Region of Primary Excitation: [Text pages 49-53]

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.

Beam-Specimen Interactions

[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.

Secondary Electrons:

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

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]

Transmitted Electrons:

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.

Bragg Equation:

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

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.

Cryopreservation

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

Cryoprotectants:

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.

Freeze Drying & Distillation:

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.

Cryosectioning:

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.

Freeze Fracture

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

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.

Photography

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.

Micrographs as Data

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.

Captions and Labels:

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."

Plate Construction:

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.

Micrographs as Art:

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.

Presentation 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.

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.

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

Contrast:

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.

Noise Reduction:

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.

Output Devices:

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.

Image Analysis:

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.

Image Processing:

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.)

Image Storage:

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%.

Digital Images as Negatives:

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.

Scanning Confocal Microscopy

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.

Scanning Aperture Disk:

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.

Laser Scanning Microscope:

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.

Digital Confocal Microscopy:

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.

Stereo Pairs:

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.

Low Voltage SEM

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!

Scanning Probe Microscopes

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.