secondary signal electrons
are focused by the electromagnetic field of the final lens
and concentrated into a smaller area. A converging lens works the same
way regardless of the direction from which the
electrons enter the lens. Thus the final lens acts somewhat like a
signal collector, concentrating the secondary electrons before
detection by the in-lens detector.
[insert Fig. 2-20 here]
A photomultiplier tube or PMT consists of a cathode which converts the
quantum energy contained within the photon into an
electron by a process known as electron-hole replacement. This generated
electron then travels down the PMT towards the
anode striking the walls of the tube as it goes. The tube is coated
with some material (usually an oxide) that has a very low
work function and thus generates more freed electrons. This results
in a cascade of electrons and eventually this amplified signal
strikes the anode. The anode then sends this amplified electrical signal
to further electrical amplifiers. The number of cascade
electrons produced in the PMT is dependent on the voltage applied across
the cathode and anode of the PMT. Thus it is in the
PMT that the light produced by the scintillator detector is amplified
into electrical signal and thus producing gain. We can turn
up the gain by increasing the voltage to the PMT which is essentially
what we do when we adjust the contrast. The electrical
amplifier increases the electrical signal from the PMT by a constant
amount thus increasing or brightness.
[illustration of PMT]
Because secondary electrons are emitted from the specimen in an omni
directional manner and possess relatively low energies
they must be in some way collected before they can be counted by the
secondary electron detector. For this reason the
secondary electron detector is surrounded by a positively charged anode
or Faraday cup or cage that has a potential charge on
it in the neighborhood of 200 V. This tends to draw in many of the
secondary electrons towards the scintillator. It also helps to
alleviate some of the negative effects of the scintillator aluminum
layer bias which because it is so much greater (10 KV vs. 200
V) can actually distort the incident beam. A second type of electron,
the backscattered electron [which we will discuss later], is
also produced when the specimen is irradiated with the primary electron
beam. Together backscattered and secondary
electrons contribute to the the signal that reaches the scintillator
and form what we refer to as the secondary electron image.
A rather new usage of secondary electrons is employed in "Environmental
SEMs." Unlike a conventional SEM the
environmental SEM is designed to image specimens that are not under
vacuum. In fact for an environmental SEM to function
properly there must be air or some other gas molecules present in the
specimen chamber. The way an environmental SEM
works is by first generating and manipulating a primary beam in much
the same way as in a conventional SEM. The primary
beam then enters the specimen chamber through a pressure limiting aperture
(PLA) that is situated beneath the final lens pole
piece. This PLA allows the chamber to be kept at one pressure (e.g.
0.1 ATM) while the rest of the column is at a much higher
vacuum (e.g. 10-6 Torr). The primary beam strikes the specimen and
produces secondary and backscattered electrons in the
same manner as does a conventional SEM. The difference is that these
secondary electrons then strike gas molecules in the
specimen chamber which in turn produce their own secondary electrons
or "environmental electrons." This results in a
cascading or propagation effect and greatly increases the amount of
signal. It is all of these electrons that are then used as signal
by the detector that is positioned near the final aperture. Because
of this unique design wet or even uncoated living specimens
can be imaged in a SEM. There are however, some very real drawbacks.
Backscattered Electrons: [Text pages 54-56]
A backscatter electron is defined as one which has undergone a single
or multiple scattering events and escapes with an energy
greater than 50 eV. Backscattered electrons are produced as the result
of elastic collisions with the atoms of the sample and
usually retain about 80% of their original energy. The number of backscattered
electrons produced increases with increasing
atomic number of the specimen. For this reason a sample that is composed
of two or more different elements which differ
significantly in their atomic numbers, will produce an image that shows
differential contrast of the elements despite a uniform
topology. Elements that are of a higher atomic number will produce
more backscattered electrons and will therefore appear
brighter than neighboring elements.
[Illustration here]
The region of the specimen from which backscattered electrons are produced
is considerably larger than it is for secondary
electrons. For this reason the resolution of a backscattered electron
image is considerably less (1.0 um) than it is for a
secondary electron image (10 nm). Because of their greater energy,
backscattered electrons can escape from much deeper
regions of the sample than can secondary electrons hence the larger
region of excitation. By colliding with surrounding atoms of
the specimen some backscattered electrons can also produce X-ray, Auger
electrons, cathodluminescence, and even
additional secondary electrons.
The detector for backscattered electrons is similar to that used in
the detection of secondary electrons in that both utilize a
scintillator and photomultiplier design. The backscatter detector differs
in that a biased Faraday cage is not employed to attract
the electrons. Only those electrons that travel in a straight path
from the specimen to the detector go towards forming the
backscattered image. So that enough electrons are collected to produce
an image, many SEMs use multiple backscattered
detectors positioned directly or nearly above the specimen.
[Diagram 3-8 here]
Backscattered Electrons Cont'd:
By using these detectors in pairs or individually, backscattered electrons
can be used to produce a topographical image that
differs from that produced by secondary electrons. Another type of
backscatter detector uses a large angle scintillator or
"Robinson" detector that sits above the specimen. Shaped much like
a doughnut the beam enters through the center hole and
backscattered electrons are detected around its periphery.
[Draw Diagram here]
Because some backscattered electrons are blocked by regions of the specimen
that secondary electrons might be drawn
around, this type of imaging is especially useful in examining relatively
flat samples.
[Draw Diagram here]
Characteristic X-rays: [Text pages 56-58]
Another class of signals produced by the interaction of the primary
electron beam with the specimen come under the category
of characteristic X- rays. When an electron from an inner atomic shell
is displaced by colliding with a primary electron, it leaves
a vacancy in that electron shell. In order to re-establish the proper
balance in its orbitals following an ionization event, an
electron from an outer shell of the atom may "fall" into the inner
shell and replace the spot vacated by the displaced electron. In
doing so the this falling electron loses energy and this energy is
referred to as X- radiation or X-rays.
The SEM can be set up in such a way that the characteristic X-ray of
a given element is detected and its position recorded or
"mapped." These X-ray maps can be used to form an image of the sample
that shows where atoms of a given element are
localized. The resolution of these X-ray maps is on the order of greater
than 1 um.
[Diagram here]
In addition to characteristic X-rays, other X-rays are produced as a
primary electron decelerates in response to the Coulombic
field of an atom. This "braking radiation" or Bremsstrahlung X-ray
is not specific for the element that causes it and so these
X-rays do not contribute useful information about the sample and in
fact contribute to the background X-ray signal.
Auger Electrons: [Text pages 58-61]
Auger electrons are produced when an outer shell electron fills the
hole vacated by an inner shell electron that is displaced by a
primary or backscattered electron. The excess energy released by this
process may be carried away by an Auger electron.
[Diagram]
Because the energy of these electrons is approximately equal to the
difference between the two shells, Like X-rays an Auger
electron can be characteristic of the type of element from which it
was released and the shell energy of that element. By
discriminating between Auger electrons of various energies Auger Electron
Spectroscopy (AES) can be performed and a
chemical analysis of the specimen surface can be made. Because of their
low energies, Auger electrons are emitted only from
near the surface. They have an escape depth of between 0.5 to 2 nm
making their potential spatial resolution especially good
and nearly that of the primary beam diameter. One major problem associated
with this is the fact that most SEMs deposit small
amounts (monolayers) of gaseous residues on the specimen which tend
to obscure those elements on the surface. For this
reason an SEM that can achieve ultrahigh vacuum (10-10 Torr) are required.
Also the surface contaminants of the specimen
must be removed in the chamber to expose fresh surface. To accomplish
this further modifications to the SEM (ion etching,
high temperature cleaning, etc.) are needed.
Unlike characteristic X-rays, Auger electrons are produced in greater
amounts by elements of low atomic number. This is
because the electrons of these elements are less tightly bound to the
nucleus than they are in elements of greater atomic number.
Still the sensitivity of AES can be exceptional with elements being
detected that are only present in hundreds of parts per million
concentration.
Cathodluminescence: [Text pages 61-62]
Certain materials (notably those containing phosphorous) will release
excess energy in the form of photons when electrons
recombine to fill holes made by the interaction of the primary beam
with the specimen. By collecting these photons using a light
pipe and photomultiplier similar to the ones utilized by the secondary
electron detector, these photons of visible light energy can
be detected and counted. An image built up in the same point by point
manner that all other scanning micrographs are. Thus
despite the similarity of using a signal of light to form the final
image resolution and image formation are unlike the image formed
in a light optical microscope. The best possible image resolution using
this approach is estimated at about 50 nm.
[insert fig. 3-16]
Specimen Current: [Text pages 63-65]
One rather elegant method of imaging a specimen is by means of measuring
specimen current. Specimen current is defined as
the difference between the primary beam current and the total emissive
current (= Backscatter + secondary + Auger
electrons). Thus specimens that have stronger emissive currents have
weaker specimen currents and vice versa. Imaging by
way of specimen current has the advantage that the relationship of
the detector to the position of the specimen is irrelevant
since the detector and is actually within the specimen. It is most
useful for imaging material mosaics at very small working
distances.
[Figure 3-20 here]
Yet another method that can be used in the SEM to create an image is
that of transmitted electrons. Like the secondary and
backscatter electron detectors, the transmitted electron detector is
comprised of scintillator, light pipe (or guide), and a
photomultiplier. The transmitted electron detector differs primarily
in its position relative to the specimen.
X-ray Microanalysis [text 332-344]
Another class of signals produced by the interaction of the primary
electron beam with the specimen come under the category
of characteristic X- rays. When an electron from an inner atomic shell
is displaced by colliding with a primary electron, it leaves
a vacancy in that electron shell. In order to re-establish the proper
balance in its orbitals following an ionization event, an
electron from an outer shell of the atom may "fall" into the inner
shell and replace the spot vacated by the displaced electron. In
doing so the this falling electron loses energy and this energy is
referred to as X- radiation or X-rays.
In addition to characteristic X-rays, other X-rays are produced as a
primary electron decelerates in response to the Columbic
field of an atom. This "braking radiation" or Bremsstrahlung X-ray
is not specific for the element that causes it and so these
X-rays do not contribute useful information about the sample and in
fact contribute to the background X-ray signal.
For each electron/specimen interaction there are specific electron replacement
events that can take place. We speak of these
events as either K, L, or M replacement events depending on which orbital
shell lost the electron.
[Fig. 15-8]
We can further dissect these electron replacement events by speaking
of them in terms of which outer orbital electrons served
as the replacement for the displaced electron. If the replacement electron
came from an adjacent orbital shell it is an alpha
event, if it came from two shells away it is a beta event, if the electron
was donated from three shells away it is a gamma event.
Within a given shell there may be several different orbitals, any of
which could donate the replacement electron. Thus we could
speak of a K alpha1 or a K alpha2 replacement. The important thing
to note is that each electron replacement event for each
element gives off a specific amount of energy as the replacement electron
goes from a higher energy state to a lower energy
state. This change in energy is released in the form of x-rays and
because of the specific nature of these x-rays they are call
"characteristic x-rays."
By using special detectors that can discriminate between the different
characteristic x-rays one can obtain information about the
elemental composition of the specimen. Let's assume that a plant cell
is found to have in thin sections an electron dense
inclusion of unknown composition. By bombarding the inclusion with
electrons from the beam we drive off a number of
electrons which are replaced by outer orbital electrons and give off
characteristic x-rays for the elements in the specimen. Since
we are primarily interested in the composition of the inclusion and
not the surrounding tissue it is very beneficial to be able to
focus the beam to a single spot and position this over the object on
interest. This can best be done in a Scanning Transmission
Electron Microscope or STEM. A STEM is equipped with a set of scan
coils and can function in much the same way as an
SEM by rastering the beam (reduced to a small spot) over the specimen
which in this case would be a section on a grid. Also
because more of a sample will contain more of the material in question
we tend to cut thicker sections for x-ray microanalysis
than we would for straight visualization. Sections of 100-250 nm are
typically used. Finally, because certain elements produce
their own characteristic x-rays that may interfere with or obscure
the ones of the unknown sample, we tend to avoid osmicating
the specimen and avoid UA and lead staining. Also the choice of metal
grid can be important as grids composed of one metal
(e.g. nickel) may not overlap whereas others (e.g. copper) may.
By collecting the x-ray signals produced over an extended period of
time (e.g. 100 seconds) certain electron replacement
events will occur more frequently than others. We collect for 100 seconds
of longer so that the more frequent events will
reinforce each other and thereby become distinct from the background
(characteristic x-rays from other elements) and
continuum x-rays. These repeated energy spectra manifest themselves
in the form of distinct peaks. Next with the aid of a
computer we can assign a numerical value for the midpoint of each peak
and scan through the values from known samples to
find the most logical match to our observed spectra. In trying to assign
a match it is important to note that for a given element
there will be more K alpha events than K beta events, and more K beta
events than K gamma events. Thus if one suspects the
presence of a given element due to the match between a collected peak
and the K beta peak of a known element, there should
be a corresponding K alpha peak for that element that is larger than
the suspected K beta peak.
There are basically two types of x-ray detectors available for TEM and
SEM. These are Energy Dispersive X-ray (EDX)
detectors and Wavelength Dispersive X- ray (WDS) detectors. They function
in quite different ways.
EDX detectors are the most versatile, cost effective
and hence most widely used type of x-ray detectors. EDX detectors are
composed of a silicon semi- conductor that has been doped with lithium
and are therefore referred to as SiLi detectors. The
EDX detector works by measuring the change in conductivity that occurs
when the semi-conductor absorbs excess energy in
the form of x-radiation. The conductivity increase is directly proportional
to the magnitude of the x- rays and so by carefully
measuring this increase one can knows the level of x- radiation that
was absorbed by the detector. Since these changes are still
relatively small the detector is kept a liquid nitrogen temperatures
to reduce electronic noise that would degrade peak
resolution. Since the internal environment of the TEM is subject to
minor (introduction of specimen) to major (venting of the
column) changes the detector must be kept in an exceptionally clean
environment. To do this it is typically shielded behind a
very thin seal or window. Beryllium is the material of choice for such
a window. Because of its low atomic weight (4) beryllium
will not block the x-rays of higher energy that are produced by elements
of higher atomic weight. It will however reduce the
ability to detect x-rays of relatively low energy (such as those given
off by elements of low atomic weight) and make the
detection of "light" elements more problematic. To avoid this some
systems have gone over to a windowless detector which
depends on the purity of the TEM environment from contaminating the
cold EDX detector.
The second type of x-ray detector is base on WDS.
In WDS crystals of known composition and structure are placed on a
movable turret relative to the x-ray source and a simple detector (alternatively
the detector itself is movable relative to the
crystal. Electrons and x-rays will move through a crystal and be reflected
or diffracted based on the particular arrangement of
molecules in that crystal. Only those energy sources entering from
a specific angle relative to the matrix arrangement of the
crystal will be so deflected. The angle by which this takes place is
known as the "Bragg angle" and is dependent to some extent
on the energy of the incoming radiation.
where = an integer (1, 2, 3, etc.)
= the x-ray wavelength
= the interplanar spacing of the crystal
= angle of incidence
[Fig 5.2 Goldstein]
The crystal is often polished to a curved surface so that the collected
x- rays can be focused onto the actual detector. The
detector is a thin wire kept at a high positive voltage in an argon/methane
environment. As the x-rays pass through a thin plastic
window they ionize the gas mixture and conduct electrons to the wire.
This current flow is carefully measured and is
proportional to the energy of the x-ray which in turn reveals information
about the source of the x- rays.
WDX is more quantitative than EDX but has a number of disadvantages
to it. The most important of these is the fact that each
type of crystal has a relatively narrow x-ray energy range that it
can deflect. Thus each crystal and detector can only detect a
small range of elements whereas an EDX detector can detect nearly the
entire spectrum of elements. Because of this one often
needs a suite of WDX detectors, each with a different type of crystal
and responsible for a different portion of the periodic
table. This means having a number of open ports available near the
specimen. On most TEMs we do not have this luxury and
so WDX systems are usually found on a special class of SEM known as
a microprobe. X-ray analysis on TEM and STEM is
usually accomplished with an EDX system.
Electron Diffraction [text 347-355]
We have seen several methods whereby we can learn more about the specimen
than just its appearance. Although not widely
used by biologists electron diffraction is a powerful TEM technique
that can provide important information about the molecular
arrangement of crystalline specimens.
Electrons are forward scattered or "diffracted" as they come in contact
with molecules in the specimen. Most of the time this
results in a random deflection of the illuminating electrons and creates
a fuzzy or muddled quality to the final image. This is
caused by the fact that a deflected electron will create just as bright
a spot on the fluorescent screen or TEM film as will an
undeflected electron. Because a randomly scattered electron may hit
the screen in a region that would normally appear dark
due to the presence of an electron dense body immediately above it.
To reduce the effect of these randomly scattered electrons
one typically places a small diameter aperture in the objective lens
immediately beneath the specimen. Although this reduces
overall illumination and reduces resolution by decreasing the angle
of the cone of incident illumination, it increases image
contrast by eliminating most of the forward scattered electrons.
[draw diagram]
The situation is quite different when the electrons of the beam encounter
a crystalline specimen. A crystalline specimen is one in
which the molecules of the specimen are arranged in such a way as to
form a close-packed lattice array with individual
molecules arranged in a very ordered and repetitive structure. If the
electrons strike a crystalline structure at the proper angle
they will all be diffracted from the individual planes of the lattice
in the same angle and same direction and brought to the same
focal point. This focal point lies in the same plane as the one in
which transmitted electrons come to focus and is known as the
back focal plane of the objective lens.
[Fig. 15-26]
Electron Diffraction Cont'd
The angle at which the incident electrons encounter the specimen is
the most critical parameter in creating a sharp electron
diffraction pattern. This angle is known as the Bragg Angle. A crystalline
specimen that is placed on grid may intially lie at any
random angle relative to the incident beam. To orient the specimen
so that the incident beam strikes at the proper Bragg angle
and generates a sharp diffraction pattern it is necessary to tilt and
rotate the specimen until a clear pattern is formed. If the
beam strikes the lattice at the proper Bragg angle electrons that are
scattered from the same point in the specimen are brought
together at a single point in the image plane. Likewise electrons scattered
from different points in the specimen BUT deflected
in the same direction and angle will converge in the back focal plane
of the objective lens.
[Fig. 15-30]
If one can obtain a picture of this pattern and carefully measure the
spacings between these spots of convergence much can be
learned about the molecular structure and composition of the specimen.
The spacing between lattice planes can be calculated from the diffraction pattern using the following equation:
d= L/R
Where d = Spacing between planes
= Wavelength of electron (based accelerating voltage)
L = Camera length (distance in mm between specimen and camera)
R = Distance from center spot to bright dots on negative
It is important that these calculations be done on the negative itself.
If done on prints the exact enlargement factor must be
known so that the R measurements can divided by this number. Since
the camera length is a critical portion of this equation it
should be regularly calibrated. This is done not by measuring with
a ruler but by creating a diffraction pattern with a standard
sample of known d spacing at a given accelerating voltage and calculating
the value for L by plugging R into the equation.
Electron Energy Loss Spectroscopy (EELS) text [344-347]
and Electron Spectroscopic Imaging (ESI)
When an electron of the primary beam interacts with the elements of
a specimen one of several things can happen. First, it may
pass by without altering either its energy (wavelength) or trajectory.
These non-scattered electrons are what are primarily
responsible for creating the bright portion of a TEM image as they
strike either the phosphor screen or emulsion of the film.
Second, an electron may pass near the nucleus of the atom and be attracted
by the positive charge. This will result in a change
of trajectory (scattering) but will not result in any loss or decrease
of energy (no change in wavelength). Such elastically (no loss
of energy) scattered electrons may contribute to the final image if
the change in trajectory is not so severe that they are
eliminated by the aperture of the objective lens. Third, a primary
beam electron may interact with one of the electrons in the
atom and lose energy to it (inelastic collision). This results not
only in a scattering of the electron but also a change in its
wavelength. When such an inelastic scattering event takes place with
one of the inner orbital electrons (K, L, or M shell) the
energy that is lost by the primary beam electron is very specific and
like a characteristic X-ray contains information about the
element that produced it. In a conventional TEM the scattered electrons
(both from elastic and inelastic collisions) serve to
degrade the final image as they strike the recording surface and cannot
be distinguished from nonscattered electrons. A small
diameter aperture eliminates many of these and increases our image
contrast but degrades our resolution by reducing the angle
of illumination.
[diagram here]
Electron Energy Loss Spectroscopy (EELS)
We can take advantage of those inelastic collisions that take place
with inner orbital electrons in one of two ways. The first
involves using a type of magnetic prism to separate those electrons
that are still traveling at their original velocity Eo from those
that have been inelastically scattered Ei = Eo - E. If one scans the
specimen in a point by point fashion (STEM) the electrons
that are produced at that point can be focused onto a magnetic prism.
This then further focuses the electrons to different focal
points depending on their energy. If an aperture or slit is placed
in this focal plane and positioned over the point in which the
electrons with an energy of Eo are focused, and then detected using
a scintillator and photomultiplier tube (PMT) similar to
what is used in an SEM, the relative quantity of electrons can be converted
into a bright or dark pixel on a CRT. By moving the
aperture so that it coincides with those electrons that have been slowed
by a specific amount one can create an image that is
indicative of where the elements are localized that would slow the
primary electrons by that amount. EELS detectors can be
put onto most commercial STEM and are placed beneath the film recording
camera.
Electron Spectroscopic Imaging (ESI)
Like EELS, ESI takes advantage of the fact that electrons can be slowed
by very specific amounts depending on which
elements (and which electrons) they interact with. One company, Zeiss,
has taken advantage of this and incorporated an
magneitc prism into the column of the TEM. By doing this they can also
separate the polychromatic (many wavelengths) beam
after it has passed through the specimen. The main advantage is that
by placing a discriminating aperture or slit above the 2nd
projector lens of the TEM they can create a typical TEM image that
can be recorded on film. With an ESI system it is not
necessary to scan the image and thus images that contain information
about the elemental composition can be created that have
higher resolution and require less time to create than either X- ray
maps or EELS. The limiting aperuture can thus create higher
contrast images by eliminating inelastically scattered electrons without
decreasing the angle of illumination and therefore
resolution. We can also increase the accelerating voltage to exactly
match that of the element we are seeking and thus create a
photographic image of that element's distribution in the specimen.
In both EELS and ESI it is essential that the specimen be
extremely thin for a primary electron that interacts with more than
one atom will no longer contain specific information about
that interaction.
A second method for examining the surface topology and structures of
specimens in a TEM employs shadowing techniques. In
this case the image contrast is produced by the uneven distribution
of fine metal particles. Once again electron dense metals are
the coatings of choice and platinum, chromium, palladium, uranium,
and gold are some of the more commonly used metals for
shadowing. Also, as the name implies information about the surface
topology is gained by creating a shadow effect which is
directly proportional to the microarchitecture of the specimen. This
is accomplished by depositing the coating metal from a low
angle (5 - 30 degrees) relative to the general plane of the specimen.
The greater the height of portions of the specimen the
larger will be the resultant shadow. The contrast difference created
by a shadow that is created is opposite to a shadow
produced by sunlight.
[diagram this]
In interpreting a shadowed preparation it is important to know the direction
from which metal was deposited. In fact if the angle
and direction of the shadowing source are known relative to the specimen
the height of the specimen can be calculated using
the equation:
H = tan O X l Where H = height of specimen
O = angle of shadowing
l = length of shadow
or H = b/c X l Where b = Height from level to source
c = Distance from sample to source
[fig L-2 Wischnitzzer]
Shadowing may be done from a fixed angle (static shadowing) or on a
rotating specimen (rotary shadowing). Rotary
shadowing allows one to resolve portions of the specimen that might
otherwise have been obscured by the shadow.
As with negative staining resolution in the TEM of shadowed specimens
is dependent on the grain size of the deposited metal.
Basically there are three methods of depositing thin metal films for
shadowing preparations these being a) heated electrodes, b)
electron beam gun (often called an e-gun or electron gun), and c) cathodic
etching.
A) Heated Electrodes - With heated electrode evaporation the material
to be deposited is heated by passing a large electrical
current through it while maintaining it under high vacuum conditions
(10-6 to 10-7 Torr). The material then begins to volatilize
(boil) and is evaporated in all directions into the vacuum chamber.
Some of the metal particles will strike the specimen and
create a shadow depending on the topography of the sample and the angle
of the incoming particles. The most common device
for accomplishing this is a vacuum evaporator and this is still the
most common means of depositing metal or carbon. The
higher the vacuum at the time of evaporation the finer will be the
grain size. For this reason liquid nitrogen is often added to the
system to act as a cryogenic pump immediately before shadowing.
B) Electron Beam Evaporation - This technique is similar to the heated
electrodes method only in this case electrons emitted
from a surrounding tungsten filament (which emits electrons due to
thermionic emission) strike the target and causes it to heat.
The fine particles are then emitted from the source and are free to
strike the specimen. Once again this type of deposition takes
place under high vacuum conditions in vacuum chamber. Because electrons
are the source of heat in these deposition devices
they are often referred to as electron guns or "e-guns" but should
not be confused with the electron gun assembly that is the
source of imaging electrons in a TEM.
C) Cathode Etching (Sputtering) - In Cathodic Etching ionized molecules
of an inert gas (usually high purity argon) are focused
and accelerated to bombard a cathode target. The target consists of
a thin foil of high purity heavy metal (gold or
gold/palladium). The gas ions displace molecules of metal from the
target which are then free to go toward the specimen
(sputter) and coat it. Because little or no heat is generated in the
process cathode etching is also known as a "cold" source
technique. Unless special equipment is used the size of the deposited
metal grains in sputtering are often quite large and
although may be suitable for SEM are not suitable for high resolution
TEM imaging.
Shadowing is used on many of the same types of samples and for many
of the same reasons as is negative staining. As with
negative staining only information about the surface of the specimen
is really obtained. One often goes through the trouble of
shadowing (as opposed to just negative staining) because of the added
resolution that can be obtained, especially with low
angle rotary shadowing. Shadow casts can be made of any stable dried
organic or inorganic molecule of organism that will not
change shape under high vacuum conditions. The shadow cast can be made
on an intermediate substrate such as a piece of
mica and then removed or directly on a Formvar or carbon film on a
grid which is then placed directly in the TEM. It is
common to deposit the electron dense metal from a predetermined angle
to create the shadow effect and then to evaporated
from directly above, a fine layer of carbon which does not add much
electron opacity but does provide strength to the shadow
cast, particularly in regions where no metal was deposited.
A modification of shadow technique is known as replication. In forming
a replica many of the same steps employed in creating
a shadow cast (metal and carbon deposition under vacuum on an intermediate
substrate) are used. The shadow cast is then
removed from the substrate by floating on water and the pieces placed
in a solution to remove the biological or mineral sample.
Strong acids (hydrochloric, chromic, hydrofluoric) or bases (sodium
hypochlorite) are used, sometimes in succession, to
dissolve away the original biological material and leave only the metal/carbon
cast or "replica" of the original specimen. This is
often extremely useful in that the original material may have been
electron dense enough to prevent visualization of the fine
shadow produced on the surface of the specimen. It is also important
in making a replica that there be sufficient carbon
deposited to make the replica strong enough so that it will hold up
in the TEM. The tiny floating replica fragments are rinsed in
water and picked up on naked 300 mesh grids and examined in the TEM.
Thus there is no support film present as there is in
shadow casts.
A modification of the replica technique is when a replica is made of
a frozen sample. This is known as freeze etching or freeze
fracture. We will discuss this technique when we cover cryobiology.
In some cases the sample may not lend itself to direct replication and
in this case a two step replica (negative replica, reverse
replica) may be made. This is done by first making a plastic replica
of the specimen by applying liquid plastic to the original
specimen. After the plastic hardens the specimen is then removed from
the either by peeling or dissolving. The first stage plastic
replica is then subjected to metal and carbon deposition as before
and the plastic removed from the second stage replica by
dissolving in an organic solvent. The metal/carbon replica is then
examined in the TEM. Cases in which one might make a two
stage replica include rare or large specimens that cannot be sacrificed
or specimens that must be used for a second purpose.
In terms of resolution shadow casting, especially low angle rotary shadowing,
can equal or exceed the resolution capable from
negative staining. Replication is really the only technique available
for examining the surface features of an electron dense
specimen in the TEM.
One alternative to standard chemical fixation is the use of low-temperature
methods otherwise known as cryopreservation. In
cryopreservation samples are rapidly frozen and then further processed
using a variety of techniques.
Essentially the same goals of standard fixation apply here namely to
arrest cellular processes rapidly and preserve the cell in as
near to the living state as possible. We have a lot of confidence that
this is the case with cryopreservation as it has been shown
that rapidly frozen cells can remain viable following warming. Cryopreservation
offers a number of advantages over
conventional fixation among these are:
1) Rapid arrest of cellular processes. One is not dependent on the speed
of penetration of the fixative. (milliseconds vs.
seconds)
2) Avoidance of artifacts induced by changes in osmolarity, pH, or chemical imbalance.
3) Because cellular constituents are not subjected to biochemical alterations
they remain in more of their natural configuration.
Labile components are retained and antigenicity is usually improved.
4) Cells can be examined without introduction of other possible artifacts caused by dehydration or embedding.
5) One can examine cellular domains that might otherwise be inaccessible
(e.g. IMPs) or from a view that is usually not
possible (e.g. 3-D view via deep etch).
There are however a number of disadvantages as well and among these are:
1) The need for specialized freezing and processing equipment (-80 freezer, cryoultramicrotome, freeze fracture device, etc.)
2) Freeze damage due to poor freezing rates.
3) Limited view of specimen and or difficulty in manipulating the frozen material.
Rapid Freezing:
The major obstacle to good cryopreservation is the introduction of artifacts
due to formation of ice crystals that disrupt the
cellular structure. The goal of rapid freezing is to prevent the formation
of ice crystals and preserve the aqueous component of
the cell in near to the vitreous state. Vitreous refers to glass or
glass like, and just as glass is really a supercooled liquid and not
a solid, water can also exist in this quasi-solid state. In general
this is very difficult to accomplish with biological samples and
usually we simply strive to keep ice crystal formation to a minimum
which is often defined as whether or not the crystals are
visible in the electron microscope. This cannot be accomplished by
simply putting the sample in the freezer.
Cryopreservation Cont'd
Perhaps the most important aspect of rapid freezing is the choice of
cryogen or freezing medium. A good cryogen should have
several properties.
1) Low freezing point - need to have a good thermal gradient between the sample and the cryogen.
2) High boiling point - must minimize the formation of a vapor barrier
near specimen due to latent heat of sample. The
formation of an insulating vapor barrier around the sample is known
as the leidenfrost or "bad frost" phenomenon and prevents
the cryogen from making direct contact with the surface of the sample.
This tends to slow the freezing rate and produce ice
crystals.
3) It should have a high heat capacity and thermal conductivity (latent
heat). In plain terms it should be able to absorb heat
without increasing its own temperature. Because of this low molecular
weight liquids such as N2 and He tend not to very good
cryogens.
Cryogen melting pt. boiling pt.
Freon 22 -160 -40.8
Freon 13 -181 -81.1
Freon 12 -155 -29.8
isopentane -160 27.85
propane -189 -42
nitrogen -209 -196
ethane -183 -88.6
helium -272 (1o K) -268.9
An alternative to liquid cryogens is the use of a nitrogen slurry or
slush. By lowering the pressure of liquid nitrogen it can be
induced to freeze and become a solid. When brought back to room pressure
the liquid and solid nitrogen exist side by side.
Just as a glass of ice and water remains at 4 degrees longer than does
a glass of pure 4 degree water, the nitrogen slush has a
higher latent heat and can thus absorb more heat from the sample before
boiling. This reduces the leidenfrost effect and
improves freezing rates.
The rate at which a specimen freezes is usually the determining factor
in the amount of ice crystal formation and subsequent
damage there is. Slow freezing rates such as 1 C/min results in significant
damgae. The extracellular water freezes first and pulls
out the water from the cell as the concentration gradient changes.
In general cells do not contain large amounts of unbound
water so the formation of very large ice crystals usually does not
happen but the specimen can become shrunken and distorted.
Rapid freezing is usually defined as a change in temperature in excess
of 10,000 C/sec. (vs. 1 C/min). One of the major
problems associated with rapid freezing is the total amount of heat
that must removed from the specimen. If internal heat from
the specimen continues to warm those portions that are cooling it will
prevent the water from undergoing a rapid phase change
and large ice crystals can form. For this reason the size of the specimen
should be kept to a minimum regardless of the freezing
method used and the specimen carrying device should be made of a small
amount of material that has excellent thermal
conductivity. Thin pieces of copper or gold are usually used.
Cryopreservation Cont'd
Specimens are then rapidly placed or "plunged" into the cryogen and
held there for 20 - 30 seconds. It is important that the
specimen be as small as possible as good freezing will only occur on
the outer surface and one wants to reduce the heat load
placed on the cryogen. Plunge freezing is best used on very small specimens
or cell suspensions.
One problem associated with plunge freezing is the fact that as the
cryogen removes heat from the specimen it begins to warm
up. This is a localized effect but results in either a decrease in
the thermal gradient between the cryogen and specimen or even
worse in the formation of leidenfrost. To avoid this it is desirable
to have a fresh supply of cryogen constantly moving over the
sample and taking away any excess heat with it. This can be done by
either moving the sample rapidly through the cryogen
(projectile freezing) or moving the cryogen past a stationary specimen.
This is the theory behind jet freezing. The most
commonly used cryogen for jet freezing is liquid propane and the device
is known as a propane jet freezer. Basically the unit
operates by putting the specimen on a very thin support foil or holder
and then placing it between two thin pipe with opposing
ports. Liquid propane (which was liquefied by a bath of liquid nitrogen)
is stored in a bomb underneath the output ports and is
then forced out from the ports under great pressure by introducing
dry nitrogen to the the propane bomb. Two opposing
streams of liquid propane the hit the specimen from both sides and
carry away the excess heat. Cooling rates of 30,000 C/sec
have been claimed for propane jet freezing and heat exchange is 2 -
30 times faster than with plunge freezing alone. These are
dangerous to use and we are experimenting now with a device I helped
to design which uses six ports (3 above, 3 below) that
uses a stream of liquid nitrogen.
A second alternative to rapid freezing samples with liquids is to bring
them in rapid contact with a very cold surface. Although
this will result in severe ice damage in the sample that is not immediately
in contact with the surface, it can produce excellent
results in the region immediately adjacent to the surface. Contact
freezing is accomplished by pre-cooling a large metal block
(usually polished copper, brass, or gold) and then rapidly bringing
the sample in contact with the block. Because latent heat and
leidenfrost is not a concern in this method one simply wants to create
the largest thermal gradient possible. For this reason
liquid nitrogen or even better liquid helium is used. The primary reason
that most researchers choose to use liquid nitrogen is
that it costs approximately 45 cents per liter whereas liquid helium
costs $200 per liter.
One problem with bringing the sample in contact with the block is the
possibility that it will bounce and thus damage the
specimen. For this reason a special freeze slamming device is used
that has a glycerol hydraulic damping system to drop the
specimen onto the block but prevent it from bouncing. A modification
of this procedure involves grabbing the specimen
between two precooled metal surfaces. These cryopliers are widely used
in cryopreservation of specimens such as muscle
fibers.
A modification of surface freezing is known as spray freezing. In spray
freezing the sample in the form of a suspension is spray
or atomized onto a precooled metal block or into a cryogen. This avoids
the problems of bouncing and keeps specimen size to
a minimum (1 ul or less volume). It has the disadvantage that the specimen
must be one that can be sprayed and is often difficult
to handle afterwards as it must be collected without rewarming the
sample.
Cryopreservation Cont'd
The latest in freezing devices is known as a high pressure freezer.
At extreme pressures of 2100 bar (Bar = 1 ATM = 760 mm
Hg) the nucleation of ice is significantly reduced. A second thing
that happens is that the melting point of water is lowered to -
22C (vs. 0 C at 1 ATM). This is one reason that cold water on the ocean
bottom does not freeze. At these pressures the
critical cooling rate is raised to 100 C/sec (vs. 10,000 C/sec at 1
ATM). The device works by initially pressurizing the
chamber with isopropanol followed by liquid nitrogen. Because the cells
are pressurized for only a few milliseconds before the
LN2 is introduced they are generally not harmed too much. LN2 can be
used because at these pressures it will not boil so no
leidenfrost is formed.
Device freezing depth cost
Plunge freezer 10 - 20 um $ 0.50 - 50
Spray Freezer 10 - 20 um $ 10 - 50
Slam Freezer 20 -40 um $ 2000
Propane Jet 40 um $ 10,000
High Pressure 50 - 100 um $ 150,000
Regardless of the freezing method used many specimens are treated with
a cryoprotectant to reduce the possibility of ice
damage. Cryoprotectants function by both increasing the number of ice
nuclei and retarding the growth of ice crystals. By either
binding to water molecules or substituting for water molecules cryoprotectants
reduce the number of water molecules available
for binding to growing ice nuclei and thus greatly slow the growth
of these crystals. Generally cryoprotectants are viscous and
in this way they also slow down the rate of diffusion of water from
the specimen as the exterior water freezes. This helps to
reduce the shrinkage effects of slow freezing. Some commonly used cryoprotectants
are glycerol (penetrating type) or sucrose
(non- penetrating type) and are generally used in concentrations of
10-30%. One of the disadvantages of cryoprotectants is
that it has been shown that extensive exposure to cryoprotection can
alter the internal structure by applying osmotic pressure to
the cytoplasm. Usually marine organisms have a number of dissolved
salts in the medium which act as cryoprotectants and
often these can be frozen without further cryoprotection.
One of the things that can be done with rapidly frozen samples is to
replace the aqueous component of the specimen with an
organic solvent without allowing the to change from its frozen arrested
state. During the freeze subsitution process a rapidly
frozen sample is held for one to two days in a vial of organic solvent
at -80 C. Over this time period the frozen water molecules
are replaced or "substituted" by molecules of the organic solvent.
This happens despite the fact that the water is never allowed
to return to the liquid state. Acetone is usually the solvent of choice
although ethanol and methanol have been used as well. The
organic solvents have some fixitive properties of their own which can
be enhanced by the addition of standard fixitives such as
osmium tetroxide. Recently anhydrous glutaraldehyde has become available
for use in organic solvents during freeze
substitution. Thus the cells are chemically cross linked and fixed
before their components have an opportunity to change from
their frozen positions. The samples are then gradually brought to room
temperature (done slowly to prevent renucleation of ice
crystals), the fixitive, if any, rinsed out with pure organic solvent,
and infiltrated and embedded as usual. Thus in freeze
substitution the fixation and dehydration steps are combined into a
single step.
One great advantage of rapid freezing and freeze substitution as oppposed
to standard chemical fixation is that many of the
artifacts associated with chemical fixation can be eliminated or greatly
reduced. A prime example of this is in the study of
membranes and membrane bound organelles. The length of time a fixitive
takes to penetrate a cell and the changes it induces in
terms of periability often results in shrinkage or wrinkling of membranes
and membrane bound organelles. If one compares
these to chemically prepared cells the smoothness and roundness of
freeze substituted material is quite surprising. Also rapid
cellular processes such as the fusion of membrane bound vesicles can
be captured because although the fusion process itself is
very rapid, the freezing rate is even faster.
A second great advantage of freeze substitution is seen when one uses
the fixation properties of the organic solvent alone to
preserve the cell. This has the great advantage of hlding all cellular
components in place while at the same time not cross linking
the cell so completely that not cytochemistry can be done. In fact
cells preserved in this way have better ultrastructural
preservation and greater ability to react in cytochemical treatments
than any other method. A variety of methacrylate resins
have been developed which facilitate immunocytochemical processing
of cells including Lowicryl which remains a liquid down
to - 40 C and can be polymerized at that temperature using U.V. light.
Thus cells are freeze substituted, infiltrated, and
polymerized without ever regaining the unfrozen state. Cell structures
and biochemicals can therefore be preserved in nearly
their native state.
A modification of the freeze substitution process is known as freeze
drying or in some cases as "cryodistillation." In freeze
drying the rapidly frozen specimen is held cold under vacuum and its
water is allowed to sublimate (go directly from solid to
gas). Once all the water has been removed a low termperature embedding
resin (Lowicryl) is introduced, allowed to infiltrate
under vaccum and eventually polymerized and sectioned. Cryodistillation
has the advantage that water soluable components
are not extracted from their nave position during the substitution
process and thus much can be learned about the natural
biochemical composition of the cell.
Yet another technique that can take advantage of rapidly frozen specimens
is cryosectioning or "cryoultramicrotomy." In
cryosectioning the specimen is sectioned while still in the frozen
state and before any post processing (substitution, distillation,
etc) has been done. Frozen sections are thin enough for examination
in the TEM and this can be done either on the cold
sections using a cryotransfer system which keeps the sections at liquid
nitrogen tempertures or on warmed specimens that have
been allowed to dry down onto a grid. Generally the ultrastructural
preservation of cryosectioned material is quite poor. The
primary reason for using cryosections is the enhanced antigenic reactions
that one can get from unfixed, unembedded material.
The major drawback (other than poor structural preservation) is that
cryosections are exceptionally difficult to make and the
technique and equipment needed are tough to master and expensive. Despite
this cryoultramicrotomy can allow one to
immunolocalize structures at the TEM level that would otherwise be
impossible to do with conventional methods.
At times it is important that one examine a replica of a specimen that
has not been dried but rather is in the hydrated state. For
these applications one would use the technique of freeze etching or
freeze fracture. The key element of freeze fracture is that
the platinum/carbon replica is made on a frozen specimen that is contained
within a vacuum evaporator. In those cases where
actual fracturing of the specimen is important a mechanical microtome
that can be cooled to liquid nitrogen temperatures and
operated within the vacuum evaporator is also employed. As could be
expected, these specialized vacuum evaporators or
Freeze-fracture devices, are quite expensive often costing as much
or more than the TEMs for which they prepare specimens.
Freeze fracture operates on the principle that a specimen that is held
in place frozen in ice can be treated like a solid rigid
structure and broken or fractured in various regions of the specimen.
These newly fractured surfaces may run along the original
surface of the specimen but are more likely to pass through the internal
portion of the specimen. Thus a replica made of these
newly exposed surfaces can reveal important information about the internal
composition of a specimen, not just the exterior as
in normal dry shadow casts or replicas. As with other cryotechniques
the size of the ice crystals formed is especially important
in freeze fracture and specimens are usually prepared using one of
the rapid freezing techniques previously discussed (plunge
freezing, jet freezing, slam freezing, high pressure freezing).
To prepare a freeze fracture replica a small amount of the sample is
placed on a small metal carrier sometimes referred to as a
"hat." These hats are often made of gold due to the ability of this
metal to conduct heat rapidly away from the specimen. The
hats are then rapidly frozen and stored in liquid nitrogen until ready
for use. In the mean time the freeze fracture device is
warmed up and brought down to high vacuum using a diffusion/mechanical
pump system. The cold specimen on hats are then
rapidly transferred to a stage which has been cooled under vacuum by
liquid nitrogen flowing through the stage. The chamber is
then rapidly pumped down again while the stage and specimens remain
at LN temperatures. Now the microtome arm assembly
with attached razor blade is cooled to -195 C with LN while the stage
and specimens are gradually raised to about -100 C.
The cooled knife is then rotated over the specimen until contact is
just made and thin shavings are removed from the top
surface. These shavings are not sections and the specimen is not so
much sectioned as it is scraped. Although a razor blade is
used the analogy is closest to a huge snow plow clearing a snow covered
dirt road. As it makes contact small pieces and
chunks are torn loose from the road revealing exposed frozen surfaces.
After a sample has been scraped and a clean surface
exposed the sample is often "etched" for a period of 1-3 minutes. During
this process the cold knife hovers above the fractured
specimen while both are held under vacuum. The combined effect of high
vacuum and a temperature differential (-150 vs.
-100) causes some of the frozen surface water of the specimen to sublimate
(go directly from water to gas) and be removed by
the vacuum system. As this happens the non-aqueous components of the
specimen become more an more prominent relative to
the flat background. A variation on this technique involves deep etching
followed by rotary shadowing. Using this technique
large relief images can be created of structures that are only visible
in the TEM.
A modification of this technique is known as double replica or complementary
replica formation. In this process the sample is
initially frozen sandwiched between two planchets which are then inserted
into a special precooled holder. This holder is then
flipped apart while on the cold stage and the specimen is split in
two exposing matching surfaces. A replica of each surface is
then made and examined. In this way both surfaces can be viewed whereas
the opposite surface is scraped away in
conventional fracturing.
One of the most useful and widespread applications of freeze fracture
is in the study of biological membranes and their various
protein components. To understand why we need to look at how a biological
membrane is organized. Basically all biological
membranes are composed of two layers of phospholipids arranged so that
their hydrophobic regions face one another.
Embedded in this phospholipid sandwich are intramembranous particles
(IMPs) which are proteins or protein complexes that
span from one hydrophilic side of the membrane to the other. In addition
to these IMPs there may or may not be additional
protein complexes that are embedded in one half or the other of the
membrane.
[Fig. 14-1]
When a cooled razor blade contacts a frozen specimen the membrane selectively
splits apart at the hydrophobic junction. This
occurs because at reduced temperatures the energy needed to split the
hydrophobic junction of the membrane is less than that
needed to split the ice or aqueous components of the cell. A replica
made of a fractured surface typically reveals large portions
of the internal region of biological membranes. In fact, freeze fracture
is about the only technique available that allows one to
visualize the hydrophobic regions of membranes. Of course other structures
such as nuclei, flagella, and cell walls are also
fractured during this process.
As difficult as it is to make a good freeze fracture replica, it is
often even more difficult to interpret one. Part of the reason for
this is made clear in looking at the following illustration. Conventional
scientific illustration usually places the light source in the
upper left hand corner of the image at an angle of about 45 degrees
relative to the specimen. Most SEMs follow this
convention when designing the scan pattern, detector position, and
display monitor. Based on this we conclude that an object is
convex when the dark shadow produced by light is in the lower right
hand corner of the image and concave when it falls in the
upper left hand corner. Because cells are mostly composed of spherical
vesicles and curved membranes, the freeze fracture
image is a case study in this type of illustration. The first problem
that one then encounters in interpreting freeze fractures is the
fact that lights and darks of shadows are reversed from those made
by light. For this reason some people initially find it easier
to interpret their micrographs from the photographic negative rather
than the positive image. A second problem is the fact that
when a replica is placed into the TEM there is virtually no way to
know before hand the angle of shadow (direction from which
metal was deposited). After cleaning, and picking up tiny replica fragments
on grids and then placing them into the TEM nearly
any orientation is possible. Two things can help to orient the viewer
of a freeze fracture replica. The first is any known structure
that the operator knows to be convex in nature. IMPs are an excellent
example of these. Using the shadow produced by the
convex structure the direction of shadow can be determined and the
micrograph oriented so that convex structures appear
convex and concave ones appear concave. A strategically convenient
piece of dirt that fell on the surface of the sample
immediately before the replica was made can also fill this function.
One problem that arose when freeze fractures began to be widely used
by electron microscopists was that of terminology.
Before freeze fracture a biological membrane could be thought of as
a single sheet with two (hydrophilic) surfaces. Now
suddenly scientists had four different surfaces to deal with and a
way was needed to clearly distinguish between them. A paper
by [?] created the guidelines by which all other freeze fracture images
would be labeled. The first rule that was suggested is that
the membrane be broken down into surface (hydrophilic) and fracture
(hydrophobic) profiles. These were abbreviated as the
"S" and "F" designations. The other way distinguishing which surface
is being discussed is to determine whether the half of the
membrane in question was in contact with the protoplasmic (P) portion
of the cell or the endoplasmic (E) portion. Thus any
given biological membrane can be spoken in terms of four surfaces or
"faces"; going from the outside of the cell towards the
cytoplasm the plasmamembrane would be designated as having a ES face,
a EF face, a PF face and a PS face. This
designation system becomes tricky when one begins talking about double
membrane bound systems (nuclear envelope,
mitochondrion, chloroplasts) but none the less is clear and unambiguous.
Double replica formation is especially useful in this
case for both the EF and PF faces of a given membrane can be viewed
and the relative abundance of IMPs on each can be
determined.
Immunoelectron microscopy as defined here has a broader definition than
strictly antibody-antigen reaction. Under the broad
definition it includes the labeling of biochemicals so that their localization
can be visualized in the TEM. In order to visualize this
in the TEM we must in some way tag or label the biochemical of interest
with an electron dense marker that distinguishes it
from other cellular components. Some techniques that come under this
category are lectin-horseradish peroxidase reaction,
biotin-avidin conjugates, as well as antibody-antigen reactions.
An immuno response is one in which an organism exposed to a foreign
body develops a resistance to that type of body so that
it is resistant or "immune" to infection from future exposure to a
similar type of body. Any substance capable of eliciting an
immune response is referred to as an antigen.
There are two broad classes of immune responses: 1) Humoral antibody
responses involve the production of a antibodies
which circulate in the bloodstream and bind specifically to the foreign
antigen that induced them and 2) Cell-mediated immune
responses which involve the production of specialized cells that react
mainly with foreign antigens on the surface of host cells. In
immunoelectron microscopy we are primarily concerned with humoral responses
that produce soluble antibodies.
Antibodies are produced by a class of cells known as B lymphocytes.
The only known function of B lymphocytes is in fact to
make antibodies. Antibodies are a unique group of proteins that can
exist in millions of different forms each with their own
unique binding site for antigen. Collectively they are call immunoglobulins
(abbrv. Ig). Most antibodies are bivalent, that is they
have two identical antigen binding sites. The antigen binding site
is composed of a heavy and a light chain each containing about
220 amino acids. They are hinged by way of their heavy chains to an
Fc (Fc stands for Fragment Crystallization).
[Figure 17-17 here]
There are five different classes of antibodies; IgA, IgD, IgE, IgG,
& IgM. They differ from one another in the composition of
their heavy chains. IgG antibodies constitute the major class of immunoglobulin
in the blood and are copiously produced during
secondary immune responses. It should be remembered that when using
monoclonal antibodies (single antigenic site vs.
Polyclonal = multiple antigenic sites on that antigen) the right portion
of the antigen must be presented to the surface of the
section in order for the antibody to recognize it and bind to it.
Immunogold labeling can be done in one of several ways. The colloidal
gold particles (5- 40 nm) are conjugated either directly
to the antibody being used or to an IgG or IgA protein. In an indirect
method the sections or tissue is first incubated in the
antibody of interest. Next the sample is exposed to a secondary antibody
that reacts to the IgG or IgA antibody of the first
animal. This secondary antibody is conjugated to a colloidal gold particle
which because of its electron density allows one to
visualize where in the cell the primary antibody (and by implication
the antigen) is localized.
One can even do double labeling experiments if gold particles of two
different sizes and different animal IgGs are used. This
requires using sections picked up on uncoated grids. A number of other
electron dense tags that can be used with antibody
labeling as well. Ferritin molecules (the storage protein for iron
in mammals) have a diameter of about 10 nm and there iron
component imparts their electron opacity. Horseradish peroxidase (HRP)
is an enzyme that can be coupled to primary
antibody and then allowed to form an electron dense reaction product
that is visualized. One alternative to using a secondary
antibody involves the use of protein A. Protein A is produced by the
bacterium Staphylococcus and can bind to the Fc portion
of IgG. Tagged protein A is often better suited for use as a secondary
label than is an anti IgG antibody.
There are a number of rules that one must follow in performing immunoelectron
microscopy. The first involves the choice of
grids. Some of the solutions that the sections will be exposed to may
react with the metal of the grid (e.g. copper react with
high salt conc. solutions). To avoid unwanted chemical reactions one
typically chooses grids made of non-reactive metals.
Nickel is a common choice as it is fairly unreactive and relatively
cheap. Others prefer solid gold grids as these are the most
inert. Coated or uncoated grids may be used but sections should not
be carbon coated after they are picked up as this can
make the sections hydrophobic.
A second rule that should be followed is to avoid overfixation. This
is often a difficult thing to balance as we want to retain as
much structural preservation as possible while at the same time retain
biological activity of molecules. These are mutually
incompatible goals. Excessive crosslinking with glutaraldehyde can
prevent the reactive sites of a molecule from retaining its
shape and therefore function and fixation with osmium can render membranes
impermeable and make membrane bound
biomolecules inaccessible. Sometimes osmium can be used as fixative
after the antibody labeling has been carried out, but this
can only be done in cases where the specimen is labeled prior to embedding.
A typical fixative for immunocytochemistry
studies would be a mixture of 4.0% paraformaldehyde and 0.1% glutaraldehyde
in the proper buffer. This will provide
reasonable ultrastructural preservation while preventing excessive
cross linking. Often sections on grids are initially soaked on a
drop of saturated sodium metaperiodate. This reacts with any unbound
or unreacted glutaraldehyde in the sections and
prevents the glutaraldehyde from crosslinking the antibodies when they
are applied to the sections. Freeze substituted
specimens must of course be rehydrated if pre-embedding labeling is
to be done otherwise this method is an excellent fixation
choice (assuming that fixatives have been left out of the substitution
fluid. Of course unfixed material such as found with
cryosectioning offers the best cross reactivity but structural preservation
and image contrast is often very poor. They offer the
advantage of never having been fixed, retaining water soluble components,
and having not embedding medium to penetrate.
Sometimes sections are "etched" to make the antigens contained within
it more accessible. A unique application of this involves
polystyrene embedding and acetone etching. Prolonged exposure can remove
all of the embedding resin leaving only the
specimen after sectioning. This is similar to xylene extraction of
paraffin sections.
Another type of immunolableing involves the use of Avidin. Avidins are
a class of basic glycoproteins that have a MW of about
65,000 and can be found in large amounts in egg white or Streptomyces.
They are useful in immuno EM because of their high
affinity binding for biotin. Each avidin molecule has four biotin-binding
sites per molecule. Many biomolecules can be labeled
with biotin (biotinylated) including proteins, lectins, fluorescent
beads, and nucleic acid bases. When one treats a sample with
gold or ferritin conjugated avidin it selectively binds to the biotinylated
molecule and the metal atoms acts an electron dense
marker of where the biomolecule of interest is localized.
Enzyme Cytochemistry: [text 254-261]
In addition to the anitbody/antigen type of reactions there are other
biochemical reactions that can be utilized to visualize the
localization of biological compounds in the TEM. One of these is the
very specific reactions that can take place between certain
enzymes and their substrates. The reactions can be utlilized to localize
the presence of a given enzyme in a specimen. The
technique works by trapping the resultant reaction product between
the enzyme and the substrate and visualizing it.
As with immunoEM the initial fixation of the specimen must be sufficient
to preserve structure while at the same time no
degrading the enzyme's ability to react with substrate. A fixation
similar to the ones used in immunoEM is often employed.
Because enzymatic reactions are sensitive to environmental conditions
such a pH, temperature, and substrate concentrations all
of these need to be taken into account. Finally, unlike gold particles
the reaction product may be only weakly electron opaque
therefore at least some of the sections are usually viewed prior to
post staining. One interesting note is that enzymatic labeling is
often best accomplished using epoxide resins rather than methacrylates.
It is believed that the hydrophilic nature of
methacrylates allows the enzyme to easily access the substrate, carry
out the reaction, and then detach. Since we want the
enzyme to remain attached to the substrate (thus showing the localization
of the substrate) it is actually better to use resins that
are more difficult to penetrate and therefore more difficult for the
enzyme to release from.
The reaction between horseradish peroxidase (HRP) which is an enzyme
that reacts with peroxide and through the addition of
DAB and oxidation with OsO4 forms and insoluable electron dense precipitate.
Sometimes HRP is coupled to an antibody and
then a reaction product formed through the addition of the proper components
to form an insoluable precipitate. The earliest
use of this involved ferritin- HRP complex but this may have a reduced
access to the lectin binding sites due to steric hindrance.
Recently HRP has been electrostatically bound to colloidal gold and
thus used as an indirect marker for lectin binding sites.
This avoids the steric hindrance problem and gives a better indication
of lectin binding site distribution.
Alternative Methods [280-285]
Lectins:
Lectins are plant compounds that have specific affinities for certain
carbohydrates. They may be tagged and used as a probe
for the presence of these oligosaccharides.
Naturally occurring compounds:
Molecules that normally bind or react with one another can be utilized
One final type of biochemical localization involves the use of Diaminobenzidine
(DAB). DAB specifically binds to sulfated
mucopolysaccharides when exposed to them at low pH. The DAB can subsequently
be oxidized by exposure to Osmium
tetroxide. The resulting electron dense precipitate is then an indication
of where the sulfated polysaccharides are localized. A
second rather use of DAB takes advantage of the fact that DAB can become
oxidised by U.V. irradiation. If a sample is first
made fluorescent by either labeling with a fluorescent dye or conjugated
molecule, then bathed in DAB and finally exposed to
the wavelength of light that will excite the fluorochrome, the energy
absorbed will oxidise the DAB which in turn will form an
insoluable, electron dense precipitate. This precipitate will therefore
be colocalized with the fluorescent marker. This reaction
also takes place with autofluorescent compounds that are naturally
found in cells therefore making the cytochromes of
mitochondria and the chlorophylls of chloroplasts sites where DAB precipitation
will take place. The technique has the
advantage of allowing fluorescent and EM studies to be done on the
same sample and is an excellent way of positively
identifying the biological structure that was originally labeled.
Sections have thickness to them and are not really flat. Things generally
bind only to exposed molecules. Size of probe and
porosity of the embedding medium are two factors that influence immunolabeling.
For this reason hydrophilic methacrylate
resins such as LR White and Lowicryl are often used in immunoelectron
and cytochemical microscopic studies and epoxy
resins generally avoided. This is not to say that epoxy resins cannot
be used, only that if labeling is poor the choice of resin
should be re-evaluated. Labeled sections are usually post stained after
immunolabeling with uranyl acetate or lead citrate to
provide contrast to the sample.
Basic Principle:
The process of photography is basically a series of chemical reactions.
A specific class of compounds known as silver halide
salts are light sensitive. Usually these salts consist of silver bromide
(although iodide and chloride are sometimes used). When
these salt grains are struck by a given number of photons the energy
of the photons is imparted to them and they undergo a
change to their activated state. In this activated state, these particular
silver halide grains can undergo a chemical process
known as development to become black silver grains. The unexposed silver
grains are dispersed through a gel matrix known as
an emulsion. This emulsion is then supported by either a clear backing
(acetate or glass plates) or on paper.
The activated silver halide grains are developed to black silver particles
by a reducing agent, the developer. Like all reducing
agents, developer is basic having a pH higher than seven. Because developer
will eventually reduce even those grains which are
not in a highly activated state or which have received very few photons,
the development process must be stopped. This is
accomplished by either using a stop bath which is usually a mild acid
solution or by putting in running water which has a low
enough pH to stop the development process. This step is known as the
stop process.
The remaining silver grains still have the potential of undergoing reduction
and becoming visible as black grains even after the
stop step. To prevent light from later developing these grains and
causing the image to darken with time, these undeveloped
grains must be removed in a process known as fixation. Photographic
fixatives are usually thiosulfate salts. These have the
ability to remove from the emulsion the unactivated silver halide grains
that do not come out in the developing or stopping steps.
Thus the photographic process is a series of 1) light activation, 2) development, and 3) fixation.
The two primary factors in choosing a photographic emulsion are light
sensitivity and grain size. The term grain size litterally
refers to the size of the exposed and developed silver particles in
the emulsion. These can range from 0.2 m to 20 m in size with
"fine grain" high-resolution films being at the smaller end of the
spectrum. Rememer that 0.2 m is equal to 200 nm and begins to
approach the resolution limit of a light microscope! This is an important
feature of a film in that it allows a negative to be
enlarged greatly before one begins to see the actual grains. The distribution
of these grains is also important with low speed
films having a uniform distribution of grains whereas high speed films
tend to have a wide distribution of different sized grains.
There are basically two types of emulsions which distinquished by their sensitivity to different energy sources.
Panchromatic emulsions are sensitive to all wavelengths of light and
for this reason must be handled in total darkness until the
fixation stage is complete.
Orthochromatic emulsions are sensitive to only certain wavelengths of
light and can usually be handled under a safelight. The
polycontrast paper that you are now all familiar with has a variety
of different sized silver grains in it emulsion. This allows the
activation of specific sized grains depending upon which filter wavelength
is used. The size of these grains and their dispersion
changes the exposure curve for the paper are what are responsible for
making a print of different contrasts.
Exposure:
In order to activate the silver grains of an emulsion it must be exposed
to an illumination source. Exposure is defined as the
darkening effect of light upon the silver halidses of the emulsion.
It is the product of intensity of illumination (I) times the length
of exposure in seconds (T).
E = I x T
This is the Reciprocity Law. Image density relates to the ability of
the image to impede the transmittance of light. However this
relationship does not follow a straight line equation for films and
each film has a characteristic curve which reflects its reaction
when exposed under a variety of conditions. This characteristic curve
has three portions the toe (underexposure), the straight
line portion (proper exposure) and the shoulder (over exposure). Each
film and developer combination produces its own
unique curve. The slope of the straight line portion of the curve is
known as gamma. This is important for gamma relates to the
ultimate contrast found in the emulsion.
A steep curve will yield an emulsion with high contrast whereas a low curve will yield one with lower contrast.
As a microscopist your final data, the material that you will present
to colleagues for peer review, are images. As such they
should be both scientifically informative and aesthetically pleasing.
Let's take a look at how they can be both.
Micrographs as Data:
As with scientific writing, scientific micrographs need to be brief,
informative, and well crafted. With the exception of review
articles, taxonomic treatises, and other similar publications, one
tries to use the fewest number of figures to communicate the
data. Perhaps the best example of this "brevity is everything" concept
can be found on the pages of Science. Micrographs in
this journal are known for being very small and very few. Unlike other
forms of data presentation (graphs, tables, charts, line
drawings, etc.) it is unusual for a single micrograph to contain a
great deal of information. In fact most micrographs contain
information about only a single feature or in the case of three dimensional
reconstruction, a single image may contain only a
small portion of the information that the author is trying to convey.
Most professional publications limit the authors to a certain number
of plates or in some cases, a certain number of printed
pages. When one considers how much written material can be presented
on a page of text, the need for image brevity becomes
apparent. Thus the first rule of image publication is to use as few
micrographs as possible to illustrate a given point and if a
single micrograph can be used to illustrate multiple points then it
should be given preference over others.
The second rule is to make the micrograph as small as is possible without
losing the data. More micrographs per plate
translates to more data per page. This is why it is important to not
fill the entire image with the specimen when one is using large
format negatives (TEM and Polaroid). One can always safely enlarge
a negative 2-3 times the original size but image reduction
is often very difficult to do*. A good way to evaluate if an image
is too small is to photocopy it on a standard, poor quality
photocopier. If the data within the micrograph is lost, it is probably
too small. Also be certain to check the "Instructions to
Authors" section for the particular journal that you intend to submit
the manuscript. Some will mention that image reduction is at
the publishers discretion while others will insist that the final plate
size be of a specific dimension to avoid further reduction. It is
a good idea to assemble the final plate so that it will fit within
the standard size of that particular journal without further
reduction and to specify this in your letter to the editor.
A third rule to bear in mind is that it is still VERY expensive to publish
in color. If one can convey the data in a black and white
micrograph then this should be done, even if it requires the use of
2-3 separate micrographs to convey the data contained in a
single color micrograph. This is NOT the case with presentation images
which will be discussed separately. Even when using
techniques such as 3-D confocal image a pair of stereo black and white
micrographs, or
* The same is true for image contrast which can usually be increased in the darkroom but rarely reduced.
even a single 2-D volume projection, can often convey the essential
information. Color micrographs should be taken as well as
black and white ones and for this reason many fluorescent microscopes
are equipped with two cameras, one loaded with color
slide film and one loaded with black and white print film.
The labels and captions that accompany your plates are almost as important
as the micrographs themselves. A well written
figure legend should allow the reader to understand the micrographs
without having to refer back to (or even have read) the
text of the manuscript. The same is true of figure labels which when
possible should be obvious to the reader and the same as
those used in the text. It is important to define the abbreviated labels
either in the body of the captions (once defined it should
not be redefined) or as a "Key to Figures" presented before Figure
1. Other types of labels (arrows, arrowheads, stars,
asterisks, etc.) should be defined each time they are used as one often
needs an arrow to illustrate different features in different
micrographs. Labels come in a variety of styles and sizes. It is important
to use the same style throughout the manuscript. Black
on white lettering is the most versatile but pure black or pure white
can also be used.
The final thing that should be included on each figure in a plate is
the scale bar. Some authors prefer to simply include "image
magnification" as part of the figure legend but this runs the risk
of misinterpretation should the figure be enlarged or reduced
from its original size. A scale bar, incorporated into the micrograph,
will remain useful regardless of how the image is magnified
as it will always stay proportional to the original. Scale bars have
the further advantage of brevity for if a similar magnification is
displayed on a single plate of figures one can simply state "Scale
bar = ?? for all figures."
The actual assembly of the plate (group of related micrographs on a
single page) is one of the most difficult steps in publishing
micrographs. A photocopier with enlarging and reduction functions can
be an extremely useful tool and can greatly aid your
plate production. It is always best to do a plate work-up using photocopied
images as these are cheap, easy to produce and
modify, and can be cut and arranged to create the "lay out" of the
plate. Many journals require that all the figures be abutting
whereas others allow you separate the individual images with black
or white tape.
Several methods of actually attaching the micrographs to the stiff board
can be used. Rubber cement can work but tends to
wrinkle the micrographs and can be messy. Dry mount is a heat sensitive
adhesive that lays flat and is very permanent. A
number of spray adhesives come in a variety of permanence levels and
are good for different purposes. We will demonstrate in
lab how these plates can be mounted, assembled, and trimmed.
While the first requirement of any micrograph is that it be scientifically
informative, a second requirement is that it be
aesthetically pleasing. This means that the contrast, overall brightness,
neatness of labeling, and general flow of the individual
micrographs that make up a plate should all go together. A good photographer's
rule of thumb is that one takes 8-10 pictures
for everyone published. The same ratio applies to scientific photography,
only the ratio may be quite a bit higher. Attention to
detail goes a long way towards getting reviewers and readers to take
you seriously. Micrographs with knife marks, poor
fixation, sloppy darkroom technique, etc. suggest that you are not
serious about your science or your data. If you are not
serious why should your colleagues take you seriously. When deciding
which journal you should submit you micrographs too,
consider how well that particular journal reproduces black and white
halftones. If you are not happy with the quality they give
to the work of authors, assume that you will not be happy with the
way your micrographs are reproduced. In today's world
there are too many journals and you should be able to choose at least
one that meets your high standards for micrographs.
Micrographs prepared for presentation are quite different from those
prepared for a manuscript. First of all color is not a major
obstacle and in fact with today's slide maker software etc, people
have come to expect color, even when one is dealing with
SEMs and TEMs where color has to be artificially added to the image.
In the case of fluorescent micrographs color is an
essential and when using double or triple labeled specimens it is a
necessity. Even in the case of images captured with a SIT
camera or confocal microscope people have come to expect color being
added to these otherwise black and white images.
When one is preparing images for a poster presentation size is as important
as it was when preparing a manuscript plate. In this
case the images must be large enough to be comfortably viewed from
a distance of four to five feet. If you cannot read the text
or see the data in the micrograph from this distance then things are
too small and you should work to enlarge it. With a poster
one can usually have a little more latitude with the number of figures
used but bear in mind that many poster sizes are quite
restricted and you may be very limited in the figures that you can
use. When giving an oral presentation it is usually better to err
on the side having too many figures because the eye quickly gets board
when it has no text to read. If the audience is only
listening to your words then having multiple images, even if they all
illustrate essentially the same thing, works to your
advantage. My personal record is 115 figures in a 15 minute talk but
30 to 40 is my average. Here is where aesthetics can
really come into play. be certain that when you see that "really gorgeous"
shot that you take it, even if there is nothing
scientifically important about the image. You will someday be glad
that you did.
In order to make an image more useful we often employ a type of image
processing. Basically there are three types of image
processing and these can be defined as Optical processing, Analog Processing,
and Digital Processing. We have all had
experience with optical processing. By using the glass lenses of an
enlarger to focus and magnify a negative we are practicing a
type of optical processing. We are changing the original data contained
in the image. Such things as burning and dodging a
negative during the exposure process and altering the brightness and
contrast by choosing different exposure conditions and
type of photographic paper can all be thought of as optical image processing.
This is the oldest form of image modification.
Analog processing requires that the image be manipulated through electronic
means. Most of us have also practiced this type of
image processing. The image on a television screen is controlled by
the voltage signal that the electronic gun at the back of the
CRT receives. By electronically altering this signal we alter the final
displayed image. Changing the amplitude of the signal
(difference between the highest and lowest point) will affect what
we refer to as the contrast (difference between black and
white). Altering the overall strength of the signal will influence
the brightness of the image. The important thing to note about
analog processing is that all of the components that go into making
the image are all altered.
Finally there is digital image processing. In digital processing the
image is represented by a series of picture elements or "pixels."
Each pixel has a discrete position in the image and a defined intensity
value. The pixel's position and intensity can be
represented by numerical values. With today's high speed computers
we can now manipulate each of these numerical values in
a number of ways. We will talk about some of these possible manipulations.
Image Capture for Image Processing:
Traditionally there were only two ways to share the data generated on
an electron microscope with other researchers. These
were to actually have the researcher looking into the same microscope
as you or to take a high quality photograph of the
sample and make publication quality prints from the negative. With
this negative a skilled microscopist could produce a high
quality print that emphasized that portion of the image that he or
she considered important. While this is still the primary mode
of data dissemination used by microscopists photography is fast becoming
an archaic practice. Today, image capture and
image processing are fast replacing photographic methods as a way to
share electron microscope images. A classic example of
this is the replacement of 8mm movie cameras by VCR camcorders in nearly
every American home. Video is cheaper, does
not require processing, reusable, captures images and sound and is
easier to view at home.
The reason that film can be replaced and the reasons for doing so lie
in advances that have been made in two fields. The first is
in the field of electronics. The second is in the field of computers
and computer software. Together, these two allow a
researcher to easily handle and manipulate images that five years ago
could only be done using very sophisticated and very
expensive hardware.
All of this is possible because of two things. First the human eye can
distinguish 256 different levels of grey. Second, every
image can be broken down into a series of small grey dots each of which
is defined by one of these 256 grey levels. This
process of turning a continuous tone image into one made up of pixels
is known as "digitizing" an image and the resultant image
is said to be digitized or "pixelated." This is essentially how black
and white photographs are reproduced in newspapers. Take
a close look at a newspaper photograph and you will see that it is
simply a series of black dots of various sizes (i.e. intensities).
A black and white photograph is essentially the same thing (a series
of black silver grain dots) the primary difference being the
size of the dots and spacings between them.
Twenty years ago no electronic device (TV monitor, Image printer, etc.)
could come close to the particle size and spacing of
photographic paper. These early attempts were crude and the large size
of the spots resulted in what is called a "grainy" image.
The resolution of the human eye is about 0.2 mm and so any two dots
that are farther apart than 0.2 mm can be seen as single
dots or "grains." The resolution of a digitized image is therefore
partially dependent on the number of pixels per unit area. The
higher number of points per unit area the greater the resolution. Since
the digitized image can be represented as a matrix of
pixels it's dimensions are given in terms of the number of pixels present.
A high resolution digitized is usually defined as having a
point density of 512 X 512 or greater. For reasons discussed later
(e.g. loss of data points due to post processing) it is always
desirable to collect the data in as high a resolution manner as possible.
Even if your output device is not of high enough
resolution to take full advantage of the data set a denser digitized
image gives you more more latitude.
Example: The same image can is represented by four digitized matrices
which differ in terms of their spatial resolution: 256 X
256, 128 X 128, 64 X 64, 32 X 32. The distance from the observer directly
affects how the spatial resolution of the image
influences how it is perceived. When viewed from a distance all four
of these images appear nearly identical but when seen up
close they are radically different.
[Fig. 3-4]
Image Processing
The eye's ability to detect grey levels is intimately linked to what
we call contrast. Contrast refers to the distribution of
brightness in an image. A high contrast image is composed primarily
of dark black and bright white and has a quality of intense
boldness to it. In contrast a low contrast image has only middle grey
tones present and appears washed out. An image with
good contrast should have all 256 grey levels represented somewhere
in the image reflecting the natural distribution all the way
from black to white. This is important not just from the standpoint
of aesthetics (i.e. creating a "pleasing" picture) but also in
terms of information. A picture that is too high in contrast will result
in a loss of image detail in those regions where there is a
subtle but important change in image brightness. Likewise, an image
that is too low in contrast may not reveal image detail
because the differences that the eye could normally detect are not
visible. Because a digital image can in fact be broken down
into at least 256 grey levels (in some cases even more) and each of
these can be manipulated separately we can "enhance" or
modify image contrast in very specific ways. Increasing the image contrast
would involve taking a digital image of limited grey
values and expanding the differences between them.
Example: A digital image is composed of pixels that range in grey value
from 100 to 160. A simple and quick calculation could
be made that first subdivides the group around the middle value of
128 (= 1/2 of 256). All pixels of a 128 remain unchanged.
Those of 127 are changed to 126 while those of 129 become 130. In the
next step all of the original pixels of intensity 126
become 124 while those originally of 130 become 132. The process continues
in this manner (new value = original value -(or
+) (N) where N = steps from 128). By this process the new image would
have a expanded grey scale that now ranges from 72
to 192. Still not perfect but much improved.
Likewise, reducing the contrast might involve artificially changing
the grey level value between pixels to spread out the tone
range.
Example: A single line of a digital image has the pixel values 0, 0,
0, 120, 120, 255, 255, 255, 255. If we were to plot these
values on a Brightness/Position curve it would look like this.
We could alter the value of these pixels to the following string:
0, 40, 80, 120, 140, 180, 240, 255, 255.
While this may give us a more pleasing final image it is important to
remember that we have essentially "created" data for these
pixel points. It is always best to collect the original data image
in as near to the "perfect" contrast balance as possible, but since
this is often difficult to do it is better to err on the side of slightly
too little contrast than too much. It is easier to artificially add a
little contrast than it is to subtract contrast. This is true not only
for digital processing but for optical processing as well. There is
always a danger when collecting a contrasty image that important information
(represented by subtle changes in grey level) will
be lost.
Ex: Contrast stretch 170 X 1.5 = 255 so if the highest value in an image
is 170, simply multiply each pixel value by 1.5 to
"stretch" it out to the full grey range.
One of the main problems in any digital image capture system is noise.
Generally this noise is the result of electronic interference
or spurious signal that is produced by the detection system or subsequent
amplification of signal. This is the same kind of noise
that is realized on an inexpensive stereo tuner when it is played at
full volume as noise generally becomes a greater problem as
one turns up the amplification of any electronic signal. In an SEM
this noise can result from increasing the voltage on a
photomultiplier tube (PMT) or the subsequent signal amplifier. As one
is always trying to maximize the signal to noise (S/N)
ratio it would be nice if there were some method of removing any noise
that was introduced to the image by way of the signal
detecting system.
There are several ways by which a digital image can be processed to
remove some of the noise. The first is a "filtering"
approach whereby we apply a mathematical algorithm to the digitized
data set and remove any spurious pixels. A spurious
pixel is defined as a pixel whose value exceeds, by some predetermined
value, the value of any of it's immediate neighbors.
Thus if we look at the following data matrix for a 3 X 3 cluster of
pixels the computer can easily determine if the value of the
central pixel is "appropriate".
127 130 129
126 248 131
128 131 133
Recognizing the "248" value as being inappropriate and therefore of
likely spurious origin it could take the average of the
surrounding 8 pixels (1036/8 = 129.5) and assign a rounded value of
130 creating a new pixel matrix of:
127 130 129
126 130 131
128 131 133
Thus eliminating the stray pixel. It must be remembered however that
this is a new data set and the original image will be lost. It
is possible that the original 248 value was correct and one must be
careful in applying this type of smoothing or noise reduction
system.
A second approach to noise reduction is known as image averaging. In
image averaging the same image is collected multiple
times and the values of each pixel are averaged to create a new value:
Example: Collect the same 3 X 3 matrix and display the averaged image.
125 179 142 127 133 140 126 156 141
130 133 128 130 137 126 130 135 127
134 136 138 130 134 139 132 135 139
Image 1 Image 2 Average of 1 & 2
Notice that the spurious value, 179, is recognized and reduced regardless
of where it happens to lie in the matrix. If one
collects the image a third time and averages it against the average
of #1 and #2 the image is further refined.
126 130 139 126 156 141 126 143 140
132 133 129 130 135 127 131 134 128
132 137 137 132 135 139 132 136 138
Image 3 Average 1 & 2 New average image
You can see that after multiple passes the image will be "cleaned" up
with each subsequent collection and averaging of the
image. What is more is that even if a spurious signal occurs in one
of the later image collections (there is an equal probability
with each collection) the more sophisticated image averaging algorithms
will account for this and minimize the impact of a
spurious signal.
In the SEM we try to minimize the S/N ratio by collecting our final
image in a slower scan speed but sometimes this can
degrade the quality of the image, especially if it is charging badly.
Although image averaging is usually employed with multiple
fast image scanning (e.g. TV rate) is can sometimes be used in image
acquisition on the SEM.
Edge Enhancement:
Another example of
Thus image resolution is dependent not only the number of pixels per
unit area but also the different brightness intensities that
can be represented by each pixel. The different intensity levels are
represented as a binary data string or bit. In the simplest
model each pixel is either black (0) or white (1). This would then
be a 1 bit (21) representation. A two bit image (22) could
represent each pixel as 00, 01, 10, or 11. Thus four shades of grey
are possible. Thus one require 8 bits (28 = 256) of data
per pixel to represent it as one of the 256 possible grey values. Many
of today's sophisticated image processing computers and
video games deal with color images (of which the human eye can distinguish
thousands of different hues). For that reason it is
not uncommon for these computers to have 16 (65,536 colors) or even
24 bit (16,777,216 colors) capability per pixel.
Printers for Digital Images
Dr. Alan D. Brooker JEOL (UK) Ltd., Welwyn Garden City, UK
Inkjet printers
Inkprinters are now very affordable, and can offer very good printing
resolution. A quantity of ink is ejected as a droplet (by
thermal and/or electrostatic means) and fired at the paper to form
a dot. In principle any form of paper can be used, but results
are clearest if special low-absorbency paper is used. If transparency
film is used a short drying period must be allowed or the
ink will smear. Virtually all inkjet printers can print in color or
monochrome - though they print much faster in monochrome. For
inkjet printers the grey scale of color range is limited, and the resolution
is relatively low (but improving rapidly), the overall
image density can be low, and images take a long time to print (up
to 20 minutes), but eh printers are very cheap to buy and
run.
Thermal printers
There are two types of printer to consider - thermal wax (relatively
cheap), and dye-sublimation (expensive). Thermal wax
printers work on a dot-matrix principle (halftoning) to produce grays
and colors - a heated element transfers dye from a carrier
onto the paper (or whatever) in dots. The process is relatively slow
(more than 5 minutes) but produces excellent density even
though non-primary colors are produced by dithering. The resolutions
currently available are comparable to inkjet printers.
Dye sublimation printers represent stat of the art as far as photo-realistic images are concerned.
A single heating element sublimes dye from a carrier film onto a specially-prepared
paper into which the ink diffuses. The
amount of dye transferred from the film depends on the heat applied
to the element - this therefore determines the grey or
color-level. The diffusion process results in continuous tones (like
conventional photographs) on the paper. Dye-sublimation
printers are expensive to buy and run, but can generate very high quality
color or monochrome images in reasonable times -
less than 5 minutes. Even though the resolutions may not sound impressive,
it is important to remember that for dye-sublimation
printers dpi equals pixels per inch.
Laserprinters
A laser is focused onto a drum which behaves such that where the laser
impinges becomes charged. The charged drum then
picks up magnetic toner particles which are subsequently deposited
onto paper, and sealed by heated rollers. In principle the
magnetic susceptibility of the drum and the focus of the laser determine
resolution, but in practice the toner coarseness and
delivery are more important. The vast majority of laserprinters are
monochrome, although color laserprinters are now becoming
available.
Laserprinters are cheap to buy, and cheap to run, the latest models
boast 600-1200 dpi. The images produced by such
printers are not photographic quality, but are easily recognized and
show good grey scale reproduction. It is well worth paying
the extra for laserprinter paper.
Conclusion
While the above is by no means an exhaustive summary of the current
marketplace, it is hoped that some of the more pertinent
areas have been highlighted. So what is the most suitable printer to
buy? My personal prejudice is as follows:
For low-resolution, reduced color (and grey scale) images (e.g. X-ray
maps, SPM images, Auger maps, etc.), destined for lab
notebook copies, giveaways, or internal reports - an inkjet printer
is a good compromise.
For grey scale images from anything destined for lab notebook copies,
giveaways, and internal reports - a Laserprinter would
suit most users.
For grey scale or color images, destined for top-copies of reports,
publication, or exhibition - a dye-sublimation printer will
give the required photo-realistic quality.
**************************************************************
A properly processed digitized image is still of little value unless
one can share it with others. For this reason the final output
device is of critical importance. One obvious way of sharing a digitized
image is to send interested parties the actual image data
set. Provided that the receiver has the appropriate computer hardware
and software they too can view the same image. This is
not as wild as it seems. Sales of Nintendo cartridges and computers
attest to the lengths people are willing to go to exchange
images. The use of data transmission over telephone and computer dedicated
wire systems (Bitnet, Internet, etc.) will make the
distribution of image files more and more common in the future. Already
journals such as Cell Motility and Cytoskeleton accept
manuscripts (including micrographs) on disk and in video format. When
one views a micrograph in a high quality scientific
journal today one is not seeing the same image that the author of the
paper did. First, the author recorded the image on
photographic film. Ideally this was using a camera on the microscope
and represents the best possible primary image. Next, the
author makes a high quality photographic print using optical image
processing techniques. Some researchers are better at this
than are others. Next, the publisher of the journal takes a paste up
of the figures and photographs the whole plate on a large
format internegative. Finally, this internegative is used in the printing
of the figures onto the pages of the journal. This represents
an image that is four generations removed from the original image.
If the digitized data set were distributed all interested parties
could look at a first generation image.
For the time being however photographic prints and other forms of hard
copy will be a necessary part of image processing.
Some of the options available today are:
Film Chain: This is essentially a high resolution CRT that is dedicated
to image capture. A photographic camera of some sort
(Polaroid, 35 mm, large format sheet film, etc.) is permanently attached
to the CRT. The camera may or may not contain lens
which focuses the image from the CRT onto the film. It is important
that the resolution of the CRT be high enough to take
maximal advantage of the digitized image. The photographic system on
the SEM is a film chain as is the small image capture
device on the confocal microscope. To capture high resolution color
images one can use a high resolution black and white CRT
in the film chain and break down the image into its red, green, and
blue components (RGB). A three color filter wheel then
rotates in front of the CRT while the color film is being exposed and
the composite image then results in high resolution
micrograph that has good color balance.
Thermal Printers: Thermal printers use a special paper and thermal transfers
to produce an image onto paper. The printer takes
the incoming video or digital signal and by heating the paper from
behind transfers a tiny dot of black (or color) onto the paper
which corresponds to a pixel. One of the things one looks for then
in a video printer is the number of dots per inch (DPI). The
greater this number the more points of information per unit area, the
greater the resolution. Most of todays B&W thermal
printers have a rating of approximately 300 DPI. Color printers use
colored transfers of yellow, magenta, and cyan. The
number of different colors available depends on the combination of
these. A 2 X 2 matrix can produce nearly 1000 different
colors whereas a 4 X 4 matrix can produce 4,096 colors. The more colors
however the bigger the dot matrix required and the
lower the image resolution. A good color printer may have a DPI rating
of only 186. They range in price from $4000 to
$22,000.
Plain Paper Printer:
Today's laser printers can achieve surprising quality and are increasing
used as output devices for black and white graphics.
Because they can produce much smaller dots than can thermal printers
there are now on the market plain paper laser printers
that can produce continuous tone B&W images with a DPI of 1200!
At 1200 DPI this equals one dot every 0.2117 mm. This
is near to the resolution of the unaided human eye. A second reason
for choosing one of these printers is the fact that plain
paper is significantly cheaper to use than is specialty thermal paper.
Its ability to withstand long term archival is also superior to
thermal papers which last only a few years under ideal conditions.
They are not necessarily cheaper however with good laser
printers starting at about $14,000. They also are incapable of producing
color images.
In order to perform good image processing on a digitized image something
has to be known about the composition of the
image. This quantification of the data stored in an image falls under
the general title of Image Analysis. One of the most useful
tools in image analysis is the image histogram. A histogram is basically
a graphic representation of the data contained in the
image data file. A simple example of an image histogram would be a
plot of how many pixels fall into a certain grey level
categories. We could represent three different images with the following
histograms.
{FIGS 4-1 to 4-3}
Using this information we can often subdivide portions of the image
that have similar brightness intensities together. Since similar
objects or objects of similar composition will have nearly the same
grey levels when viewed under identical conditions we can
make use of this numerical information to gather quantitative data
about the sample.
Example: Identical strains of bacteria are grown on two different test
media. When viewed in the microscope it is apparent the
cells grow better on medium A than on medium B. The researcher would
however like to quantify this so she collects 30
random images of each preparation all taken under the same conditions
(magnification, staining, light intensity, etc.). Using the
histograms generated for each image she identifies a subset of grey
levels that go into making up the images of the bacteria (e.g.
from 175 to 225). She now goes back and uses a subroutine to calculate
the percentage of pixels from each image that fall
within this range and ignores all others. Using this information she
learns that 17% of the area has these intensity values in
sample A whereas 42.5% of the area in sample B falls within these boundaries.
Thus growth of bacteria on medium B is 2.5
times that of medium A.
Another use of image histograms would be to define the grey levels that
correspond to the edges of the structure of interest.
Using sophisticated sub-routines one could then define the boundaries
and fill in that portion of the image that was contained by
the boundaries. One could then recalculate a new histogram for the
processed image and produce quantitative data about the
sample regardless of the values of the original grey levels. Other
sophisticated software can analyze the image and recognize
shapes defined by the user. This can be useful in cases of pattern
detection that might be difficult to see otherwise or in
separating out objects of interest from objects that have a similar
grey level intensity. These are just some of the ways that
information about the brightness intensity of each pixel can be used
to analyze the image.
In addition to the simple image processing mentioned before (contrast
stretch, regional highlighting, etc.) many other image
manipulations are possible using digital image processing. Some of
these involve changing pixel position and would include such
things as image rotation, image inversion, digital magnification, etc.
Another way in which pixel location can be used in
processing involves the merging or combining of two or more separate
images. This can be useful in reconstructing an image
that was previously sub-sampled (e.g. serial sections) or two views
from different collections (e.g. double labeling, 3-D
projections, etc.).
Differences in brightness intensities can also be used in a number of
different ways. Subtle shifts in brightness can be
accentuated to bring out detail of boundaries. This type of edge enhancement
can be very useful in clearly showing slight
changes. Likewise stray electronic noise or spurious pixels can be
removed by doing a next nearest neighbor algorithm or by
collecting multiple copies of the same image and averaging each new
image against the previous ones. One could also produce
an negative image by flipping all of the brightness intensities around
a middle value of 128.
Example: An image has significant noise introduced by the electronics
of image capture system. These appear as single white
pixels randomly distributed throughout the image. If the operator uses
a sub-routine that checks each pixel's intensity against its
neighbor's and if the difference between them is greater than some
value (say 50 grey levels) it will change the brightness value
of that pixel to a grey value that is an average of its nearest neighbors
and thus remove the spots. A second way would be to
collect the same image several times (6-10) and only save those pixels
brightness values that remain nearly the same in all the
separate images. This too will help in eliminating electronic noise
from the image.
In addition to being used to remove electronic noise from the image
one can perform image processing that will increase or
accentuate the differences between adjacent pixels to "enhance" the
boundary between the two pixels. This is often referred to
as "Edge Enhancement." It can be calculated using the Laplacian operation
for a 3X3 pixel matrix:
a b c
d e f
g h i
and the equation L = [e - (a+b+c+d+f+g+h+i)/8]
e will be replaced by L only if the the L value is greater than the
critical value or "threshold" otherwise the new value for e will
be ignored.
A final way in which brightness intensities can be processed is by assigning
various colors to the image based on the grey level
intensity. Often this results in a loss of resolution (it takes more
pixels to make a color than a grey value) but can have benefits.
One of the benefits is to make the micrograph pretty enough that it
will be published on the cover of Nature or in one of the
popular journals. A more useful application is to accentuate certain
structures in an image so that attention can be drawn to
objects of interest without drastically affecting the remainder of
the image. Another useful application would be once again for
use in merged images when one wants to still be able to distinguish
between the two images (double labeled, 3-D projections,
etc.)
One of the problems with digital image analysis is the tremendous amount
of computer memory storage that is
required. In a normal image processor 8 bits (= 1 Byte) are required
for each pixel. Since there are
262,144 pixels in a 512 X 512 image this means that 262,144 Bytes are
required to store one image. Most display monitors
are not squares but rectangles and an image format of 740 X 512 (378,880
Bytes) is more typical. A computer with a 10
MByte hard drive could hold only 26 of these images before its storage
capacity was exceeded. The data file that contains the
raw image is essentially the microscopists negative and it must be
preserved. For this reason mass storage capacity and some
form of data compression is an essential part of image processing.
Although computers now can be routinely outfitted with
large capacity hard drives (1 gigaByte = 1000 MBytes or more) even
these will reach full capacity in a relatively short time.
For this reason other high capacity storage media are used. Some of
these removable hard disk drives (Winchester, or
Bernoulli boxes), optical disk drives (WORM = Write Once Read Many;
Re- writeable), tape drives (1/4" cassettes), etc.
Each of these has their own advantages and drawbacks and decisions
are usually made on the basis of such considerations as
cost, convenience, accessibility, and capacity.
One type of image compression uses a technique known as run-length coding.
A simple example would be to scan a single line
of an image. There may be many pixels in a single line that have the
same grey value (e.g. in a good fluorescence image a large
portion of the pixels may be black). Rather than code this string of
pixels each as a separate 8 bit point we could code the
whole string with just two 8 bit numbers, one to represent the grey
level and the second to represent how many in a row have
that grey level. Essentially any string longer than 3 pixels in a row
would produce some savings and long strings could be
substantial. Of course if the pixel intensity changes between every
pixel this would double our storage as we would dedicate
two 8 bit numbers per pixel instead of just one.
Image Processing
A second type of compression uses Differential Pulse Code Modulation
or DCPM. This algorithm assumes that although the
pixel intensity levels will be changing as we move across the line
the changes between adjacent pixels will not be great. Thus
rather than code the absolute grey level using 8 bits per pixel we
can record only the change that occurred between one pixel
and its neighbor. If this change is small (e.g. 8 grey levels or less)
than we only need 3 bits to record it not 8. If we apply this to
the whole image we can achieve a savings of nearly 63% (= {8-3}/8 =
5/8). Even if we allow for a greater change in pixel
intensity (e.g. 32 grey levels vs. 8) DPCM could save us nearly 38%.
One final advantage of digital image files over photographic film is
the fact that they can be replicated with perfect fidelity many,
many times. Even in the best scientific journal or publication the
image that the reader sees is at a minimum a fourth generation
image (1= original negative, 2= original print for plate, 3= publishers
plate negative, 4= publisher's printed page). As the
standardization (or flexibility) of computer hardware and software
becomes more universal and more and more researchers
become linked by way of their computers and computer networks the rapid
dissemination of image files will become routine.
Even today it is not uncommon for researchers and product engineers
to swap image files either by wire or through the
exchange of floppy disks (which can hold up to five or more compressed
images depending on the format). In the future
readers and authors will be able to independently examine the same
image and the reader may even be able to perform their
own processing and analysis to either confirm or refute the author's
conclusions. Even if this does not immediately occur, the
first logical step would be to distribute the original image files
to the outside reviewers for their evaluation and even perform
their own processing if warranted. The fact that electronic backups
of valuable image data files means that even if a catastrophe
occurs the data can remain safely stored away somewhere else. The same
can never be done with photographic negatives.
Digital image processing is fast replacing optical and analog image
processing and will soon become the primary means by
which microscopists share images.
In addition to storing images as easily copied first generation images
these data files can rapidly be distributed to interested
researchers around the world via data transmission over telephone and
computer networks, and copies on disks and tapes and
physically distributed. In the future most publications will be distributed
in such electronic media virtually putting many
publishers out of business or at least changing the way they do business.
Libraries will become central electronic media
processing centers where researchers will access data bases and journals
remotely via their office computers.Image Processing
In order to make an image more useful we often employ a type of image
processing. Basically there are three types of image
processing and these can be defined as Optical processing, Analog Processing,
and Digital Processing. We have all had
experience with optical processing. By using the glass lenses of an
enlarger to focus and magnify a negative we are practicing a
type of optical processing. We are changing the original data contained
in the image. Such things as burning and dodging a
negative during the exposure process and altering the brightness and
contrast by choosing different exposure conditions and
type of photographic paper can all be thought of as optical image processing.
This is the oldest form of image modification.
Analog processing requires that the image be manipulated through electronic
means. Most of us have also practiced this type of
image processing. The image on a television screen is controlled by
the voltage signal that the electronic gun at the back of the
CRT receives. By electronically altering this signal we alter the final
displayed image. Changing the amplitude of the signal
(difference between the highest and lowest point) will affect what
we refer to as the contrast (difference between black and
white). Altering the overall strength of the signal will influence
the brightness of the image. The important thing to note about
analog processing is that all of the components that go into making
the image are all altered.
Finally there is digital image processing. In digital processing the
image is represented by a series of picture elements or "pixels."
Each pixel has a discrete position in the image and a defined intensity
value. The pixel's position and intensity can be
represented by numerical values. With today's high speed computers
we can now manipulate each of these numerical values in
a number of ways. We will talk about some of these possible manipulations.
In recent years a new type of microscopy has become popular that utilizes
the principle of a raster or "scanning" pattern to
image a specimen. These microscopes are called scanning confocal microscopes
or SCMs. The term confocal means "having
the same focus" and in practice this refers to two lenses aligned so
as to be focused at an identical point in space. There are
two radically different designs presently employed to achieve this
phenomenon of confocality. They differ principally in the
manner in which the raster pattern is established on the sample. Each
type of design has advantages and disadvantages over the
other and we will discuss each separately.
Like the SEM the confocal microscope differs from standard microscopes
in that it does not function as an optical microscope
but rather as a probe forming/signal detecting instrument. Thus despite
the fact that it relies on conventional light optic and glass
lenses to function it is as different from the standard U.V. microscope
it is mounted on as the SEM is from a TEM. Like the
SEM the confocal microscope builds its image in a point by point manner
based on the signal strength reaching the detector
from the specimen. To do this one must create a point scanning or raster
pattern on the specimen.
As the name implies a confocal microscope acts by scanning its illumination
source over the specimen and some mechanism
must be created for establishing this raster pattern. There are only
two ways to achieve this. One must either physically deflect
the illumination source to create a raster pattern or leave the beam
stationary and move the sample in a raster pattern. Some of
the earliest confocals used the scanning specimen technique in which
a very small sample was place on the end of a
piezo-electric device which could then be rapidly shifted by controlling
the current going to the piezo-electric. Optically this is
the most stable design for a confocal microscope but has the severe
limitation that only very small and stable specimens can be
examined. Certainly no living or wet specimens would work nor would
any large or heavy specimens.
The primary difference between a confocal microscope and a conventional
wide field microscope is the increased logitudinal
resolution of the confocal microscope. Longitudinal resolution is defined
as the abilty to resolve objects in the optical axis or
"Z-plane". This is in contrast with the transverse resolution which
is the ability to resolve in the "X-Y plane" of a flat field. Each
can be defined by the following equations:
Transverse resolution = 0.6 / NA
Longitudinal resolution = 2 / NA2
Where = wavelength of illumination and NA = numerical aperture of the
objective lens. Thus as the NA increases, resolution in
both the longitudinal and transverse dimensions increases. LR and TR
can be compared at a given wavelength by reducing the
equations to:
LR/TR = 4/ 1.22 NA
But even under the best conditions (NA = 1.4) the LR will be about twice the TR.
Confocal imaging greatly improves our ability to resolve in the LR, almost to the point where it equals the TR.
As the name implies the scanning aperture disk type of confocal microscope
uses a perforated disk ("Nipkow" disk) to
establish the raster pattern on the sample. Spinning at high speed,
the disk is made up of a series of tiny holes that are arranged
in a very precise pattern. In one type of scanning aperture disk confocal
microscope known as the tandem scanning
microscope, the illumination enters from above the aperture disk and
proceeds towards the sample. As it does this the beam
becomes highly attenuated and is imaged as a very small, nearly diffraction
limited point in the focal plane. Passing through a
beam splitter the illumination source focused by the objective lens
of the microscope and brought to focus in a single focal
plane. The reflected light from this single point is then reflected
back through the objective lens, split by the beam splitter, and
reflected back through a corresponding aperture in the aperture disk.
Thus only light that is brought to focus at the single point
in that single focal plane is capable of being reflected back to the
viewer. All other extraneous signal is eliminated from the
image. the perforations in the aperture disk therefore act as both
point source apertures and point detector apertures.
[Diagram]
The aperture disk is constantly spinning and as light passes through
the individual apertures a raster pattern is established on the
sample. Each of these points lie in the same focal plane thus the image
acquired represents all those reflected points that lie in a
single image plane. By imaging a series of individual focal planes
a "through focus series" or "optical sectioning series" can be
produced. The focal plane is determined by the fixed strength of the
glass objective lens. Thus the only way to change the focal
plane is to change the distance between the sample and the objective
lens and this is achieved either with a conventional stage
adjuster or more precisely by a piezo-electric stage height controller.
Because the aperture disk rotates at high speed a near
real time image of the sample is produced. A modification of this design
involves a shifting slit aperture rather than a point
apertures. This system sacrifices resolution for increased illumination.
The second type of scanning confocal microscope achieves essentially
the same result by a very different method. In a laser
scanning confocal microscope the illumination source passes through
a beam splitter and is moved in a raster pattern by a set of
pivoting mirrors. These mirrors cause the beam to move in a an X and
Y pattern similar to what occurs in the SEM. The beam
then passes through a tube lens and then an objective lens which focuses
it on the specimen. This produces a diffraction limited
light spot which achieves its minimum size only in one plane of the
specimen. Traveling via the objective lens, tube lens, and
scanners the reflected light is directed to a beam splitter. Another
lens then focuses the reflected beam so that only illumination
from the one point in the focal plane is brought to focus at a point
corresponding to the pinhole diaphragm or aperture. The
transmitted light is then detected by a photomultiplier tube and the
resulting signal is represented by points on a CRT.
[Diagram]
A second type of laser scanning confocal microscope known as the Odyssey
SCM has recently been developed by NORAN
Inc. Rather than deflecting the laser illumination by mechanical mirrors
the Odyssey uses an acousto-optic deflector (AOD). An
AOD is a glass body which by way of multiple transducers can establish
sound waves in the glass. These closely spaced waves
can perform much like physical grooves in a diffraction grating and
by changing the frequency of the signal driving the AOD
transducers the beam can be deflected in the x-axis at very high speed.
Coupled with a mirror deflector in the y-axis, the AOD
deflected laser can scan a sample (512 X 480) at 30 frames per second
(7 times faster than a dual mirror system). Also by
varying the amplitude of the AOD transducers the intensity of the laser
can be continuously varied. Another difference between
the AOD and other laser SCMs is that it employs a variable final slit
rather than an aperture. Thus true confocality is only
achieved in one axis rather than two.
All types of scanning confocal microscopes can thus acquire images from
a single focal plane. By eliminating extraneous signal
from above and below the an image is produced that is significantly
improved in contrast and resolution over conventional light
microscopes. Furthermore, because images can be acquired as single
planes these can be individually stored, manipulated and
recombined to produce three dimensional representations of serial optical
sections. Ignoring factors such as cost, the different
systems have pluses and minuses. The spinning aperture disk microscopes
can image in near real time, use a variety of primary
excitation illuminations ranging from white light to true U.V. While
these features are clearly advantageous the aperture systems
suffer from a great deal of loss of signal (due to passing through
two small apertures) and an inability to change the raster
pattern which is fixed by the distribution of holes in the aperture
disk. In contrast the laser scanning microscopes can alter the
scan pattern and thereby magnify the image by upwards of a factor of
eight. Also, because the signal is attenuated by only one
aperture and can be enhanced by the very sensitive photomultiplier
tube reflected light that is very low in strength can still be
seen on a laser confocal microscope. One major drawback is that because
the raster pattern is established by physically
moving mirrors, real time imaging is impossible. The best that can
be achieved now is four seconds per frame. Also because a
laser must be used as the primary beam source true colors, and true
U.V. fluorochromes can not be visualized on a laser
scanning microscope. Another disadvantage is that the operator can
only see the resultant image as it is presented on the CRT
whereas with the aperture disk systems the operator can either look
through the eyepiece or record using a TV camera and
CRT.
A new approach to using digital image processing involves collecting
a series of conventional images from a microscope that
are separated in Z space in much the same way as are the images on
a scanning confocal microscope. These images are
captured using a CCD (Charge coupled device) chip which in conjunction
with a digitizing card can store the image in digital
format. By applying a complex set of algorithms known as deconvolution.
In very simple terms what deconvolution or "digital
Confocal" does is compare each pixel in Image Processing
each image with the same pixel in the planes above and beneath it. Based
on changes in gray level intensity the program either
retains or rejects the pixel from the image plane being examined. In
this way only those pixels that were much brighter than they
were in the planes above and beneath and were therefore collected "in
focus" are retained and the image is restored as a
cleaned up, in focus image with all of the out of focus noise removed.
This cleaned stack of digital images can then be
manipulated for three dimensional projections and volume renderings
in the exact same ways scanning confocal images are.
Although the micrographs produced by the SEM appear to be in three dimensions,
this is actually not the case but is simply a
result of the great depth of field offered by the SEM. However one
can take advantage of this depth of field to produce true
stereo micrographs using the SEM. In order to achieve a stereo view
the same object must be viewed from slightly different
angles. A one eyed person, single lens camera, or single micrograph
cannot produce this effect. When the same object is
viewed separately by each eye, the regions of overlap can be fused
by the brain and great spatial information can be gained. In
doing this in the SEM certain factors must be taken into account.
Stereo micrographs are produced in the SEM by taking two separate micrographs
of the same object from different angles.
Depending on the magnification a tilt difference of between 5 degrees
and 10 degrees is usually ideal. This can be
accomplished in one of two ways. First, the specimen can be physically
tilted, recentered, and refocussed at an angle different
from the previous micrograph. Care should be take to refocus the image
using specimen height since large changes in the
strength of the final lens current will produce a micrograph of a different
magnification and of a slightly different rotation.
Alternatively, some SEMs offer the capability of angling the incident
beam. This has the advantage of not having to translate the
specimen but also it is possible to rapidly switch back and forth between
incident beam angles. Using this technique real-time
stereo imaging can be done in the TV. mode.
In order to create a stereo pair from two separate micrographs certain
rules must be followed. First, the images must be
aligned in the proper fashion, otherwise regions that are actually
peaks will appear as valleys. To do this a certain convention is
followed in which the lower or less tilted micrograph is viewed by
the left eye while the more tilted micrograph (positive tilt) is
viewed by the left eye. Next the images should be arranged so that
the tilt axis runs parallel to the interocular plane. Finally, the
center to center spacing of each object must be carefully positioned
so that the two images can be easily fused. This is
accomplished either by looking at each micrograph separately with each
eye (a difficult trick to master) or by using a stereo
viewer or glasses. Total magnification is also a concern here as for
if the micrographs are too big the center to center spacing
will be so large that the images cannot be fused together.
Although the separate images are the result of different shadows created
by interaction of the beam with the specimen we
cannot create a stereo view simply by having two detectors separated
by a few degrees. The beam must actually strike the
specimen from different angles. This is done either by tilting the
specimen or, as Leica does it, by shifting the beam slightly to
strike the specimen from slightly different angles.
The conventional way of displaying a three dimensional image is as a
side by side stereo pair. This can be done for either color
or black and white images. A second method is to display each black
and white image as either a blue or red image. This is
known as an anaglyph projection. When viewed with red and blue filtered
glasses each eye sees only one image and a stereo
view is formed. Alternatively the images can be projected through polarized
filters and then viewed with polarized glasses. This
requires two projectors each with a polarized projection lens, careful
alignment of the images, and a special "lenticular" screen.
Lateral resolution in the SEM is dependent on four limiting factors:
a) the size of the primary beam probe at the specimen surface.
b) the amplitude (stability of the energy spread) of the primary beam current (which affects the signal to noise ratio).
c) the penetration depth of the primary beam and the size of the region of signal production.
d) the effect of charging on the specimen.
Each of these is affected by different variables which we can discuss.
While any SEM can be run in a low (5 KeV or less)
mode, only a field emission gun (FEG) SEM has a probe size and stability
that truly allows us to take full advantage of low
voltage imaging.
Size of Primary Beam Probe:
The final size of the beam probe striking the specimen is dependent
on a number of factors. First, the size of the region from
which the primary beam electrons are generated is of critical importance.
On a bent tungsten wire filament this region is
approximately 106 , for a LaB6 emitter it is 105 , and for a Field
Emission source is 102 . Thus the electron source for a FEG
is 4 orders of magnitude smaller than for that of a standard tungsten
emitter. All of the lenses that are in the column of the SEM
and further focus this spot (condenser, final lens) and so with very
similar lenses the ultimate size of the probe hitting the
specimen will always be much smaller in an FEG SEM.
Amplitude of Beam:
Changes in amplitude of the beam, or energy spread, can be very detrimental.
Not only do such changes manifest themselves in
increased chromatic aberration in each of the condensing lenses of
the column but they can also result in differential signal
production (since this is dependent on how many primary beam electrons
strike the specimen). Once again an FEG-SEM has a
significant advantage over conventional SEMs in that they typically
have an energy spread of 0.2-0.3 eV whereas tungsten and
Lab6 emitters range from 1-4 eV. This may not sound like a lot when
one considers accelerating voltages of 15 to 20 KeV but
it is an order of magnitude difference which when amplified by chromatic
aberration can become significant. The issue of beam
stability becomes even more important when one considers low KeV beams
of less than 1 KeV.
Depth Penetration of Beam:
Just as the size of the region of primary excitation is proportional
to the size of the beam probe it is also dependent on the depth
to which the primary beam penetrates into the specimen. The lower the
KeV the better but often in order to efficiently collect
most of the electrons being produced by the emitter one must use an
anode/cathode difference of 10 KeV or more. One way
around this is to decelerate the primary beam electrons before they
reach the specimen. This can be done either up in the gun
assembly or closer to the specimen. Most SEMs can only do this in the
region of the anode/cathode and thus trade away a lot
of primary beam electrons and at the same time introduce the potential
for chromatic aberration.
The effects of increasing beam penetration can be seen on thin, low
atomic weight specimens. In the example below the
cuticular hairs of an insect are easily penetrated by the beam at relatively
low KeV (10 KeV) and signal is produced from a
greater volume of the specimen resulting in dramatically decreased
resolution of the surface of the specimen.
Charging:
Charging effects can be minimized by coating the specimen or by reducing
the total number of electrons needed to generate a
useful signal. Because a FEG SEM crams so many electrons into such
a small probe one can generate a comparable signal
without having to oversaturate the specimen with electrons. This coupled
with the reduced energy of the beam results in less
specimen damage and reduced charging.
In-Lens Detector:
The ability to image a specimen in the SEM is often limited not so much
by the specimen or the signal it produces but the ability
of the detector to collect this signal. This becomes a critical issue
at very short working distances (5 nm or less) which are
necessary for very high resolution work. A secondary electron detector
positioned to the side of the specimen is sometimes
blocked from receiving signal by the specimen and stage itself. This
is similar to the situation with a specimen that has a deep
cavity from which signal cannot escape despite the fact that it is
producing a significant amount of signal.
One attempt to overcome this limitation in signal collection is to place
a secondary electron detector within the final lens of the
SEM. In this way the detector is on nearly the same optical axis as
the primary beam itself thus the position of the detector
relative to the source of the signal is not the limiting factor in
signal detection. Because the secondary electron detector does not
need to be positioned between the specimen and the final lens very
short working distances can be used and very high
resolution obtained. The secondary electrons of the signal can be distinguished
from the electrons of the primary beam by both
their significantly lower energy and their directional vector (i.e.
opposite in direction to those of the primary beam. The
secondary electrons produced by the specimen do not interfere with
the primary beam electrons, the situation being analogous
to shooting a water pistol into the air during a driving rainstorm.
The chances of water droplets in from the water pistol actually
hitting the individual raindrops is vanishingly small despite the greater
numbers and significantly higher energy of the rainstorm.
Like the electrons of the primary beam, the secondary signal electrons
are focused by the electromagnetic field of the final lens
and concentrated into a smaller area. A converging lens works the same
way regardless of the direction from which the
electrons enter the lens. Thus the final lens acts somewhat like a
signal collector, concentrating the secondary electrons before
detection by the in-lens detector.
Intermediate And High Voltage EM [text 360-367]
Theoretical resolution in a transmission optical instrument can never
exceed 1/2 the wavelength of the illumination. de Broglie's
equation for calculating the wavelength of an excited electron is
= h/mv
Where = wavelength
h = Planck's constant (6.626 X 10-23 ergs/sec)
m = mass of electron
v = velocity of electron
By plugging in known values this equation can be reduced to
= (1.23/ V )nm Where V = Accelerating Voltage
In theory then the higher the accelerating voltage, the shorter the
wavelength, the greater the resolution capability! If 100 kV is
good 1000 kV (one million volts or 1 MV) is better! This type of accelerating
voltage is known as High Voltage Electron
Microscopy or HVEM.
A second advantage of HVEM is the ability of the beam to penetrate a
specimen. Even a 125 kV TEM cannot penetrate a
very thick specimen and most of our knowledge of the three dimensional
nature of biological structures is from reconstructions
made of serial thin sections each of which was laboriously sectioned,
photographed, and pieced back together. Electrons that
are accelerated to only 125 kV are more widely scattered than those
of a 1 MV TEM. Because of this thicker specimens can
be used than are possible with a conventional TEM. By viewing a greater
portion of the specimen at a single time stereo pair
images can be formed of a single thick section and viewed to gain three
dimensional information about the specimen. This is
best done on specimens that contain no embedding resin which would
only serve to scatter the electrons.
A final advantage of HVEM is that because fewer of the beam electrons
interact with the specimen (they are moving by too
fast) specimen damage tends to be less in a HVEM than in a conventional
TEM (assuming the same thickness sections). Of
course, since one of the primary reasons for using a HVEM is to look
at thick sections, this advantage is often canceled out by
the increased number of interactions with the specimen.
One major drawback to HVEM is $. Although the optical systems are essentially
the same as those found on conventional
TEMs the components associated with the 1 MV accelerating system usually
means that HVEMs are several stories tall and
require a special building dedicated to their use. There are only a
handful of active HVEMs in the U.S. and less than 30 world
wide. Most of these were built in the 1950's or 1960's. In an effort
to gain some of the advantages without all of the expense a
number of TEM manufacturers have introduced Intermediate Voltage Electron
Microscopes (IVEMs). Although generally
costing more than a conventional TEM, IVEMs can be housed in the same
places as conventional TEMs and can also be
operated at lower (80 - 100 kV) accelerating voltages. IVEMs and HVEMs
are most popular among materials scientists use
images of lattice images are often only possible at very high accelerating
voltages.
One of the major discoveries of cellular structure was found through
the use of HVEM, this being the complex microtrabecular
lattice that is believed to extend throughout the cytoplasm of cells.
There are many however who believe that this lattice is an
artifact of dehydration and specimens are properly critical point dried
no such lattice exists!
Typically resolution in an optical system is limited by the wavelength
of the illumination source. Due to the properties of
diffraction one can only image objects that are greater than 1/2 wavelength
of illumination. However, if one passes the light
through an aperture that is markedly smaller than the wavelength of
the illumination then based on the light transmitted or
reflected by the sample one can detect (i.e. image) objects smaller
than 1/2 the wavelength. A new type of light microscope,
the scanning near-field microscope takes advantage of this property
by passing light through a pinhole and bouncing it off a
very flat object that lies just beneath the opening. By moving either
the specimen or the aperture in a raster pattern and
recording the amount of signal that is produced an image of the object
can be produced.
[diagram]
This has been taken to the ultimate extreme in the case of scanning
probe microscopes (SPM). In a SPM the aperture is
replaced by an extremely fine probe or tip. Often this is a crystal
of tungsten that has been electroetched down to a very fine
tip, in some cases only an atom or two across. In the case of a scanning
tunneling microscope (STM) the tip is brought very,
very close to the surface of the sample and a small voltage is applied
to it. Electrons from the specimen then move or "tunnel"
across this gap and create a small current. If the tip moves even a
tiny distance closer or further away from the atoms this
tunneling current changes dramatically. The STM works by establishing
a constant tunneling current and then moving the tip
across the surface of the specimen in a raster pattern while keeping
the current constant. The only way to do this is to move the
tip up and down relative to the specimen and thus keep the distance
between the tip and the specimen constant. This up and
down movement of the tip is then recorded by a computer and the X,Y,
& Z coordinates can be graphically displayed as a
topographic map or image of the specimen surface.
[diagram]
The precise X, Y, and Z movements of the probe are controlled by piezoelectric
controls which are devices that can move a
very small and precise amount depending on the amount of current that
is passed through it. Today's piezoelectric devices are
sensitive enough to record changes at the atomic level and thus a STM
can create topographic images at the atomic level. One
problem associated with an STM is the fact that the sample must be
relatively flat and also conductive (otherwise tunneling will
not occur). As this is not terribly useful for most biological specimens
a second different type of SPM has been developed. The
Atomic Force Microscope (AIM) uses the same type of basic tip movement
and position recording system as does a STM
(e.g. X & Y piezoelectric controller, computer position recorder
and topographic display, etc.). It differs primarily in the type
of tip or probe that is used. In an AIM the tip is mounted on a spring
and is literally dragged across the surface of the specimen
in much the same way as is a stylus on a record. As the tip interacts
with the atoms in the surface it is repelled by atomic forces
(hence the name) and is deflected up or down. These up and down movements
are recorded by measuring either the tunneling
effect change between the top and bottom of the spring or by optical
deflection of a laser light bouncing off of the tip.
[diagram]
Sums allow us to use scanning technology to image objects at the atomic
level. Depending on the type of detector tip used wet
and or non- conductive biological specimens can be examined. New probe
designs (e.g. ion probes, etc.) are allowing us to
use this basic technology to create three dimensional maps of a wide
variety of specimens at the atomic level without being
limited by the boundaries of standard light and lens based optics.