Manual of Electron Microscope

Electron Microscopy: A Handbook of Techniques for the Biologist Preface Acknowledgments INTRODUCTION TO ELECTRON MICROSCOPY UNIT 1 - PREPARATION OF BIOLOGICAL SAMPLES FOR TEM Chapter 1 - Chemical Fixation Chapter 2 - Ultrathin Sectioning & Ultramicrotomy Chapter 3 - Post-Staining Chapter 4 - Grids & Grid Supports UNIT 2 - PREPARATION OF BIOLOGICAL SAMPLES FOR SEM Chapter 5 - Hard Tissue Preparation Chapter 6 - Soft Tissue Preparation Chapter 7 - Alternative SEM Specimen Preparation UNIT 3 - BLACK & WHITE PHOTOGRAPHIC PRINCIPLES... Chapter 8 - Film and Paper Composition Chapter 9 - Processing (Films and Papers) Chapter 10 - Negative Handling and Exposure (TEM & SEM) Chapter 11 - Enlargement Printing

Electron Microscopy:

A Handbook of Techniques for the Biologist

Stephen J. Beck Nassau Community College

Electron Microscopy: A Handbook of Techniques for the Biologist

Electron Microscopy: A Handbook of Techniques for the Biologist

Stephen J. Beck Nassau Community College

CONTENTS

TABLE OF CONTENTS Preface ○ ○ ○ ○ ○ ○ ○ ○ Acknowledgments

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INTRODUCTION TO ELECTRON MICROSCOPY

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UNIT 1 - PREPARATION OF BIOLOGICAL SAMPLES FOR TEM Chapter 1 - Chemical Fixation ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ Additives Fixation Times & Temperatures Preparation of Fixatives Methods of Fixation “Routine” Biological Soft Tissue Protocol Tissue Processing Note Embedding Media Fixation Schedule Fixation Schedule Worksheet Final Chemical Fixation Considerations

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Chapter 2 - Ultrathin Sectioning & Ultramicrotomy ○ Processing of Embedded Blocks - Block Trimming Glass Knife Making Diamond Knives Ultramicrotomy MT-2B Ultramicrotome Sectioning Procedure Troubleshooting Guide to Ultramicrotomy Materials Required for Ultrathin Sectioning Chapter 3 - Post-Staining ○ Uranyl acetate Lead Citrate Post-staining Procedure

vii ix 1 6 6 10 10 11 15 16 19 20 23 24 25

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26 26 29 33 34 39 44 46

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48 48 49 49

Chapter 4 - Grids & Grid Supports Grids Grid Supports Formvar Films Carbon Coating

v

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52 52 52 53 56

CONTENTS

UNIT 2 - PREPARATION OF BIOLOGICAL SAMPLES FOR SEM Chapter 5 - Hard Tissue Preparation ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○

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58 59

Chapter 6 - Soft Tissue Preparation ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ Critical Point Dryer Operation Alternatives to Critical Point Drying Fluorocarbon Drying Organo-Silicon Compounds Conductive Coating Vacuum Evaporation Sputter Coater Denton Desk II Operation Comparison of TEM and SEM Soft Tissue Protocols Fixation Schedule Fixation Schedule Worksheet

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60 63 67 67 67 68 69 69 71 73 74 75

Chapter 7 - Alternative SEM Specimen Preparation Uncoated Specimens Cryofracture Technique Microbial Specimen Preparation Microbial Fixation Schedule Worksheet

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76 76 77 79 83

UNIT 3 - BLACK & WHITE PHOTOGRAPHIC PRINCIPLES... ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ Chapter 8 - Film and Paper Composition ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ Emulsion Film Speed (ISO/ASA) Supports (Base) Routine Photographic Films and Papers Used For TEM and SEM Chapter 9 - Processing (Films and Papers) Development Stop Bath Fixer Washing Drying

vi

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84 85 86 86 87 88

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89 90 91 92 93 94

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95 95 96 98 99

Chapter 10 - Negative Handling and Exposure (TEM & SEM) TEM - Hitachi HS-8 HS-8 Camera System and Film Exposure SEM - Hitachi S-2400 S-2400 Camera System and Film Exposure Chapter 11 - Enlargement Printing Photographic Paper Grades Process of Enlargement Printing Enlargement Printing Variables Printing Tricks

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102 102 103 104 105

PREFACE The development of electron optics has had a profound effect on the study of life; its structure and function. Without the high technology tools known as the transmission (TEM) and scanning (SEM) electron microscopes, we would understand little about the world of the cell. As proof, I offer any typical college level introductory and advanced biology course, the textbooks these courses utilize, and any number of biological periodicals/journals. Biology professors will lecture on a variety of cellular structures and their related functions. Textbooks at all levels are crammed with TEM and SEM photomicrographs in an effort to illustrate any number of rudimentary concepts. Even to this day, cutting edge research relies on high resolution images as can be seen in many biological publications. In today’s high technology environment, the tools of scientific inquiry are easily overlooked and often taken for granted. Of course, one must understand that these instruments are tools; a means to an end. It will require a scientist to make sense of the images presented, someone who understands how the images were produced, even to the point of anticipation of a given result. The novice in electron optics must first learn technique and the theory behind it. The purpose of this handbook is to provide a detailed explanation and procedural guide to the many tedious procedures of biological electron microscopy, TEM and SEM. It can be used in the laboratory as a step by step guide and outside of the laboratory as the student attempts to comprehend the many concepts of biological electron optics. This handbook is intended for introductory college level courses in TEM and SEM. Depending on the college, this could mean the undergraduate (two and four year institutions) or even the graduate level. At Nassau Community College, the specific relevant courses are Transmission Electron Microscopy (BIO 221) and Scanning Electron Microscopy (BIO 222). In my ten years of teaching these courses at NCC, I have had the pleasure to introduce this valuable discipline to students with varied backgrounds and experiences, from traditional two year college students to those with earned Ph.D.’s. Regardless of prior education, students begin the EM courses at essentially the same level. This handbook was created to assist any individual in the attainment EM skills so that they will be able to utilize these important tools as they strive to answer the questions of life processes. The book is divided into three main units; Unit 1 covers the many topics of TEM biological sample preparation and is an excellent starting point in your understanding of biological EM. Many later topics in the book refer back to concepts introduced in this unit. Unit 2 involves the preparation of biological samples for the SEM. Even if you are only taking a course in SEM, much of the chemistry of chemical fixation is found in Unit 1, therefore, the SEM student is urged to refer back to the pertinent points. At NCC the ideal course sequence begins with the TEM course and is followed by the SEM course, which is logic I have followed in organizing this handbook. The final Unit 3 covers the concepts of black and white photography relative to both TEM and SEM. Here you will find general concepts of silver based photography followed by specific treatments of both TEM and SEM photomicrography and image capture using the Hitachi HS-8 TEM and the Hitachi S-2400 SEM, both of which are easily related to other common instruments in use today. As you progress in your education, I expect that these skills you will acquire and the discipline that it takes to master them will serve you well, whether you actually use electron microscopy in the future or not. S.J.B. August, 1996 vii

ACKNOWLEDGMENTS I would like to thank a number of individuals who have helped to make this book possible. Firstly, my wife and lifelong partner Mary and my children, Jonathan, Bradley and Jessica, for allowing me the precious time to write the manuscript. To my parents, Jack and Mary for providing me with the opportunity for an education and serving as such positive role models. In addition, I acknowledge the influence of my mentors, Dr. Kenneth Erb and Dr. Gary W. Grimes, both of Hofstra University. As my graduate advisor, Ken Erb guided my development as a scientist capable of conducting original research. I must confess that most of the information in this manual is taken from the electron microscopy experiences provided by Gary Grimes, a true expert and innovator in the field of electron microscopy. Finally, I would like to thank any of the faculty of Nassau Community College who have supported my endeavors over the past ten years, especially Dr. Dudley Chin, who as biology department chair, always encouraged even my failed attempts. His vision has initiated the technological revolution in the department as we approach the turn of the century. In addition, I thank Dr. Baruch May and Dr. Patricia Cassin, who as co-authors of two successful NSF grants, have brought an awareness of technological innovation to the college and our students. I also acknowledge the 1993/94 NCC Sabbatical Committee members who approved the sabbatical which made it possible for me to write the bulk of this manuscript. I finally acknowledge the support and foresight of the NCC administration, President Sean Fanelli and Vice President Jack Ostling for their continuing support of electron optics and other high technology endeavors at the college.

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INTRODUCTION TO ELECTRON MICROSCOPY

1

Introduction to Electron Microscopy When Max Knoll and Ernst Ruska were designing the original Transmission Electron Microscope (TEM) in Germany in the late 1920’s, they envisioned no biological applications for their instrument, provided it would even function as theoretically conceived. Today, the impact of electron optics, Transmission (TEM), Scanning (SEM), and a variety of others, is apparent. Our ability to directly resolve the microanatomy of the cell using these instruments has revolutionized our very understanding of life and its requisite processes. We take it for granted when a classroom instructor or a textbook describes the cristae of a mitochondrion, the 9+2 ciliary microtubule arrangement, ribosomal subunits and the phospholipid bilayer of a unit membrane. Where many cellular processes have been elucidated using a biochemical “grind & spin” approach (cell homogenization and cell fractionation via ultracentrifugation) followed by characterization of bio-molecules by isolation and purification using techniques such as gel electrophoresis, the cellular biochemist will often finally desire an image to support their chemical findings. The old maxim applies here, “a picture is worth a thousand words”. Even though the operating environment of the EM is a high vacuum (10-5 Torr) and biological samples must be preserved or fixed for examination, we have a unique insight to the molecular architecture responsible for life sustaining functions through the high resolving power of these instruments. Since structure begets function, and the EM can produces images of fixed cellular ultrastructure, it follows that we can come to a better understanding of cellular physiology through such images/electron micrographs. Of course, it becomes vital to comprehend how electron images are produced and how samples are handled in order to derive valuable data from a micrograph. This handbook will provide the novice with the procedural and theoretical information they need in order to make sense of the final product - the electron photomicrograph. The Transmission Electron Microscope (TEM), as the name implies, transmits a high energy electron beam through a specimen in a high vacuum environment. The vacuum environment is required to prevent electron interactions with air molecules which would serve to randomly scatter the electrons. In order for the electron beam to penetrate a sample, it must be extremely thin, approximately 600-900Å thick. Most samples will have to be sectioned using an instrument known as an ultramicrotome. Soft tissues will need to be mechanically strengthened by epoxy resin embedment to withstand the forces of cutting. The TEM can be viewed as an inverted light microscope (LM), with the source at the top of the instrument. The source of the TEM is a high voltage gun (50,000 volts or higher) which contains a pointed tungsten hairpin filament across which the high potential (voltage) is applied. The filament is housed in a metal cylinder with a central aperture (circular opening). This cylinder is known as the Wehnelt cylinder or Bias Cap since it is held at a slightly negative potential (the “bias”) with respect to the filament. This causes initial electrostatic repulsion of electrons coming off of the filament and the saturation of the Wehnelt cylinder. An anode, also with a central aperture, is held at ground potential. Due

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ELECTRON MICROSCOPY: A HANDBOOK OF TECHNIQUES FOR THE BIOLOGIST

to the large potential difference between the cathode, now saturated with electrons, and the anode, the electrons are accelerated through the distance between these two points this is known as the accelerating voltage. Wide-angle electrons will be grounded out and some will continue at high velocity through the anode aperture in order to contribute to image formation. The electrons will continue to the first lens in the TEM, the condenser. This lens will refract or bend the source electrons to the specimen. It should be noted that the lenses of an electron microscope are electromagnetic consisting of an iron shroud with a central bore and external copper windings. By varying the current through the copper windings, the focal point of the lens can be modified. This effect can allow for adjustments to brightness, magnification and focus. Once the electrons are focussed by the condenser lens, they will encounter the ultrathin specimen which is mounted on a grid and placed in a finely machined mechanical stage (with fine micrometer X,Y movements). Based on the electron density of various regions of the sample, some electrons will be backscattered while others will continue to transmit through the specimen. These transmitted electrons will next encounter the short focal length objective lens assembly which serves as the main imaging lens in the TEM and is critical to final magnification and focus of the image. This region of the TEM also includes an electromagnetic stigmator which is designed to reshape objective lens asymmetry arising from imperfect lens bores and lens and aperture contamination. Astigmatism (stigma meaning spot) results in stretched images of poor resolution. The octupole (8-pin) stigmator reshapes lens asymmetry by creating an asymmetric elliptical field to counter the lens distortion. The electron beam finally passes through a magnifying/demagnifying intermediate lens and then a magnifying projector lens which also functions to project the final real image on a fluorescent viewing screen. Since humans are not sensitive to electrons, the fluorescent material of the view screen is necessary to form an image that we can see. Electrons which pass through the electron transparent regions of the specimen will strike the fluorescent material causing it to emit photons which our eyes are sensitive to. Such region will appear bright. Specimen regions which are stained (usually with heavy metals) are electron dense and will not allow the transmission of electrons. They will not come in contact with the fluorescent screen and these regions will appear dark. This provides the contrast vital to image formation. A piece of film can be introduced via a camera mechanism beneath the view screen and a permanent exposure recorded. This will be explained in Unit 3. In the TEM, a direct image is formed by the differential absorption of a transmitted electron beam. Electron dense versus electron transparent specimen regions are ultimately responsible for contrast. Resolving power (RP), the ability to distinguish two points as two separate and distinct points, is based on the wavelength of the transmitted electron source. Resolving power and source wavelength are inversely proportional. As wavelength decreases, the resolving power increases as given by the Abbe equation: 0.61λ RP = ———— n (sin α)

INTRODUCTION TO ELECTRON MICROSCOPY

3

where λ is the source wavelength, n is the refractive index of the medium through which the source passes and α is one-half the objective lens acceptance angle. The relation n (sin α) is also known as the numerical aperture (NA) of the lens and under most ideal situations is usually equal to approximately 1.0, making the numerator of the equation most significant. In simple terms, the resolving power is essentially equal to approximately one-half of the source wavelength. For visible light, the shortest violet range wavelength is on the order of 400nm (equivalent to 4,000Å or 0.4µm). Given the above equation, the highest resolving power attainable using visible light is about 0.2µm (bacterial cell range). Electron wavelength is based on the de Broglie relationship stated as follows: h λ = ———— mv where λ is the source wavelength, h is Planck’s constant, m is the mass of the particle (such as an electron) and v is the velocity of the particle. Substituting electron mass and velocity at one-third the speed of light (achieved using a 50kv electron gun/accelerating voltage, the wavelength of the electron is approximately 0.05Å. Given the Abbe relationship, the theoretical resolving power of a 50kv TEM would therefore be 0.025Å. Unfortunately, due to lens aberrations (spherical aberration, chromatic aberration and astigmatism) which cannot be totally corrected for in an EM, the actual resolving power limit for a modern TEM is about 2Å - easily molecular resolution, approaching the atomic level (for example, the naked DNA double helix is 20Å in width). It becomes clear that the higher the accelerating voltage at the gun, the greater the electron velocity will be and the shorter the electron wavelength leading to a higher resolving power. Most modern TEM’s have maximum accelerating voltages exceeding 100kv. In order to achieve the highest resolving power, a single constant electron wavelength is required otherwise, chromatic aberration will degrade resolution. Voltage stabilization circuitry is critical with the absence of a source spectrum and therefore, the absence of “color” images - unless one uses computer enhancements or Dr. Martin’s dyes applied directly to photographic prints. In the year 1938, Knoll and von Ardenne constructed the first Scanning Electron Microscope (SEM) prototype. They suggested that secondary electrons could be collected from the tops of opaque surfaces, the resultant signal then amplified and used to modulate the grid of a cathode ray tube (CRT). It took many years of refinement to produce a commercial SEM (1963) - the Cambridge Stereoscan. By comparison, the first commercial TEM (1938) - the Siemens Elmiskop, was available soon after the initial TEM development. At first glance, the SEM appears to have a simpler design than the TEM with a noticeably shorter column. The TEM and SEM have an identical electron gun/anode design, however,

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ELECTRON MICROSCOPY: A HANDBOOK OF TECHNIQUES FOR THE BIOLOGIST

the SEM maximum voltage is approximately 25-30kv when using a tungsten hairpin filament. What follows is simply a series of 2-3 condenser lenses which serve to demagnify the primary electron beam diameter. In the SEM, resolving power (~30-40Å) is dictated by the diameter of the primary electron beam which scans the specimen surface in a raster pattern, much like the electron gun(s) in your TV scan the phosphorus pixels (picture elements) coated on the inside of the picture tube, in order to form an image. The SEM scan generator is responsible for the raster scan of the sample surface by the primary electron beam. When the primary electron beam contacts the sample surface, a variety of energetic phenomena arise which can be detected and collected (provided the SEM is outfitted with the appropriate detector). The most common type of energetic phenomenon emitted is the secondary electron signal. This secondary electron signal is collected to form the typical, virtually three-dimensional image that is usually encountered in textbooks and publications. Other types of signal include backscattered electrons (BSE), characteristic x-rays (which can be used to create an elemental surface map, and photons or cathodoluminescence. Once again, each require a specific detector which are easily added to a modern SEM. The weakly negative secondary electrons are emitted after surface contact with the primary electron beam. These secondary electrons are collected by a scintillatorphotomultiplier detector. The signal is then amplified and routed to the electron gun of a black & white viewing CRT. Whether the pixels remain dark or light up is related to the level of signal arising from a given specimen area. Maximum signal gives rise to a bright/ white pixel, whereas, minimum signal results in a dark/black pixel. The final result is contrast and a black and white image on the viewing CRT. The amount of signal emitted from the specimen surface is a function of its topography or relief. High points of relief, in direct line of sight with the detector and primary electron beam, will produce the maximum signal and appear brighter than low lying areas, which will appear dark. The final image is indirect, based on point by point differential contrast due to the yield of secondary electrons from the sample surface. Magnification is a function of the length of a line scanned on the sample as compared with the length of a line scanned on the viewing or photo CRT. Since the CRT dimensions never change, a magnification increase is effected by simply modifying your scan generator control to scan a smaller area/shorter line on the sample. Samples are mounted on aluminum stubs which are placed into the finely machined stage of the SEM. The stage has provisions for movements in the X, Y, Z (also known as working distance) directions, including T (tilt) and R (continuous 360˚ rotation). A benefit of the SEM image is its high depth of field and focus which leads to the striking, almost three-dimensional images. In addition to the components described above, a stigmator is required in the design of a SEM and is one of the most difficult adjustments to teach the novice. Once again, the X, Y stigmation is required to compensate for lens asymmetry which arises from lens imperfections and contamination. One must learn to see the image stretching as you go

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INTRODUCTION TO ELECTRON MICROSCOPY

through fine focus in order to correct it with the stigmator. The following illustration compares the design of the LM, TEM and SEM:

Light Microscope (inverted) illuminator/bulb

Transmission Electron Microscope (TEM) source

condenser lens

glass: fixed focal length

(focus source at specimen level)

TEM double condenser (increases brightness)

iris diaphragm

+

-

+ electron gun

electromagnets: variable focal length via variation in lens current

Demagnifies beam from 50,000 Å to 100 Å

First Condenser Lens camera

condenser aperture

aperture (reduces extraneous source)

specimen

glass slide

Scanning Electron Microscope (SEM)

grid

Second Condenser Lens

insertion mechanism

objective lens

Magnification Control

(magnify & focus)

objective aperture (TEM) Third Condenser Lens

(grounds stray electrons which increases contrast)

intermediate lens (1-3)

Final Aperture

(TEM only - magnify)

Signal (secondary electrons)

projector lens ocular(s)

(magnify & focus final real image

Detector

on retina of eye [LM] or fluorescent view screen [TEM]) eyes/retina

fluorescent screen (TEM) (direct imaging)

camera film

(to record permanent, high resolution image)

film

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ELECTRON MICROSCOPY: A HANDBOOK OF TECHNIQUES FOR THE BIOLOGIST

UNIT 1 - PREPARATION OF BIOLOGICAL SAMPLES FOR TEM The high vacuum environment (10-5 Torr) of the transmission electron microscope presents a major and obvious obstacle for the study of biological samples. Since death is the inevitable result of introducing a living organism into a vacuum environment, we are limited to the examination of dead specimens. Another problem that prevents the study of live organisms is the requirement that samples be sectioned ultrathin. Unless samples are thin enough, between 600-900Å, the electron beam cannot transmit or pass through it. Once cut, a cell cannot typically be expected to live. Unfortunately, as a result of these two limitations, we are not able to study actual processes occurring directly within the cell, even though the TEM has the resolving power to do so.

Chapter 1 - Chemical Fixation Chemical fixation involves killing and preserving the organism/organ/tissue/cell in as lifelike conditions as possible. The biological structures and their functions are fixed or “frozen” in time and space. Our ability to observe fixed biological ultrastructure using the TEM allows us to infer and come to understand function. The true goal of fixation is to enable the investigator to examine the structure(s) they are interested in (to see what you want to see!). Since we now understand that structure and not direct function is studied using the TEM, we must determine the major factors which influence cellular structure. Cellular organelles such as ribosomes, mitochondria, etc. are a variety of molecules arranged in a specific three-dimensional architecture, with the TEM capable of resolution at the molecular level. What is responsible for this cellular ultrastructure that we can distinguish with the TEM? Firstly, is the importance of the most abundant molecule in the living cell, water, and its effect on other cellular molecules. Water is a charged dipolar molecule. Being charged, it easily interacts with other charged molecules including the charged “R” groups of the protein’s amino acids (proteins being the second most abundant molecule of the cell). Polar water is incapable of interacting with neutral and non-polar molecules/groups which are designated as hydrophobic (“water fearing”). The charged groups are known as hydrophilic (“water loving”). The primary order of protein structure is its amino acid sequence which includes a number of both hydrophobic and hydrophilic units. Aside from many types of interactions between amino acids of the protein (hydrogen bonding, ionic interactions, disulfide bridges, etc.), the interaction of water with these hydrophobic and hydrophilic units is crucial to the higher order protein structure, meaning, how it folds into a three-dimensional protein structure. In this scenario, hydrophilic amino acids would be found to the outside of the protein molecule, while the hydrophobic units would be found at the center of the molecule, “hiding” from the external, much more numerous, water molecules. These interactions also explain the orientation of phospholipid molecules which makeup cellular membranes (phospholipid bilayer). Secondly, we must recognize that cellular physiology and metabolic processes are responsible for maintaining cellular conditions favorable to the continuity of molecular,

UNIT 1 – PREPARATION OF BIOLOGICAL SAMPLES FOR TEM

7

hence, organelle structure. Examples of some conditions which must be maintained (homeostasis) would include temperature, pH, and osmolarity. Any changes in these conditions during fixation could lead to structural distortions mainly through the process of denaturation. Denaturation, the alteration of the 3-D shape of a molecule (i.e. protein), must be considered when selecting fixatives for TEM specimen preparation. With the high resolving power of the TEM, any minor alteration in the threedimensional conformation of molecules is undesirable and would lead to the formation of artifacts (structures not normally present in the cell which are produced by some external intervention or agent). You might ask what chemical fixatives are doing and what distinguishes a good fixative from a poor fixative, relative to TEM preparation of biological samples. The main requirements of a fixative are that it stabilizes cellular ultrastructure without inducing distortions/artifacts. In stabilizing cellular ultrastructure, the fixative must prevent an undesirable process known as autolysis (meaning self-dissolution/self-digestion) from occurring. When an organism dies, cellular lysosomes begin to burst open, releasing a flood of hydrolytic, digestive enzymes into the cell. These autolytic enzymes would obviously degrade the very cellular microanatomy which the TEM has the power to resolve. Since autolytic changes proceed rapidly after death, it becomes important to introduce the fixative to the tissue as soon after death as is possible, ideally within 30 minutes. In summary, fixation must kill and simultaneously stabilize cellular components through the prevention of autolysis upon death of the organism (whether it be unicellular or multicellular). As noted earlier, it is critical that the fixatives do not denature cellular molecules as a result of their stabilizing components and preventing autolysis. There are a variety of fixation methods that are utilized world-wide in order to kill and preserve living materials. Most of these methods are not suitable to the fixation of samples for TEM due to the structural distortions/denaturation which result. A list of these methods is presented below with an explanation of their value or uselessness to TEM fixation. It should be pointed out that each method is capable of halting autolysis associated with the death of the organism. This fact qualifies each as a preservative. • Air Drying This method is employed for the preservation of harvested grains (wheat, etc.). Drying eliminates the water necessary for autolytic enzyme shape, and therefore, activity. Since all cellular proteins are denatured, this method is poor for TEM fixation. • Pickling The pickling process is used to preserve cucumbers (pickles), beets, etc. Preservation is made possible through a change in pH which denatures autolytic and other cellular proteins. The pH is usually lowered (acidic) through the introduction of acetic acid (vinegar). Due to denaturation, this method is also poor for TEM

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ELECTRON MICROSCOPY: A HANDBOOK OF TECHNIQUES FOR THE BIOLOGIST

fixation. • Alcohols Alcohols, such as ethanol (CH3CH2OH) generally make good fixatives since they are all dehydrating agents. Removal of water denatures autolytic enzymes along with other cellular proteins making alcohols unsuitable for TEM fixation if used alone. In TEM (and SEM) fixation, dehydration is an important step in the protocol, however, this process is carried out only after the tissues have already been stabilized with the primary and secondary fixing agents. • Heat The addition of heat energy causes the breakage of numerous bonds which are responsible for the 3-D conformation of cellular proteins, including the autolytic enzymes. The enzymes are denatured and the organism is preserved. Heat is used in the canning process. An example would include the cooking of vegetables prior to vacuum packing in the cans. Due to the resulting denaturation caused by heat, TEM samples cannot be simply “cooked” in order to preserve cellular ultrastructure. • Oxidizers/Precipitants Powerful oxidizing agents such as Osmium Tetroxide (OsO4) and Potassium Permanganate (KMnO4) were the first solitary fixatives used for TEM biological sample preparation. Both react at the double bonds present within unsaturated lipids, such as those found in abundance in the composition of biological phospholipid membranes. In addition, and as a result of their reaction with unsaturated lipids, they introduce an electron dense, heavy metal (Os/Mn precipitate) to those redox reaction sites. This resultant staining enhances contrast of TEM samples. It should be noted that OsO4 was the primary and only TEM fixative prior to 1963. Since that time it has been discovered that these violent oxidizers can destroy delicate, labile (changeable) cellular structures such as microtubules. In 1963 a different primary fixative was proposed for TEM samples which eliminated such redox reaction damage, of especially, proteinaceous components. KMnO4 is such a powerful oxidizer that all of the internal cellular cytoplasmic components are destroyed, with the exception of the membrane systems. If your goal is the study of a particular membrane system, you might wish to try KMnO4 in conjunction with other protocols. Currently, OsO4 is used as a secondary or postfixative due to its ability to react with and stain membranes of the cell. It may also react with and stain some protein elements of the cell which have been previously stabilized using aldehydes. CAUTION: Osmium tetroxide is a highly reactive and potent fixative. The vapors alone can fix exposed epithelial surfaces of the mouth, nose, and especially, eyes (cornea) resulting in temporary blindness. Crystalline or in aqueous or buffer solution, osmium tetroxide should be handled with extreme care. Gas tight goggles, double gloves and a fume hood capable of 150 cfm flow

UNIT 1 – PREPARATION OF BIOLOGICAL SAMPLES FOR TEM

9

rate are the minimum safety recommendations. Polyunsaturated oil, such as corn oil, should be at hand in the case of a spill. It has been advised that approximately twice the volume of oil be added to an osmium tetroxide spill in order to neutralize it. If spilled in the open, the area should be vacated immediately and the proper authorities notified. No one should be allowed to reenter without a proper respirator. Waste osmium tetroxide should not be put down the sink but rather, stored in a clearly labeled waste bottle for later disposal by environmental carting firms.

• Aldehydes Aldehydes, containing the reactive carbonyl group (C=O), have been used for years in the preservation of biological specimens (embalming, light level histology, etc.). It is somewhat surprising that they were not considered for TEM fixation prior to 1963 when Sabatini, Bensch and Barrnett introduced them as the primary fixative in TEM sample preparation. The role of aldehydes in fixation is their stabilization of the cellular protein matrix into a somewhat gelatinous state. The carbonyl groups react with any reactive amino acid “R” group which in turn results in the methylation (-CH3) of the protein molecules. Neutral methyl groups interact between adjacent protein molecules resulting in velcro-like linkages and stabilization. Since the aldehydes do not denature the proteins, their active sites remain intact and histochemical and immunocytochemical (ICC) localization of proteins/enzymes can be performed. The fact that they do not denature make them ideal primary fixatives for TEM biological samples. The most common types of aldehydes used in TEM fixation are formaldehyde (HCHO), acrolein/acrylic aldehyde (CH2 • CHCHO) and the dialdehyde known as glutaraldehyde (CHO-(CH2)3-CHO). While both formaldehyde and acrolein are smaller molecules which diffuse more rapidly into tissue blocks, glutaraldehyde, the dialdehyde, is doubly reactive and allows for increased stability of the cellular protein matrix through the cross-linking of protein molecules. Although glutaraldehyde is the most commonly used single aldehyde fixative, an examination of the literature often reveals the use of a combination aldehyde primary fixative. In this case, glutaraldehyde is usually used in combination with one or both of the other aldehydes mentioned above. Conventional formalin solutions are not suitable for TEM fixation. Since formaldehyde is a gas in nature, it must be prepared as an aqueous solution. Through reaction of formaldehyde with water, formic acid formation results. This lowers the pH and serves to denature proteins. In addition, formalin contains methanol which is a dehydrating agent and also denatures. Paraformaldehyde, a purified crystalline trimer of formaldehyde, is the compound of choice for TEM fixation. Under low heat for 30min duration, paraformaldehyde goes into aqueous solution without formic acid formation and without the addition of methanol. Although acrolein is an excellent primary aldehyde fixative, it is difficult to work with since it is highly explosive and a potent lachrymator (tear gas).

10 ELECTRON MICROSCOPY: A HANDBOOK OF TECHNIQUES FOR THE BIOLOGIST

CAUTION: As fixatives, all aldehydes should be handled with gloves, goggles and under a fume hood, especially acrolein since it is a lachrymator. Acrolein is also highly explosive and should be kept away from direct light, heat and flames. All used aldehydes should be stored in clearly labeled organic waste bottles for later safe disposal. Never put toxins down the drain!! ✥ Additives In the literature, you will notice that most fixatives are carried in a vehicle known as a buffer and may also include other additives such as salts (CaCl2) or even sucrose. The main purpose of these additives is two-fold. Firstly, the buffer is important to the maintenance of the natural physiological pH of the tissue being fixed. Buffer solutions can react with and counteract the release of excess H+ and/or OH- ions from the tissue being fixed. As discussed earlier, a change in pH would denature and is therefore not desired. Many types of buffers have been used in TEM fixation including veronal-acetate, PIPES, chromate, phosphate and cacodylate. Even though it has a short shelf life due to the eventual growth of bacteria, phosphate buffers are a good choice since they are non-toxic and therefore, easy to work with out in the open. Once prepared (there are many recipes such as Sorensen’s), phosphate buffers should be refrigerated at 4˚C to reduce bacterial growth. Cacodylate buffer should be handled with care under a fume hood since it contains arsenic. Fixatives are typically made up in the buffer solutions just prior to use. Buffers should be adjusted to the physiological pH levels of the organism’s internal (or external as in the case of unicellular protozoa) environment. Mammals and a number of other animals range between pH values of 7.2-7.5 with botanical samples approximately 6.8. The use of additional salts and/or inert substances such as sucrose are for osmolarity/ tonicity considerations. The fixative should be isotonic with the internal/external fluid environment of the tissue under study. Although trial and error is a common practice in determining the proper osmolarity, reference sources that list the osmolarity of specific animal blood/body interstitial fluids are available. Some labs also have access to an osmometer in order to determine precise tonicity requirements for the fixative vehicle. Another additive involves the use of dimethyl sulfoxide (DMSO) which is a penetrant used to enhance infiltration of fixatives into the tissue block. ✥ Fixation Times & Temperatures In the EM literature, you will find a wide range of fixation times and temperatures used. Who is correct? Can they all be right? Perhaps. One must consider the quality of the final micrograph that is being produced. Relative to the duration of each step in a TEM fixation protocol, there are optimum times of exposure and times which work around technician schedules for the sake of convenience. In the specific fixation of soft mammalian tissues which follows, the indicated times are optimal for ideal preservation with the minimum of

UNIT 1 – PREPARATION OF BIOLOGICAL SAMPLES FOR TEM

11

tissue component extraction and artifact production. In the literature you may see primary fixation in glutaraldehyde for 1hr to overnight. Under ideal circumstances, fixation through the tissue block will occur within 1hr, provided it is small enough in dimension (0.5 cu.mm.). Optimal times should be used whenever possible and practical. There are two schools of thought on the proper temperature of fixation. One suggests that fixation should be conducted in the cold, on ice, at 4˚C to slow autolytic enzyme activity and reduce extraction of cellular components. The other suggests that room temperature fixation hastens fixative infiltration and the actual biochemical process of fixation. In the cold, we are actually slowing the desired process and in some cases, preventing it (if fix time is inadequate) which can lead to the production of artifacts. The best recommendation is to begin the process/protocol at 4˚C and allow it to come to room temperature (probably by the time you reach the ethanol dehydration series). ✥ Preparation of Fixatives • Phosphate Buffers: Used as a vehicle to carry the various fixatives and as a wash. The solution is prepared using monosodium (NaH2PO4) and disodium (Na2HPO4) phosphate. Solutions are usually prepared with molarities that are consistent with the tonicity of the sample, such as 0.02M, 0.05M, 0.1M, 0.2M, 0.5M. The molecular weight in grams should be added to one liter of distilled water to yield a 1.0M solution. Simple calculations can be performed to reduce the molarity from 1.0M and to prepare 100ml of solution vs. 1,000ml (divide by 10). Take note of whether the sodium phosphate is hydrated since the addition of one or more water molecules will increase the gram molecular weight. By way of example, to prepare a 1.0M solution of monohydrated monosodium phosphate (NaH2PO4 • H2O), the gram molecular weight, 138g, is added to 1,000ml of distilled water. A 0.1M solution would be prepared by adding only 13.8g to 1,000ml of DH2O. To prepare 100ml of the 0.1M solution, only 1.38g would be added to 100ml of DH2O. In order to adjust the pH of the buffer solution, the proportions of monosodium relative to disodium phosphate must be varied according to the chart below.

pH

6.0

6.2

6.4

6.8

7.0

7.2

7.4

7.6

7.8

8.0

NaH2PO4 (in ml)

87.7

81.6

73.5

51.0

39.0

28.0

19.0

13.0

8.5

5.3

Na2HPO4 (in ml)

12.3

18.4

26.5

49.0

61.0

72.0

81.0

87.0

91.5

94.7

Phosphate buffers should be freshly prepared and stored in the refrigerator at 4˚C to reduce the growth of bacteria. The pH should be checked with a pH meter and adjusted as necessary. Buffer solution(s) should obviously be prepared first, in advance of any fixative since the fixatives are mixed with the buffer.

12 ELECTRON MICROSCOPY: A HANDBOOK OF TECHNIQUES FOR THE BIOLOGIST

• Glutaraldehyde: This primary fixative is available in bottles in aqueous solutions of 25% and 50% and in ampoules sealed under nitrogen gas in 8% aqueous solution. Due to polymerization of the glutaraldehyde in high concentrations, the 8% sealed ampoules afford increased shelf life. Whatever concentration is used, it must be added to an appropriate amount of buffer solution to yield the final concentration (approximately 3% is ideal for most purposes). For example, to prepare 3.2% glutaraldehyde in buffer, 20ml of 8% glutaraldehyde is added to 30ml of phosphate buffer. The buffered glutaraldehyde should be made just before use and stored in the refrigerator in the dark. • Osmium Tetroxide: OsO4 is available as crystals in 0.5g and 1.0g quantities, in sealed ampoules, or in aqueous solution (such as 2% and 4%) in sealed ampoules. Crystalline OsO4 is ideal for making larger quantities (50-100ml) of the working solution which is usually set at 1-2%. A meticulously cleaned and dry ground glass stopper bottle with a teflon seal should be used to prepare the working solution. Since crystals of OsO4 are large and dissolve slowly, it is recommended that the ampoules be held under running hot water which allows the crystals to melt. The ampoule is then gently rolled between the gloved hands to permit recrystallization of the OsO4 in a thin layer inside the ampoule. The ampoule is inserted into the ground glass bottle and the stopper introduced. The bottle is shaken which causes the ampoule to break inside the bottle (Note: EM supply companies currently use pre-scored ampoules, some in conjunction with a plastic external cylinder to prevent a cutting injury. The ampoule should be opened, placed in the bottle, and the ampoule filled with buffer using a pipette. The bottle is then filled with the remaining buffer. It is important that the ampoule sink to the bottom of the bottle in order for the OsO4 to go into solution). Once the ampoule is broken, the appropriate amount of buffer is added to yield the final working solution. To prepare a 1% OsO4 solution, use 1.0g of crystalline OsO4 in 100ml of buffer (or 0.5g in 50ml of buffer). When using the solution, be careful not to pipette near the bottom of the bottle since you may pick up some glass shards and introduce them to your tissue samples causing physical/mechanical damage. Once again, it must be cautioned that osmium tetroxide is extremely toxic and should be handled under the fume hood wearing gas tight goggles and double gloves. Cooking oil/ corn oil should also be available for possible spills. As with all fixatives, OsO4 is prepared just before use. It is ideal to prepare the working solution one day before use and leave it out at room temperature overnight. After use, the solution should be stored in the refrigerator. It should be tightly stoppered and placed inside another container of glass or metal to prevent escape of vapors which will also react with and blacken the inside of the refrigerator. • Ethanol Dehydration Series: For TEM specimen dehydration, 70%, 95% and 100% ethanol are required. Ethanol concentrations of 95% and 100% are available. It is ideal to maintain the volume of absolute (100%) ethanol since air in the storage vessel contains water vapor which mixes with the ethanol and reduces the concentration. Small pint bottles are available for one time use since it is imperative that no water remain in the tissues. Another storage technique involves standing the 100% ethanol over a layer of anhydrous cupric sulfate which serves to absorb water. It should be changed when it begins to turn blue which indicates that it is hydrated. Do not mix up the cupric sulfate

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and EtOH prior to use. The cupric sulfate must be allowed to settle on the bottom of the container. In order to prepare ethanol dilutions of less than 95%, one should not use the more expensive 100% ethanol. In this case, 95% ethanol is used as follows: 1. The amount of 95% ethanol in ml, equal to the desired final concentration, is measured out (for example, a 30% EtOH solution begins by measuring out 30ml of 95% EtOH). 2. Distilled water is added to make up a total of 95ml of solution (to complete the above 30% EtOH solution, 65ml of DH2O is added to the 30ml of 95% EtOH - total volume is 95ml). Ethanol solutions can be refrigerated in the dark until ready for use. • Propylene Oxide: Propylene oxide is available from EM supply companies usually in 250ml quantities. It should be used undiluted. The addition of water would defeat the purpose of the prior ethanol dehydration series and prevent the infiltration of epoxy resin into the tissues. It is important to know that propylene oxide is extremely volatile. If you are not careful in the solution exchange process, your tissues will dry down leading to artifacts. Propylene oxide (1,2 epoxy propane) is a small, rapidly diffusing, molecule which introduces the epoxy monomer into the tissues thus aiding infiltration. It serves as a transitional solvent as it is miscible with both ethanol and the mixed, unpolymerized epoxy resin. CAUTION: Propylene oxide is extremely flammable and should be stored in a cool dark environment. Never expose it to direct heat or flame. It should be handled with gloves, goggles and under a fume hood since it is a suspected carcinogen. • Epoxy Resins: The use of epoxy resins is ideal for the embedment of biological samples for ultrathin sectioning for the TEM. Due to its three-dimensional polymerization, samples are provided with excellent mechanical strength capable of tolerating the forces of sectioning. The preparation of epoxy resins involves mixing the epoxy compound(s) such as “Epon” 812 and/or Araldite 6005, with specific curing agents including acid anhydrides (DDSA) and tertiary amines (DMP-30). Additionally, depending on the epoxy resin used, a plasticizer such as dibutyl phthalate (DBP) may be necessary. It is critical that all components be thoroughly mixed. The Epon-Araldite mixture used below has proven excellent for the embedment of soft mammalian tissue samples. The stock solution and the final working solution should be hand mixed for at least 15 minutes each. The stock solution can be frozen for future use. A plastic syringe can be used to store the stock solutions in exact volumes and with minimal trapped air which could lead to condensed water vapor contamination of the stock solution as it is allowed to come to room temperature. Stock solutions must be allowed to come to room temperature prior to their use in the preparation of the working solution.

14 ELECTRON MICROSCOPY: A HANDBOOK OF TECHNIQUES FOR THE BIOLOGIST

Mixing of epoxy resins introduce tremendous amounts of air which should ideally be removed. Air molecules present in the resin mixture would prevent proper infiltration of the resin into the tissues. Due to the viscosity of the mixed resin, outgassing through standing at atmospheric pressure would be a lengthy and incomplete process. Therefore, mixed resin should be put under a low vacuum environment in order to remove air. Care should be used in the rate of achievement of vacuum since the resin mixture will overflow the container, carried out by air bubbles, and contaminate the bell jar. The vacuum should be achieved slowly by regulating the air inlet valve. Contamination of the work area with unpolymerized resin should be a definite concern. Be careful not to touch doorknobs, refrigerator handles, etc. with contaminated gloves. Never pour unpolymerized resins into the sink. Resins should be mixed in disposable plastic beakers using glass rods. Excess resins should be polymerized which renders them safe for disposal. CAUTION: Epoxy resins and curing agents can cause contact dermatitis and may be carcinogenic. All resin components should be handled in a fume hood and with double gloves (immediately discard the outer pair if they become contaminated). When outgassing mixed resins, the low vacuum setup should be put in a fume hood. Ovens for polymerization of resins should be vented to the outside or placed in a fume hood. Although the Epon-Araldite mixture below is prepared volumetrically, the most accurate way to measure it is by weight. Due to the viscosity of the resins, a quantity is certain to remain behind in the serological pipette used to measure it. Epon-Araldite Mixture Stock Solution (can be frozen) Araldite 6005 Poly/Bed 812 Dibutyl phthalate

Small Volume 12.5ml 15.5ml 2ml

Large Volume 25ml 31ml 4ml

Final Working Solution Stock Solution DDSA DMP-30

4ml 8ml 10ml 20ml 14 drops* 28 drops* (*Drops introduced with a Pasteur Pipette)

Another useful formulation is an Epon mixture ideal for embedding directly in plastic or glass containers such as tissue culture plates, petri dishes, microfuge tubes, etc. This

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mixture introduces nadic methyl anhydride (NMA) in addition to DDSA. Note the slight difference in the volume of NMA used in conjunction with original Epon 812 resin vs. the “812” resins available today from EM supply houses (Poly/Bed 812-Polysciences, Inc., EM Bed 812-EMS, Inc., etc.)

Epon for Glass/Container Work Component

Volume using Epon 812

Volume using “812” substitute

Epon 812 or 812 substitute

12.5ml

same

DDSA

6.5ml

same

NMA

6.25ml

9.0ml

DMP-30

1.0ml

same

DBP

0.25ml

same

Mixed resins will polymerize at 60˚C for 48 hours. If heat is a concern, epoxy resins will also cure under ultra-violet (UV) light. ✥ Methods of Fixation • Immersion: In this type of fixation, the tissue of interest is removed/excised from the organism (or if small enough, the entire organism) and placed into the primary buffered fixative such as glutaraldehyde. The buffered primary fixative can be put into petri dishes or as large drops on a card of dental wax. When a complex animal (mammal, etc.) is used, it must be sacrificed quickly, followed by opening the body cavity, identifying the organ(s)/ tissue(s) of interest, and rapidly excising them using a razor blade or scalpel. Once the tissues are transferred to the fixative, they must be minced to blocks which are less than 1.0mm3 in dimension (0.5mm3 is optimal) using clean single-edged razor blades. Mincing should be done with care since mishandling will result in mechanical damage artifacts. Artifacts will also arise if the tissue pieces are allowed to dry. It is imperative that they be kept submerged in the fixative solution at all times. Once minced, the tissues are transferred to small vials containing the primary fixative and allowed to remain for the allotted time (usually about one hour). All subsequent steps in the protocol (with the exception of epoxy resin embedding) can be carried out in these vials through decanting the current solution and quickly, yet carefully, introducing the next solution in the protocol.

16 ELECTRON MICROSCOPY: A HANDBOOK OF TECHNIQUES FOR THE BIOLOGIST

• In-situ: In-situ fixation, meaning ‘in the original place’, is accomplished by bringing/ applying the primary fixative to the specimen of interest. By example, fixation of a thin plant leaf is conducted by placing a ring of petroleum jelly on the leaf surface and filling the ring with the primary fixative. Once fixed, the area of interest is cut out, minced and placed into vials to continue with the protocol. Another example involve the fixation of tissue culture cells. The fixatives are simply administered to the culture dishes through pouring them on and off of the cellular monolayer. In the animal fixation described above under immersion fixation, once the animal is humanely sacrificed and the body cavity cut open, primary fixative should be poured into the cavity in order to initiate fixation. • Vascular Perfusion: This technique uses the vascular system of an animal to deliver the primary fixative deep into the tissues and directly on target. This is probably the most optimal means for administering the fixative. Delicate tissues subject to rapid postmortem change must be fixed in this manner. A good example would be nervous tissues. In this procedure, the animal is anesthetized and the heart and major blood vessels exposed. Typically, a canula is inserted into the aorta and a balanced saline solution is allowed to gravity feed into the blood vessel. Eventually, the saline solution is cut off by a clamp and the fixative is allowed to flow into the vascular system. The tissues of interest can be later excised, minced, and placed into vials in order to continue the process. • Fixation by Vapors: Small delicate surfaces, such as membranes, may be fixed in this manner. The specimens are simply suspended over a solution of osmium tetroxide, usually overnight. The samples will blacken to indicate that fixation has occurred. Samples can be dehydrated and embedded in resin for ultrathin sectioning. In our lab, we use a combination of in-situ and immersion fixation of biological samples as described above. Once again, care and common sense should be used in the handling of all fixatives! ✥ “Routine” Biological Soft Tissue Protocol In fact, no single “routine” protocol for the fixation of soft biological tissues exists. There are as many protocols as there are investigators, with no one better than the other. As long as your protocol enables you to observe the cellular features that you are interested in, it is valid. Of course, protocols will vary based on the particular organism being studied. It is apparent that botanical samples should be fixed using different protocols than animal samples, however, protocols will vary even among related organisms. By way of example, metazoans of different classes, orders and even genera will have varied physiologies which give rise to a wide range of body/interstitial fluid characteristics. As discussed earlier, in order to prevent artifacts, the buffer pH and tonicity would have to be considered for each animal investigated. In addition, conditions of fixation should be considered. Are you fixing under ideal laboratory conditions or are you in a tropical field setting? How much time will you have to mince the tissues after excision? You may have to store tissues in tropical climates for days to weeks without the ability to precisely mince the tissue blocks until you return to the lab. Even under these unfavorable conditions, you

UNIT 1 – PREPARATION OF BIOLOGICAL SAMPLES FOR TEM

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can modify your fixation protocol to yield quality results. In a study of neo-tropical bats, Nagato, Tandler and Phillips used a trialdehyde-DMSO primary fixative which consisted of 1% glutaraldehyde, 1% paraformaldehyde, 0.5% acrolein, 2.5% dimethyl sulfoxide, and 1 mM CaCl2 in 0.05 M cacodylate buffer (pH 7.2). The tissue was stored for 14 days in tropical conditions before being transferred to 3% glutaraldehyde at 4˚C. Given the circumstances, the resulting TEM micrographs were of extremely good quality. The protocol which shall now be outlined relates primarily to soft mammalian tissue samples prepared under ideal laboratory conditions. The solutions are initially at 4˚C and are allowed to come to room temperature. The steps in this “routine” protocol are as follows: • Primary Aldehyde Fixation - used singly, buffered glutaraldehyde (usually 3-4%) is the most common choice for TEM fixation of biological tissues. Using methods of fixation described earlier, the primary fixative is delivered to the tissue of interest. In this lab, the most common method employed for soft mammalian tissues is in-situ followed by immersion fixation in petri dishes. The tissue is minced carefully using two clean singleedged razor blades into pieces no larger than 3.0 cubic mm, with the ideal tissue block size no larger that 1.0 cubic mm. If you wish to complete mincing in the primary fixative, you will need to reduce the tissue blocks to 0.5 cubic mm, however, this can wait until the final buffer wash just prior to osmium tetroxide (OsO4 does not penetrate as well as the aldehydes). Once minced, the tissue blocks are transferred to small glass vials containing the primary fixative. The aldehydes will penetrate the tissue block to fix cellular proteins as described earlier. The ideal fix time for aldehydes is approximately 1 hour, however, tissues can be stored in glutaraldehyde at 4˚C for a number of days. I recall having stored samples in glutaraldehyde for as long as 1 year without the development of major structural artifacts. • Buffer Wash - usually the same buffer is used as is used to prepare the aldehyde fixative. In this lab, phosphate buffers are used. The buffer wash is done in order to wash out any unbound aldehydes which would react with the OsO4 in the next step, leading to a contaminating precipitate reaction. This wash is usually done a number of times (3 times) for ten minute intervals. • Osmium Tetroxide Postfix - buffered OsO4 in 1-2% solutions is administered to the tissues next. At this point, the tissue blocks must be on the order of 0.5 cubic mm in dimension. The OsO4 primarily reacts with unsaturated fats in the cell and imparts an electron dense heavy metal staining to regions of the cell which are composed of these lipids (such as cell membranes). OsO4 also reacts with some cellular proteins, however, because aldehydes were used first, the proteins should now be stable and not subject to denaturation by this powerful oxidizing agent. OsO4 postfixation is usually carried out for 1 hour with the tissue samples blackening to indicate the activity of the fixative.

18 ELECTRON MICROSCOPY: A HANDBOOK OF TECHNIQUES FOR THE BIOLOGIST

• Distilled Water or Buffer Wash (Optional) - many investigators use a brief wash of either distilled water or buffer to remove excess OsO4. The wash is performed since the OsO4 may react with the ethanol in the following step. At times, a cloudiness may develop in the fixation vials if this step is avoided. In this lab, we usually dispense with this wash since the cloudy result is rare and short-lived, usually lasting for the initial ethanol change. If this step is performed, usually 2-3 changes of ten minutes each is sufficient. • Dehydration Series (Graded Ethanol/Acetone Series) - the tissues are passed through an ascending dehydration series of either ethanol or EM grade acetone. To prevent distortion and shrinkage, many individuals ascend slowly to 100% ethanol or acetone, starting at 10% and ascending in 10% steps (10%, 20%, 30%, 40%, 50%, 60%, 70%, 80%, 90%, 100%). In order to preserve delicate, internal labile structures, it is also important to get the tissue out of an aqueous environment as soon as possible, therefore, we start dehydration of samples for TEM at 70% (two changes for ten minutes each). We then proceed to 95% (two changes for ten minutes each) and then to 100% (two changes for ten minutes each), typically using ethanol. The 100% ethanol vials must be completely filled to prevent air (containing water vapor) from reducing the concentration. The absolute ethanol should be standing over anhydrous cupric sulfate before use to ensure it is not hydrated. This is a critical step since any water remaining in the tissue will prevent epoxy resin infiltration. The dehydration series is performed since water and most embedding media (epoxy resins) are not miscible. • Transitional/Intermediate Solvent (Propylene Oxide) - this is an optional step in the protocol which will ensure successful epoxy resin infiltration. Propylene oxide (1,2 epoxy propane) is a highly volatile, flammable, small molecule which rapidly penetrates the tissue blocks. It serves to enhance infiltration of the tissues by epoxy resin since it carries in the reactive epoxy monomer. If used, three changes for ten minutes each is adequate. • Propylene Oxide : Mixed Epoxy Resin (1 : 1 ratio) - in this step, an equal volume of propylene oxide and mixed (unpolymerized) epoxy resin is introduced to the tissues. The purpose is to make a gradual transition to pure epoxy resin and enhance its infiltration into the tissue blocks. This step is carried out in vials for 1 hour. • Vacuum Infiltration - after the pure mixed epoxy resin has been placed under a low vacuum (bell jar and rotary pump setup) to eliminate air introduced through the mixing process, a small volume is poured into a small diameter petri dish (enough to fill the bottom and produce a thin uniform layer). Tissue blocks are selected and placed into the resin using toothpicks or bamboo sticks. The labelled petri dishes are placed into the low vacuum setup and the bell jar is pumped out. This process is carried out for 1 hour to prevent gas from impeding epoxy resin infiltration. • Embedding - During the 1 hour vacuum infiltration process, BEEM (Better Equipment for Electron Microscopy) capsules (which have a preformed truncated pyramid tip of 1.0 square mm) are stuffed with an identification label and filled with epoxy resin mixture to

UNIT 1 – PREPARATION OF BIOLOGICAL SAMPLES FOR TEM

19

a slight negative meniscus. Allow air bubbles to break naturally and further fill capsules as needed. Identification labels should identify the investigator and the tissue type. Write the labels in pencil only since pen and other inks can react with the resins. Once written, the labels are wrapped around a wooden stick (bamboo) and placed into the BEEM capsule. Don’t wrap the label too tightly around your stick since you want the label to spring against the inner walls of the capsule. When viewed from above, the labels should be barely visible. If they project into the center of the BEEM capsule, try to push them against the sides with two sticks. You do not wish the labels to impede the descent of the tissue blocks to the tip of the capsule. With the BEEM capsules labelled and filled with the epoxy resin mixture, individual tissue blocks can be transferred from the petri dishes to the center of the filled BEEM capsules using a pointed bamboo stick. The tissue blocks are heavy as a result of osmium impregnation and will sink to the capsule tips prior to the polymerization of the resin. BEEM capsules should be supported in some type of holder. In this lab we use old micropipette tip holders. Some EM supply companies manufacture BEEM capsule holders and some individuals simply punch appropriate sized holes into cardboard squares. With the tissues transferred to the BEEM capsules, the holders are placed into an oven at 60˚C for at least 48 hours in order that the epoxy resin polymerize. All containers (plastic beakers, petri dishes, etc.) which came in contact with the epoxy resin should also be put in the oven and be allowed to polymerize for safe disposal. Vials which held the 1:1 propylene oxide/resin mixture should be left open under a fume hood for a number of hours to overnight to allow for the evaporation of the propylene oxide (never put propylene oxide near a heat source/oven). The vials can later be placed in the ovens to polymerize the epoxy resin and finally discarded. After 48 hours, the BEEM capsules can be removed from the ovens and stored at room temperature. The tissues are ready for trimming and ultrathin sectioning. ✥ Tissue Processing Note From the mincing of tissues in the primary fixative and up to the point of vacuum infiltration, tissue blocks are handled and contained in small glass vials with plastic snapcaps (8ml capacity). Chemical agents are simply decanted using a Pasteur pipette and the replacement agent added with a different pipette. Do not allow your tissues to dry down during processing. This will lead to denaturation and artifacts. When decanting, always leave a small amount of the original solution to cover the tissue blocks, especially when decanting the highly volatile ethanol (especially 100%), acetone and propylene oxide.

20 ELECTRON MICROSCOPY: A HANDBOOK OF TECHNIQUES FOR THE BIOLOGIST

✥ Embedding Media In order for electrons to pass through a sample in the TEM, it must be reduced in thickness to between 600-900Å. Therefore, the sample must be sectioned ultrathin. Ultrathin sectioning would be impossible unless the biological soft tissues we study are strengthened to withstand the incredible forces encountered during sectioning. Unless a suitable embedding media is used, the soft tissues would simply disintegrate at the cutting edge along with the fine structures we wish to see. An adequate embedding media for TEM samples must meet certain requirements which follow: 1. consistency - a formulation must yield the same results for each mix. 2. availability - components must be readily available. 3. purity - components must be identified and characterized to avoid artifacts. 4. solubility - in common solvents. 5. miscibility - with other embedding media/curing agents/dehydrating agents (e.g. alcohol). 6. viscosity - low is ideal from a standpoint of convenience. 7. polymerization - must be controllable, uniform, and occur in a reasonable amount of time. 8. transparent - to light (to view tissue in blocks and sections) and electrons. 9. stability - under the electron beam. 10. sublimation - avoids liquid interface and resultant surface tension forces. 11. cross-linked polymer - for good ultrastructural preservation / strengthens tissues. 12. stainability - allows for heavy metal (TEM), pigment (LM) and histochemical staining. 13. no shrinkage - little volume change during polymerization. 14. stores well - long shelf life (some preliminary mixtures can be stored in the freezer). In general, resin components should be mixed thoroughly to avoid uneven polymerization and embedment. Hand mixing with a clean glass rod for 15-20 minutes is usually sufficient. Of course, this mixing will introduce a great deal of air to the mixture. Since the mixture is viscous and lengthy staining times may cause separation of the components, a low vacuum (bell jar / rotary pump) setup can be used to eliminate gas from the mixture. Such a setup is best run under a fume hood to avoid resin fumes. Another important consideration is that the hardness of the resin should closely match the hardness of the tissue in question. This will be considered in the discussions of the various embedding media which follows.

UNIT 1 – PREPARATION OF BIOLOGICAL SAMPLES FOR TEM

21

• Methacrylates The first embedding media to provide good results for the embedment and sectioning of TEM samples were the methacrylates which were introduced by Newman, et al. in 1949. These resins are low in viscosity and their hardness can be adjusted by varying proportions of methyl versus butyl methacrylate. The catalyst typically used to polymerize the methacrylates is benzoyl peroxide. Initially, they were hailed as the ideal TEM embedding agent until it was demonstrated that they polymerize unevenly. Since they are linear polymers, it was noted that they shrink nearly 20% during the polymerization process. This leads to a flowing of cellular components and gross distortions known as “explosion artifacts” due to the presence of large, seemingly vacuolated regions. In addition, the methacrylates are very unstable under the electron beam. This clearing of the resin produces an increase in contrast (since only tissue components are left behind). However, due to surface tension which arises at the liquid resin/tissue interface and the flowing and subsequent collapse of cellular structures, distortion artifacts are a certainty. Formvar (plastic) coated grids were typically used to support methacrylate sections. • Epoxy Resins Introduced in Denmark for TEM by Maaløe and Birch-Anderson (1956) and later in England by Glauert, et al. (1956), the epoxy resins are and continue to be the best embedding media for biological samples. The first quality epoxy resin developed for use in TEM sample embedments was “Araldite”, an extremely viscous compound which was used for many years in England by model makers. Later, the epoxy resin known as “Epon” 812 was developed and eventually marketed by the Shell Oil company. Today, many “812” epoxy resins are available from EM supply companies (PolyBed 812-Polysciences, Inc., EM Bed 812-EMS, Inc., etc.). As discussed earlier, in this lab, we use an Epon-Araldite mixture which is suitable for a variety of tissues. Epoxy resins are transparent, yellowish, cross-linked, thermosetting (with the application of heat) polymers which are highly viscous in their unpolymerized form. The modern epoxy is a diepoxy which has an epoxy group at each end of the molecule. The epoxy group is a strained, three-membered ring (C-O-C) which when ruptured, provides the energy 22Kcal/ mol) to drive the polymerization forward. Spaced along the organic molecular chain between the two epoxy groups are 1-5 hydroxyl (-OH) groups. Both the epoxy groups and the hydroxyl groups react with a variety of “curing agents” and either heat or ultra-violet (UV) light to yield the three-dimensional, cross-linked polymer. The final mechanical properties of the embedment are based on the type of epoxy resin and curing agents used. The epoxy group reacts with any reactive hydrogen atom which leads to its rupture and subsequent release of energy to drive the polymerization. A good source of reactive hydrogen atoms is the organic molecule class known as tertiary amines. The tertiary amines such as Benzyldimethylamine (BDMA) and Tridimethyl amino methyl phenol (DMP-30) are also known as catalysts or accelerators and are responsible for linear (end-to-end) polymerization. They should be added last and just before use of the epoxy mixture. Another class of organic molecules known as acid anhydrides react with the hydroxyl groups of the epoxy resin molecule to provide cross-linking. Acid anhydrides such

22 ELECTRON MICROSCOPY: A HANDBOOK OF TECHNIQUES FOR THE BIOLOGIST

as Dodecenyl succinic anhydride (DDSA) and Nadic methyl anhydride (NMA) are also known as hardeners. The mixture of epoxy resin (Epon/Araldite), acid anhydride hardener (DDSA), tertiary amine catalyst (DMP-30) and heat (or UV) yields a strong three-dimensional polymer which is resistant to solvents and heat. The three-dimensional polymerization provides excellent mechanical strength for ultrathin sectioning without shrinkage and resultant explosion artifacts. The ultrathin sections are extremely stable under the electron beam, so much so that uncoated grids can be used to pick up the sections. As mentioned earlier, it is important to match resin and tissue hardness. Hard tissue embedded in a soft resin would break out of the resin block. Soft tissue in a hard resin would be distorted or disintegrate as it is moved across the cutting edge. Much of this matching is a trial-and-error process of working with different resins and curing agents. By example, when using Araldite with the hardener NMA, the resultant polymer is too hard and brittle. When using DDSA, the block is still too hard for biological tissues. In this case a modifier such as Dibutyl phthalate (DBP) is used. This compound is nonreactive, however, it increases elasticity to produce a softer embedment. DPB is also known as a “plasticizer”. In another example, when DDSA is used with Epon, the block is too soft. Conversely, when NMA is used with Epon, the block is too hard. This situation is ideal since final hardness can be adjusted by varying the proportions of DDSA and NMA. One should be reminded that most epoxy resins are immiscible with water and that complete dehydration must be carried out prior to the introduction of epoxy. Any water in the tissue will result in a poor embedment. The transitional solvent, propylene oxide, aids in the infiltration of epoxy resins into the tissue and its use is recommended when possible. • Other Embedding Media Other types of embedding media for TEM samples exists. Polyester resins such as Vestopal W and Rigolac provide an improvement over methacrylates since they are not subject to shrinkage due to three-dimensional polymerization. They are polymerized by heat, light and oxygen and should be protected from such sources by keeping mixtures refrigerated in the dark. Another useful resin mixture is known as Spurr’s resin (vinylcyclohexene dioxide-VCD) which is a low viscosity resin ideal when infiltration of the sample may be difficult. Botanical samples are often embedded with Spurr’s resin since the cell wall may provide a barrier to the penetration of high viscosity resins. Water soluble resins are available such as Aquon which is prepared by extraction of the water soluble fraction of Epon. Aquon is completely miscible with water at 15˚C and below. Glycol methacrylate (GMA) is also relatively water soluble for sample preparation where dehydration would be impractical or detrimental to the tissue.

UNIT 1 – PREPARATION OF BIOLOGICAL SAMPLES FOR TEM

Fixation Schedule Mammalian Soft Tissue Protocol - In-situ and Immersion fixation - 0.5mm3 tissue blocks Initial Fixation at 4˚C - Ascending to Room Temperature

Schedule

Duration

3% Glutaraldehyde (0.2 M phosphate buffer, pH 7.4)

1 hour

Buffer Wash

3 x 10 minutes each

1% OsO4 (buffered as above)

1 hour

Buffer or DH2O Wash (optional)

2 x 10 minutes each

70% Ethanol 95% Ethanol 100% Ethanol (fill vials completely)

2 x 10 minutes each 2 x 10 minutes each 2 x 10 minutes each

Propylene Oxide

3 x 10 minutes each

1 Propylene Oxide : 1 Mixed Resin

1 hour

Vacuum Infiltration (in small petri dishes)

1 hour

Embed (in pure Epon/Araldite mixture) Place labels into BEEM capsules, then add degassed resin, then tissue blocks. Tissue blocks will descend to the tip of the EEM capsules before polymerization of resin.

48 hours at 60˚C

23

24 ELECTRON MICROSCOPY: A HANDBOOK OF TECHNIQUES FOR THE BIOLOGIST

Fixation Schedule Worksheet Mammalian Soft Tissue Protocol - In-situ and Immersion fixation - 0.5mm3 tissue blocks Initial Fixation at 4˚C - Ascending to Room Temperature

Schedule 3% Glutaraldehyde (0.2 M phosphate buffer, pH 7.4) Buffer Wash 1% OsO4 (buffered as above) Buffer or DH2O Wash (optional) 70% Ethanol 95% Ethanol 100% Ethanol (fill vials completely) Propylene Oxide 1 Propylene Oxide : 1 Mixed Resin Vacuum Infiltration (in small petri dishes) Embed (in pure Epon/Araldite mixture) Place labels into BEEM capsules, then add degassed resin, then tissue blocks. Tissue blocks will descend to the tip of the BEEM capsules before polymerization of resin.

Time In

Time Out

UNIT 1 – PREPARATION OF BIOLOGICAL SAMPLES FOR TEM

25

✥ Final Chemical Fixation Considerations Throughout the process of chemical fixation, the electron microscopist must be reminded that this is not a natural event and that the final result must be considered artifact. All images of fixed biological samples produced by the TEM are artifactual, not “true-to-life”. Does this mean that the cellular structures which have been previously described truly do not exist? Do double membrane bound structures known as mitochondria, which possess internal infolded cristae, really exist? Are they simply an artifact of preparation? One can answer those questions from an examination of the scientific literature. When you see a recurring feature which possesses the identical structural intricacies and occurs in regions of the cell with specific requirements, in this case energy, we can be confident as to the existence of this structure. At the same time, one must be extremely careful not to induce artifacts of preparation which can arise from improper handling of the samples. Poor technique throughout the entire fixation protocol will lead to glaring defects and artifacts when finally viewed under the high resolution of the TEM. Finally, while conducting a fixation protocol one must be cautioned that each step may involve your exposure to highly toxic chemical agents. Remember that fixatives kill and preserve living tissues, including yours! A healthy respect for all potentially hazardous chemicals should be developed by the student of science/biology. Use common sense in working with all chemicals! Assume that all chemicals you are unfamiliar with are hazardous and use appropriate precautions such as the use of disposable gloves (two pairs if necessary), goggles, and the fume hood. The ideal course of action is to consult the Material Safety Data Sheet (MSDS) for the chemicals you will be exposed to and follow the listed safety guidelines. MSDS should be available in any location that involves the handling of chemicals. Specific precautions will be found both in association with any chemical listed in any fixation protocol in this book. Whether a chemical agent has an immediate effect (contact dermatitis, blindness, death) or a latent and/or cumulative effect (cancer), the goal is to eliminate exposure of both yourself and future students to the agent. Be careful not to touch bare tabletops, door handles and knobs, etc. with contaminated gloves. You will be contaminating the work area for unsuspecting personnel that would come into bare-handed contact with these surfaces, possibly for years to come (especially in the case of resin components). Also, be aware of the location of emergency equipment in the lab, such as fire blankets, fire extinguishers, eye wash and shower station(s). The phrase “Look before you leap”, meaning, think before you do anything, is most appropriate when dealing with dangerous chemicals in the laboratory.

26 ELECTRON MICROSCOPY: A HANDBOOK OF TECHNIQUES FOR THE BIOLOGIST

Chapter 2 - Ultrathin Sectioning & Ultramicrotomy Unless the electron beam produced by the electron gun of the TEM can pass through the specimen (or transmit - hence the name Transmission Electron Microscope), achieving an image of high quality and resolution, or any image at all, is impossible. In order for the electron beam to pass through a sample, it must be sufficiently thin. This level of thinness required has been determined to be between 600-900Å, what is conventionally known as ultrathin. Since most samples, biological or otherwise, which one would consider viewing under the TEM are much thicker than this, a process known as ultrathin sectioning is required. It wasn’t until the early 1950’s that quality ultrathin sections were cut routinely. It is for this reason that biological TEM lagged behind the development and commercial availability of the TEM; the first commercial TEM was marketed by the Siemens corporation in 1939 ( an instrument designated the Elmiskop I). The precision instrument which was developed and makes it possible to cut ultrathin sections is the ultramicrotome. The following information covers the theory and use of the ultramicrotome along with the related requirements which are necessary for its use, including block trimming and glass knifemaking/diamond knife usage. ✥ Processing of Embedded Blocks - Block Trimming Prior to ultrathin sectioning, polymerized blocks must be trimmed freehand under a dissecting microscope. This process is conducted since tissue blocks rarely descend completely to the tip (truncated/flat face) of the BEEM capsule. This results in an embedded tissue block which is somewhere beneath the blockface. When you consider how little material is removed from the face through ultrathin sectioning (600-900Å), it obviously becomes necessary to expose the actual tissue sample at the surface of the block, otherwise, you will be sectioning epoxy resin only. Unlike in the preparation and sectioning of samples for the light microscope, TEM sections should be completely filled with tissue sample. In LM sectioning, a considerable portion of the section periphery is the embedding material, such as paraffin, with the actual tissue centrally located. Block trimming is a vital step which will ensure that your sections will contain a maximum amount of tissue. After considerable effort is taken to properly fix your tissue samples, careful block trimming is critical to success at the ultramicrotome. Block trimming is conducted entirely by hand under a dissecting microscope, preferably with a bright overhead light source. Since the process is entirely manual, many students find this to be one of the most difficult procedures to learn in TEM specimen preparation. The ideal setup is an older style Bausch & Lomb stereoscope with a “pod” head that is fit with 15X oculars and yields a maximum magnification of 45X. A bright external B&L illuminator is used in conjunction with the stereoscope, a setup which is far superior to a newer model B&L stereoscope with its less intense light source focused from overhead by a mirror.

UNIT 1 – PREPARATION OF BIOLOGICAL SAMPLES FOR TEM

27

Besides the microscope setup described above, block trimming also requires an embedded block (in a BEEM capsule), some type of chuck (such as a collet chuck) for holding the block, a trimming base or stage for supporting the chuck, a pair of pliers for removing the block from the BEEM capsule, a package of double-edged razor blades, a beaker of acetone (with a cover) to remove oil from the razor blades, Ross lens tissue to dry the acetone dipped blades, and a metric ruler in order to determine the dimensions of the trimmed blockface. It should be noted that although most individuals in the field do not recommend the use of double-edged blades, they are by far sharper than single-edged razor blades. The minor inconvenience of learning to use these highly flexible blades can be translated into much less difficulty at the ultramicrotome. The more rigid single-edged blade is not nearly as sharp and results in trimmed blocks with what we refer to as a “frosted” face and sides. These frosted areas appear smooth, however, they are actually irregular and can lead to sectioning problems such as sections which do not completely detach from the blockface as they are cut. Prior to block trimming, the polymerized block must be removed from the BEEM capsule. A simple way to proceed would be to slit the side of the capsule with a single-edged razor blade. If one is not very careful, this technique often results in numerous cut fingers. The recommended method involves using a pair of pliers to force the block out of the capsule using pressure. The BEEM capsule is first squeezed on its side which results in a noticeable separation of the resin from the capsule’s plastic. The pliers are then positioned at the capsule tip, just above the circular ring, superior to the truncated BEEM pyramid. Slow and even plier pressure is applied which results in the block moving outward from the capsule. Do not bring the jaws of the pliers together as you might damage the blockface. Instead, reposition the pliers slightly higher on the BEEM capsule and continue. Once this has been done enough, at least one-half of the block will project from the BEEM capsule. You should be able to twist and pull the block from the capsule at this point where it will be inserted about two-thirds of the way into a chuck. Empty BEEM capsules can be saved to place over the tip of a trimmed block and protect it until the time of sectioning. For the Sorvall-Porter Blum MT-2B ultramicrotome, collet chucks, with circular openings which are reduced in diameter upon tightening (as found in the design of a drill) are employed. It is best to load the blocks horizontally, from the side so that you don’t have to combat gravity. As stated earlier, blocks should be loaded about two-thirds of the way into the chuck. The threaded chuck is then screwed into the trimming base which is placed onto the stage of the stereoscope. The blockface is focused on at maximum magnification (45X) and brightest overhead illumination. The 1.0mm square blockface formed by the BEEM capsule is then set square in the field of view by rotating the trimming base. A double-edged razor blade is placed for a few seconds in a 50ml beaker of acetone. It is removed and wiped dry by pulling a sheet of Ross lens tissue away from the each edge. The blade should be held with the index fingers of each hand on top of the blade and the thumbs underneath. Fingers of each hand should be in close proximity with a slight pulling against each other to increase blade tension and rigidity. The other three fingers of

28 ELECTRON MICROSCOPY: A HANDBOOK OF TECHNIQUES FOR THE BIOLOGIST

each hand are anchored on the trimming base with the block and chuck between. Cutting should occur away from the individual, however, some cut the top (across the blockface) towards themselves with success. Initially, the blockface is cut down to the tissue. Thin sections are cut which are as parallel to the original blockface as possible. The blade should be held level or even better, at a very slight downward angle which ensures that the blade will dig in and produce a uniform and smooth new face. A slight blockface angle may result which can be compensated for on the ultramicrotome. A sharp blade results in a characteristic striated appearance on a reflective blockface. If “frosting” appears, the region of the blade being used has dulled and should be moved to a sharp area. For routine work, your goal is to trim the face down to the tissue block. You can be sure that the block has been reached when black material appears in your hand trimmed sections. This black material is your osmium fixed sample. At this point, your blockface is much larger than the original 1.0mm square and must be reduced by trimming the sides of the block. Sides should be trimmed parallel to the angle formed by the original BEEM capsule (~45˚) and not very deeply. The ideal pyramid is short and broad-based. Deep, steep sides leads to a poorly supported pyramid which vibrates when sectioned leading to a problem known as “chatter”. As the sides are reduced, so is the blockface. The ideal final dimensions of the blockface are 0.25mm on the longest side(s); not to exceed 0.5mm, with the longest sides parallel to each other. If the longest sides are not parallel, the ribbon of sections that come off the knife edge during sectioning will curve back to the edge instead of moving out in a straight line. A number of final blockface shapes can be trimmed such as a square, rectangle, and trapezoid. The trapezoid is the best shape to trim since it is most probable that sections of this shape will form what are known as ribbons. The following illustration (fig. 1) shows a block before and after trimming. Fig. 1 1.0mm 0.25mm

Tissue

Side View

Tissue

Block Trim 1.0mm

1.0mm

Tissue

Tissue 0.25mm

Top View

} Razor Blade Hand Trim Lines

UNIT 1 – PREPARATION OF BIOLOGICAL SAMPLES FOR TEM

29

In closing a discussion of block trimming, it must be noted that most ultramicrotomes can be used to trim blocks using glass knives since the knife stage and specimen arm angles can be manipulated through a wide range. Additionally, automatic block trimming devices and even gadgets to remove blocks from BEEM capsules are marketed. Although such instruments exist, for a price, most labs still trim blocks by hand. ✥ Glass Knife Making A suitable cutting edge for use in ultrathin sectioning is mandatory. Metal edges cannot be sharpened adequately to cut sections on the order of 600-900Å. The use of free fractured glass knives, introduced by Latta and Hartmann, was an important advance to the eventual production of ultrathin sections. The sharpest cutting edge that can be produced is that of free fractured glass. Even our ancestral cave dwellers knew the value of broken glass as they fashioned obsidian (volcanic glass) into tools for cutting. Today some surgeons will use obsidian scalpels to reduce “hamburgerization” of the integumentary (skin) tissues which can lead to scars. Only a handful of such courageous surgeons will use obsidian scalpels since they risk cutting their own valuable and skilled hands. In glass knifemaking (see fig. 2 on opposite page), plate glass strips are used. In this lab, LKB glass strips are used. LKB (now Leica) manufactures fully stress flown glass strips in which lines of stress are virtually eliminated. These high quality glass strips produce the sharpest eventual cutting edges. When a piece of glass is free fractured, it is precisely scored using a carbide scoring wheel or diamond tipped scribe. The score line must not run entirely across the length of glass you are trying to break, from one edge to the other. On the contrary, the score must be along the midline of the length of glass to be broken, at or near the center. Pressure is then applied beneath the score line and the glass free fractures to yield the sharpest cutting edge known. The simplest and least expensive way to apply this pressure is through the use of glazier’s pliers. The only problem here is inconsistency in the quality of the glass knives. In the early days of electron microscopy, one individual would be hired for the job of making glass knives. This person would become proficient at glass knifemaking through repetition and could routinely make quality edges for ultrathin sectioning. For many years, instruments have been available to somewhat automate the process of glass knifemaking. These instruments, such as the LKB 7800 series knifemakers, allow anyone to manufacture quality glass knives. The LKB knifemaker has provisions for precisely aligning, clamping, scoring and breaking glass strips of various dimensions. The most common dimensions of the glass strips purchased to make glass knives are 9-18" long x 1.0" wide x 1/4" deep. Shorter glass strips (9-10") are easier to handle and the LKB knifemaker can be used to cut longer strips in half. To cut a long strip in half, the strip is laid on top of the two spring loaded posts and the end of the strip is lined up even with the small black dot mark on the right-hand side of the machine’s surface. The strip is scored and broken as per the procedure for making a 1.0" square described below.

30 ELECTRON MICROSCOPY: A HANDBOOK OF TECHNIQUES FOR THE BIOLOGIST

Fig. 2 – Glass Knifemaking: 1/4 inch

1 inch

score line

free break path

1 inch

score line 1 inch 1 inch

frilling cutting edge conchoidal fracture plane

base (1 mm)

Boat/Trough Attachment: water level

tape seal

UNIT 1 – PREPARATION OF BIOLOGICAL SAMPLES FOR TEM

31

Before making a free fractured knives, the glass strip must be carefully cleaned in a nonfilming soap such as Liquinox. It should be held at one end only, the end which will not be used to make the knives, and thoroughly rinsed by holding the end used to make the knives upright. This prevents any soap on your hand which holds the glass from running down the strip and contaminating it. A thorough rinse in tap water followed by a final rinse in distilled water is recommended. The strip is then rested at an angle against lint free cloth, with the useful end upright, and allowed to air dry. Don’t wipe the glass dry, even with so-called lint free cloth, as you will produce a static charge on the glass and attract dust. Once dry, you can proceed with glass knifemaking on the LKB 7800. Initially, the LKB knifemaker should be set up with the breaking knob, the large black knob on the lower right of the machine which is used to apply pressure, fully counterclockwise. The clamping/scoring head should be elevated in place using the ball-ended clamping lever and the silver scoring adjuster should be on the “lines” setting (the three lines should be facing up - the other positions are 2.5 and 3.8). Holding the clean glass strip by one end, it is carefully lowered to the surface of the LKB 7800 with its scored/frilled edge down. The strip can be touched on the upper 1.0" wide surface but not on the 1/4" deep edge. Touching the strip on its surface, it is pulled down flush with the white plastic guide (which is set at 90˚) and slid to butt up against the first of two metal posts. The clamping/scoring head is lowered using the clamping lever until it contacts the glass strip surface. Upon contact of the clamping head, immediately release the glass strip surface with your hand. Failure to release with your hand at this point will result in the scored glass not breaking. Lower the clamping lever so that the ball end is just above the surface of the machine (finger width space). With the score adjuster set on “lines”, score the strip by smoothly pulling the white scoring lever out. When the scoring lever is pulled, a carbide wheel descends and ascends in an arc to avoid scoring the glass from edge to edge. This score is made perpendicular to the length of the strip, exactly 1.0" in from the end. The large black breaking/pressure knob is then turned slowly and consistently until the glass free breaks. When the glass breaks, the breaking knob is turned back full counter-clockwise and the clamping head is elevated using the clamping lever while supporting the head using the extended scoring lever. At this point, a separate 1.0" glass square has been produced. The remaining glass strip can be removed and placed on lint free cloth. The glass square can be slid to the top of the pair of black pressure feet and carefully rotated counter-clockwise into a diamond formation, by touching only the top surface of the square. The lower edge of the diamond should be slid between the two white plastic supports with the upper edge aligned across from the inverted “U” of the upper metal holder. The upper holder is brought into place by pushing in and rotating the small black control knob on the rear left side of the machine. The clamping head is lowered as before and the scoring adjuster is set to 2.5. Before scoring, the small black damper is moved to just touch the lower point of the diamond using the small metal lever and the fork-like knife holder is slid under the diamond (this can be done before scoring or after the knives break). The glass is scored at 2.5, once again not from edge to edge and this time slightly off the perpendicular, and broken using the slow and steady movement of the breaking

32 ELECTRON MICROSCOPY: A HANDBOOK OF TECHNIQUES FOR THE BIOLOGIST

knob. Once the glass breaks, the clamping head is elevated, the damper is reset (dot to thin line) and the upper holder is released by pulling and rotating the holder knob. The two knives will rest on the fork-like holder which can be removed. They should be left on the holder and carried to a dissecting microscope. Do not allow them to rest on their sides on a tabletop as the edge may pick up contamination. Caution: Never leave upright glass knives unattended. Unsuspecting individuals can come along and rest their hands down on top of the extremely sharp cutting instrument. A clean plastic (tri-pour) or glass beaker is recommended to cover and protect both your knives and colleagues. After preparation of the knives, they must be evaluated for quality. Obviously, poor knives should never be used for ultrathin sectioning. Handle the knives carefully so that the cutting edge will not become damaged or contaminated, especially with oil from your skin. Initial evaluation is performed by visual inspection of both knives without a microscope. From the side you will see the shape of a right triangle. The base of the knife will have a 1.0mm raised heel. The heel should be an even rectangle with its longest sides parallel. The cutting edge should be relatively straight and level, however, a convex or concave edge is not uncommon. The convex edge often has the longest usable cutting area. Looking directly at the angled side of the knife, you will notice the sloping conchoidal fracture plane. It is believed that the further to the right this fracture plane exists before descending, the longer the usable cutting edge will be. The only way to evaluate a glass knife edge’s quality is to examine it under a dissecting microscope with an overhead light source. The setup used for block trimming is ideal. For the best view, a dark background is recommended. In this case a small square of black photograph mounting board can be cut and placed on the microscope stage. The magnification should be adjusted to allow for viewing of the entire knife edge (~ 30X). The knife should be held with the angled edge facing outward and the knife set in the shape of a “V”. While looking under the stereomicroscope the glass should be rocked so as to concentrate the overhead light on the very edge of the knife. The knife edge should lie near the center of the field of view. When the knife edge is focused you should see a broken bright line on the right side of the cutting edge. The broken line is indicative of typical imperfections in the edge which is known as “frilling”. The left edge of the knife should appear as an unbroken bright line which indicates its sharpness and high quality for sectioning. In order for the knife to be usable, at least 1/3 to 1/2 of the left edge should be unfrilled. Knives with quality edges less than this should be discarded (in an appropriate sharps container to minimize risk to the maintenance staff). The knife edge can also be evaluated for debris which would lead to the appearance of sectioning artifacts. Do not try to clean the edge of debris using a duster can (compressed air) since particulate and organic matter in the can will damage the edge. Dirty knives must be discarded with the student exercising greater caution in the preparation of additional knives. Always prepare more than one good knife in the event that the knife is damaged or contaminated in transit to the ultramicrotome. Once at the ultramicrotome, you will not want to backtrack and make more knives. Once the knives are evaluated, they must be “boated” through the application of a tape trough/boat. This boat will hold floatation fluid, typically double distilled water, which will

UNIT 1 – PREPARATION OF BIOLOGICAL SAMPLES FOR TEM

33

allow the sections to come off the knife edge and float on a liquid surface as opposed to sticking to the dry glass edge. The boat is usually made of mylar adhesive tape and is available through some EM supply companies. A suitable substitute would be electrical tape strips which are cut in half (on a clean piece of glass) to reduce the width. Once a length of tape is cut, the knife glass is placed to it. You will have more control in touching the knife to the tape versus touching the tape to the knife. You should orient the knife to the tape in such a way that only a short length of free tape extends to the left of the placed knife. This will minimize the chance of the eventual two free sticky tape ends touching and transferring adhesive to the knife edge. In terms of orientation, the tape must be exactly even with the upper cutting edge. The lower edge of the tape must be parallel to the lower base of the knife. Once the knife is placed to the tape, the knife can be picked up and the long free tape end can be wrapped around to fashion a symmetrical boat having a small gap at the bottom (where the tape meets the angled edge of the knife). If the orientation is not correct, the tape can be carefully pulled off an another attempt can be made. The more attempts one makes, the greater the risk of contaminating the knife edge. Excess tape projecting from the back of the knife is cut off using a razor blade and being careful not to touch the knife edge. While boating the knife one should be careful not to touch the tape in the region which will actually form the boat. When water is introduced, oil from the skin will serve to contaminate the floatation fluid and ultimately, your sections. After the tape boat is attached, it must be sealed to prevent leakage using nailpolish (or molten paraffin wax). The polish should be applied all along the lower tape edge, especially at the gap on the angled edge of the knife. Excess polish should be removed from the brush on paper toweling and a thin line run up the two back edges of the knife. Do not apply polish to the knife edge. The polish is allowed to dry for about 20 minutes at which point, the knife can be used to section on the ultramicrotome. Since glass is a supercooled liquid, it tends to flow away from the sharp edge and become dull over time. Ideally the glass knife should be used within 24 hours of manufacture, however, if kept in a storage box and desiccator, it should be useful for a few days. Another problem with the glass knife is that it rapidly dulls with use. Only between 15-30 ultrathin sections may be cut before the appearance of visible knife marks which are noticeable dark, vertical, irregularly spaced lines in the section. These unmarked sections will be picked up followed by moving the knife to an unused cutting region. An alternative to the glass knife is the diamond knife described below. ✥ Diamond Knives Diamond knives are extremely delicate and expensive instruments which can cost as much as $5,000.00 based on the length of the edge. Obviously, only experienced microtomists should handle a diamond knife since simply touching the edge or cutting a section greater than 1.0µm may permanently damage it. The benefits to using a diamond lie in its hardness. Proper care results in a knife that is sharp for years and can cut numerous sections without having to stop and move the knife. The entire edge is sharp which results from taking a quality gemstone and cleaving it into smaller fragments which are polished

34 ELECTRON MICROSCOPY: A HANDBOOK OF TECHNIQUES FOR THE BIOLOGIST

using diamond dust (the actual process is a trade secret of the few companies which manufacture diamond knives). The diamond is mounted into a metal holder which is glued into a metal trough using epoxy. The need for making glass knives and attaching tape boats is eliminated. A common problem in using a diamond is that it is quite hydrophobic which prevents wetting of the knife edge. The remedy involves carefully running an eyelash along the inside edge of the diamond with the boat full. If the diamond still won’t wet, a bit of saliva or dilute wetting agent (such as Photoflo 200:1) can be used on the eyelash. Routine cleaning of the diamond knife involves a gentle stream of distilled water from a squirt bottle to clean the boat and knife edge. On occasion and if sections adhere to the edge, a cleaning stick made of styrofoam or pithwood can be run along the edge, in one direction only. Never apply forward or backward pressure on the edge. Cleaning sticks can be dipped in ethanol or even saliva to remove adherent material from the diamond edge. A recent addition to the types of knives used in ultrathin sectioning is the sapphire knife. As with the diamond knife, the sapphire is permanently mounted in a trough and is superior to the glass knife. It is however inferior in quality to the diamond knife and is incapable of cutting hard samples such as tooth and bone, without damage to the edge. ✥ Ultramicrotomy The development of the ultramicrotome has been critical to the formation of high quality images of biological material using the TEM. In order for the electron beam to pass through the specimen, it must be less than 1,000Å thick, ideally between 600-900Å. To produce sections thin enough for use in the TEM took years of research into mechanical engineering of the actual instrument in addition to fixation and embedding materials, blockface requirements and cutting instruments (glass and diamond knives). When TEM development was taking place in the 1930’s, its use for the examination of biological samples was not envisioned. Early investigators who were interested in examining biological samples with the TEM would attempt to use whole mounts of fractured materials with poor result. In 1939, von Ardenne proposed cutting tapering, wedge shape sections whereby a portion of the section would be thin enough for TEM observation. Modification of a Spencer 820 microtome, used for paraffin embedded LM level sections, by Pease and Baker in 1948 resulted in a 1/10 reduction in block advance to the cutting edge. They also reduced the block face to 1.0mm2 and produced sections 0.30.5µm thick. As detailed in the unit on embedding media, Newman et al. (1949) introduced methacrylates which became the first embedding media of quality for TEM samples. Using a metal knife which had been sharpened with a new technique, along with an attached trough which was used for the first time, Hillier and Gettner (1950) made further modifications to the Spencer 820 microtome and produced sections 0.2µm thick. A major advance in the field took place in 1950 with the introduction of glass fracture knives by Latta and Hartman. In 1952, Palade used methacrylate embedding, glass knives, a

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blockface smaller than 1.0mm2 on an ultramicrotome designed by Claude and Blum and was able to obtain sections at the useful thickness of 1,000Å. In 1953, the Sorvall “PorterBlum” MT-1 ultramicrotome was introduced commercially. This instrument was handdriven and allowed for reproducible ultrathin sectioning by anyone who had the patience to learn its operation. Needless to say, the first quality TEM photomicrographs of sectioned biological material appeared at this time. Since the introduction of the MT-1, numerous improvements in design lead to the development of the Sorvall MT-2 and 2B with motorized drive, and eventually the MT9000, currently available from RMC, Inc. LKB (now Leica) was the first company to introduce an ultramicrotome using the advance principle of thermal expansion. The LKB Ultrotome series also had superior lighting and improved specimen blockface to knife adjustments for greater control in orienting knife to blockface. Current state-of-the-art instruments include the Leica Ultracut with fiber optic blockface illumination which make it almost impossible to chop off your blockface, a common problem of the inexperienced microtomist. Other advances in the field of ultramicrotomy include the work of Peachey (1958) who suggested that interference colors of sections under a cold (fluorescent) light source can provide a good estimate of section thickness. He determined that the following section colors relate to their thickness:

60 nm 60-90 nm 90-150 nm 150-190 nm 190-240 nm 240-280 nm 280-320 nm

gray silver gold purple blue green yellow

Ultramicrotome Theory In the design of an ultramicrotome (fig. 3), a rigid cantilever arm is employed to which a trimmed block can be mounted. The trimmed block is held within a chuck. The most common type is a collet which tightens down on the block by turning the threaded collar, usually clockwise, leading to a reduction in the diameter of the circular opening. Other chucks, such as those used in early LKB ultramicrotomes, resemble nosecones which are split in half lengthwise and held together by a hexagonal set screw. Vise-type chucks are also available with a pair of adjustable flat jaws to accommodate flat embedments. It is imperative that the chuck holds the block securely to avoid vibration during sectioning. The ultramicrotome cantilever arm must be able to move in three degrees of freedom about various pivot points. Firstly, the arm must be able to move up and down in an arc; this is known as the cutting stroke since downward movement through a cutting edge

36 ELECTRON MICROSCOPY: A HANDBOOK OF TECHNIQUES FOR THE BIOLOGIST

will yield a section. Secondly, the arm must be able to move from side to side in an arc; this is known as the bypass stroke since it allows the blockface to bypass the knife as it is moved to the top of the cutting stroke. Without this motion, the blockface would be damaged through compression as it hit the back of the knife. In the early Sorvall “PorterBlum” MT-1, the arm moved within a parallelogram shaped cutout. Modern instruments such as the MT-2 eliminate the side to side motion in favor of a retraction of the arm at the bottom of the cutting stroke. In the LKB/Leica instruments, the knife stage is retracted electromagnetically. Either way, contact between the blockface and the back of the knife is avoided. Finally, the arm must be able to extend linearly along its axis; this is referred to as the advance since it allows the arm to advance the required minuscule amount to the stationary cutting edge. The amount of advance will determine the ultimate section thickness. In the development of the ultramicrotome, two methods of advance have been used. The first, used in the Sorvall MT-1 and MT-2(B), is mechanical which is designed around a vertical pivoting arm which rests on a micrometer lead screw. As the lead screw turns, the lower portion of the vertical arm moves to the back of the ultramicrotome, while the upper portion advances out toward the front of the machine. Since the rigid cantilever arm is attached to the upper portion of this pivot arm, it will be moved forward toward the knife edge. The vertical pivot arm will eventually reach an end point and will have to be reset. One should be sure to reset the mechanical advance machine before mounting a block. The second type of advance used is thermal which involves the uniform expansion of a metal under uniform heating. This method, used in LKB (now Leica) Ultrotomes for example, must have provisions for variable heating to control section thickness and a cooling fan in order to “reset” the mechanism, since there is a limit of thermal expansion for any given metal. The arm must be motor driven and electronically controlled. The mechanism must be able to compensate for changes in cutting speed while maintaining section thickness. By example, if silver sections are cut at 1.0mm/sec and you change the cutting speed to 2.0mm/sec, the machine will have to introduce a point of hesitation into the cycle to continue to yield silver sections. Without this hesitation, the cycle time would be shortened, therefore, heating time reduced leading to a thinner section. In terms of cutting speed, the best results are obtained if the cutting speed is uniform (usually somewhere between 1.0-2.0mm/second). Variations in cutting speed was a common problem in producing good sections using the hand-driven Sorvall MT-1. Many labs designed a reduction gear to attach to the cycling wheel leading to greater uniformity in cutting speed. Whether mechanical or thermal advance, the introduction of motor drives to the ultramicrotome was a vast improvement. Another important feature of the ultramicrotome is the stage to which the cutting instrument/knife is attached. In some cases the stage is permanently mounted to the ultramicrotome (LKB Ultrotome III) while others are removable (Sorvall MT-2). The stage must be finely machined which will allow for fine control in bringing the knife to the blockface. The stage must incorporate a knife holder (for both glass and diamond knives) and provision for setting the clearance angle, the most important angle of the knife

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relative to the blockface, which is initially set at 4˚. There is usually a device/jig to set the proper height of the knife edge in the stage. The stage will also incorporate control for the lateral movement of the knife to select the appropriate cutting area (knife edge adjustment), rotation of the stage to allow for machine block trimming and most importantly, to establish that the lower edge of the blockface be parallel to the knife edge before cutting (knife rotation). If not parallel, one side of the blockface will be cut before the other. Many thick sections would have to be taken to yield a full blockface which would probably be too large for ultrathin sectioning. The final provision is the stage advance (knife advance) with both coarse and fine adjustments. This control is critical since the knife must be manually moved to the blockface for the first facing cut. Since the ultramicrotome arm advances in such small increments, it cannot be expected that the arm would ever reach a stationary knife located some distance from the blockface. Additionally, the cantilever arms are not capable of traveling large enough distances to reach the knife. The knife must be moved to the blockface using the fine controls of the stage. A common problem for the beginner is their lack of patience in advancing the stage to the blockface. The result is a chopped blockface which must be retrimmed. It should be noted that each control on the stage will have a lock to reduce vibrations. Vibrations can lead to a sectioning artifact known as chatter. Chatter is a regular variation in thick and thin areas within a section. This produces a uniform dark and light banding pattern on the section. Fine order chatter, which cannot be seen under the light microscope, ruins the section by obscuring details. Reduction of vibration is a major concern to the microtomist. This can be accomplished by making sure all locks are engaged (chuck, stage, etc.), that the block trimmed is short and broad-based, and that the table which supports the ultramicrotome is sturdy (special anti-vibration tables are available but may be unnecessary depending on the room used for sectioning). In general, ultramicrotomes incorporate a massive baseplate in order to lessen the effects of vibration. Another important feature of the ultramicrotome is lighting. As described earlier, a fluorescent light source is used to produce section interference colors for thickness determination. Good lighting is also important for the critical advance and initial alignment of knife edge to blockface. It is recommended that the position of the light be adjusted regularly during advance of the knife to the blockface for the best possible view. Modern instruments incorporate fiber optic lighting on the blockface which leads to a more accurate approach and fewer chopped blockfaces.

38 ELECTRON MICROSCOPY: A HANDBOOK OF TECHNIQUES FOR THE BIOLOGIST

Fig. 3 Collet Chuck

Block

Water Level

Clearance Angle (2-5°)

Block

Microtome Arm

Glass Knife

Collet Chuck

Microtome Arm

Knife Edge

Ultra-thin Sections in Ribbon

Water Surface

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39

MT-2B Ultramicrotome Sectioning Procedure The following describes the procedure to follow when sectioning with the Sorvall “PorterBlum” MT-2B ultramicrotome. Many of the general steps can be applied to the proper use of any microtome on the market with a noted improvement in lighting over the MT series. Failure to adhere to the procedure as outlined may result in frustration and many chopped blockfaces. INITIAL SETTINGS & PROCEDURES • Press Reset Button • Upper Thickness Pivot Knob at 10 (10 x 10Å = 100Å) • Thickness Wheel (front left side) at 12-14 (1200-1400Å) • Cutting Speed at 1.0mm/sec • STAGE: Fully Retracted with ALL LOCKS OFF - Do NOT Place on MT-2B at this time! THICK SECTIONING PROCEDURE • Mount trimmed block and orient with fluorescent light fully extended out! Firstly, obtain a fully reflective blockface using loosened “arc” adjustment and hand tighten thumbscrew (B&L binocs set at highest magnification). Secondly, orient bottom of blockface parallel to “ground” using chuck rotation and hand tighten (This adjustment is not critical since knife edge parallel can be obtained using stage knife rotation). Use tools to completely tighten chuck (NEVER OVERTIGHTEN ANYTHING). • Mount Knife in Holder (Be sure front of knife is flush with aluminum guide plate). • Mount Knife Holder in Stage and Lock at 4˚. • Push Light Source back in and Mount Stage to MT-2B (use stage rotation lock on left). • Stage should be pushed forward completely until it stops. • Check Position and Duration and set if necessary (observe from side of setup). • Using lowest magnification setting on B&L binocs, locate block pyramid and knife edge. • Focus on knife edge and bring pyramid into focus by rotating cycling wheel clockwise. THIS MUST BE DONE OFTEN OR YOU WILL CHOP YOUR BLOCKFACE !!! • Coarse advance the stage (knife) to the blockface. Increase the binocs mag as blockface and knife edge get closer. Always refocus on the knife edge, THEN, bring the blockface into focus by cycling (focus the “cutting” stroke and not the “retraction” stroke). • Your goal is to get the knife edge and blockface in focus together in the same field of view at the highest binocs mag (“3” = 45X). • Position your knife to face the block near the center of the knife edge and gently LOCK. • Advance closer using the coarse control until you feel you might chop your blockface. • After cycling the blockface to the lowest point in the cycle, fill your boat with clean distilled water using a drawn micropipette or syringe, to a slight positive meniscus. Level and clean the water surface using Ross Lens Tissue (the water should be of a uniform reflective gray without any dark shadows near the knife edge).

40 ELECTRON MICROSCOPY: A HANDBOOK OF TECHNIQUES FOR THE BIOLOGIST

• Lock out coarse advance and go to fine advance using thumbscrew (NEVER TIGHTEN SCREW WITH STAGE IN FULLY RETRACTED POSITION). Only the large diameter advance barrel should turn with each line equal to 1.0µm increments. • Critically focus on the knife edge and cycle the blockface back into sharp focus. • Fine advance without cycling until extremely close to knife edge. Determine if bottom of blockface is parallel to knife edge. If not, carefully rotate stage until corrected and gently LOCK stage rotation. • When critically close, begin to interleaf cycling with fine advance with a final reduction in advance to 1.0µm per cycle prior to your first section coming off. • You may note a light reflection of the knife on the blockface (see diagram). The reflection (best seen in a faced block) indicates you are at least 10.0µm from the face. The reflection will shrink and finally disappear when you are 10.0µm from the face:

• Your goal is to take 4-6 full face thick sections (1.0µm green) by manual cycling. • With the blockface at the lowest point in the cycle, pick up some thicks using your eyelash and place them in a drop of water at the center of a clean microscope slide. • Allow to air dry and observe using the phase contrast microscope for the presence and quality of tissue. • Proceed to thin section if all checks out. ULTRA-THIN SECTIONING PROCEDURE • With block in lowest point of cycle, refill and clean and level boat (Ross Tissue). • Retract with fine advance (50µm at this step and 10-20µm thereafter). • Cycle blockface into critical focus with knife edge (should see reflection). • Unlock and move knife edge just inside of quality area of knife as was determined by prior evaluation. Gently RELOCK. • Fine advance until reflection disappears then with cycling and 1.0µm advance until the first section is cut. • Lock the stage advance and START the MOTOR (depress red button). • Note section colors (probably purple) and reduce thickness using side (NOT PIVOT) thickness wheel until PALE GOLD/SILVER is obtained. WALK AWAY - LET MACHINE DO ITS JOB !!! • After 15-20 sections are cut or until noticeable vertical knife marks appear, STOP MOTOR. • Isolate floating sections using the eyelash and pick up on the dull side of 200-400 mesh GRIDS (use 200 mesh for ribbons, 300-400 mesh for loose sections (undesirable). Hold slightly bent grid in jeweler’s forceps and touch dull side down on top of floating sections. Turn grid dull side up and blot dry on filter paper. Unlock forceps and place grid on grid gripper or on edge in grid box.

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• Follow ultra-thin sectioning instructions above and obtain more silver sections. REMEMBER TO UNLOCK THE STAGE ADVANCE LOCK BEFORE RETRACTING THE STAGE AND MOVING THE KNIFE FURTHER LEFT !!! BE PATIENT & GOOD LUCK !! Final Ultramicrotomy Comments Some additional comments are warranted with regard to the MT-2B procedure which was just outlined. Under “initial settings and procedures”, the required steps should be obvious. Recommended initial cutting speeds are usually in the 1.0 to 2.0mm/sec range. The stage should be carefully checked to ensure that it is fully retracted otherwise, you could easily ram the knife into the blockface when the stage is first mounted to the ultramicrotome. Under “thick sectioning procedure”, the initial blockface orientation is critical. The goal is to obtain an initial full-face thick section manually. This is impossible unless the blockface is properly oriented. In order to accomplish correct orientation, the proper lighting on the face is important. With the MT series, the fluorescent light must be fully extended so that the blockface is illuminated from the front. After blockface orientation, the light must be placed in a rear position in preparation for advancing the knife to the face. When mounting the knife in its holder, be careful not to break the tape/nail polish seal. Make sure that the knife seats on top of the height adjusting set screw and not behind it. If the knife is not flush with the front aluminum guide plate, the clearance angle you set will be inaccurate. A feature of the MT-2B is position and duration controls (the MT-2 did not have these controls). They provide fine control over the ultramicrotome’s cutting range and are best demonstrated at a slow cutting speed (0.33mm/sec). At this slow speed you will notice the actual cutting range of the instrument. The actual range exists over the position and duration of this slow rate of cantilever arm travel. Above and below the cutting range, the arm moves faster. Obviously, you want the cutting range to coincide with the knife edge. Looking from the side of the stage/block-arm setup, one must ensure that the slower range of blockface/pyramid travel is coincidental with the knife edge. If not, the position of the cutting range can be raised or lowered and/or the duration (length of cutting range) can be lengthened or shortened. Initial facing of the block should be done at or slightly to the right of the center of the knife edge. The left side should be reserved for ultrathin sectioning. When filling the boat, it is important that the blockface be positioned at the lowest point of the cycle. The indicator of a proper water level is a uniform reflective silver-gray surface appearance. If the block is at a high point in the cycle, it will cast a shadow on the water surface and you will be unable to determine the proper level. Relative to trough fluids, double distilled water in a sterile centrifuge tube works best. Some investigators use a low concentration of 1-3% acetone or ethanol to facilitate wetting of the knife edge. Although rarely seen with glass knives, failure of distilled water to wet the diamond knife edge is common. Carefully

42 ELECTRON MICROSCOPY: A HANDBOOK OF TECHNIQUES FOR THE BIOLOGIST

running an eyelash along the inside edge of the diamond is often a solution, especially if the eyelash has been dipped in a wetting agent (Kodak photoflo) or even saliva. Only experience and practice will enable you to determine when to begin fine stage advance and ultimate interleafing of advance with cycling in order to avoid ramming your blockface. The reflection of the knife edge in the blockface is the best indicator of distance, however, it is often difficult to see in a hand trimmed blockface. Once thick sections are obtained, picking them up becomes difficult for many beginners. An eyelash which is brought up quickly from underneath a group of sections works well. If the eyelash proves too difficult, a syringe tip or shaved toothpick can be tried. The toothpick should be dipped in acetone or you may introduce a large amount of debris into the boat. The thick sections will be used to verify the presence of tissue in the section along with its quality and orientation. Under “ultrathin sectioning procedure”, you must be sure to first unlock the stage advance and then retract the stage before moving the knife to section in a new area. The knife/ stage will be fine advanced with a good reflection of the knife edge in the faced block. The reflection will also aid in the determination of parallel relative to the knife edge and bottom edge of blockface. Once the first semi-thick section is cut, the stage advance lock must be engaged to prevent chatter. Once the motor is started, ultrathin sectioning will begin. Reduce the side thickness control until silver sections are obtained. If the block was properly trimmed, a ribboning of sections will occur. Once knife marks appear, the motor is stopped with the blockface at the lowest point in the cycle and the sections are picked up on the dull side of a grid. Pickup is easily accomplished by simply touching the dull side of the grid to the floating sections. To protect the tape boat seal, a bend is put in the edge of the grid (do not attempt to bend formvar coated grids). With the edge of the grid locked into the jeweler’s forceps, it is laid flat, dull side down, on filter paper. The forceps are then raised to create an approximate 45˚ angle with the filter paper surface. The bent grid held by the forceps is put aside and the eyelash is used to separate long ribbons into ribbons of 15 to 20 sections and isolate them in the center of the boat (isolated silver sections can also be rounded up at the center of the boat if ribbons are not formed). The grid is picked up and using the unaided eye, positioned over the boat in a flat, horizontal orientation. Looking through the binocs, the grid can be arranged so that the sections/ribbon will be centrally located after pickup. The grid is gently lowered to the sections so as not to disrupt the surface tension forces and taken away. Do not submerge the grids! The dull side of the grid is used for a number of reasons. Since the dull side is rough in comparison to the polished (shiny) side and the sections themselves are rough, they adhere better to the dull side of the grid. Another advantage is that you establish uniformity and always know what side your sections are on and on the dull side, you can actually observe the reflective sections after you collect them (fig. 4). After sections are collected, grids are blotted dry on edge (see post-staining procedureChapter 3) and stored dull side up on a “grid gripper” (also known as a grid mat) to air dry, after which they can be post-stained. The grid gripper prevents grids from attaching to the lid of the petri dish via static electricity. The safest place to store grids is on edge in

UNIT 1 – PREPARATION OF BIOLOGICAL SAMPLES FOR TEM

a grid box. Serial sections collected on slot or hole grids should never be put on a grid gripper or blotted flat down on filter paper. Fig. 4

43

44 ELECTRON MICROSCOPY: A HANDBOOK OF TECHNIQUES FOR THE BIOLOGIST

Troubleshooting Guide to Ultramicrotomy When an instrument capable of cutting sections on the order of 600-900Å is used, it becomes obvious that a number of problems may be encountered. The following section provides information on troubleshooting the most common problems associated with ultrathin sectioning. Although at times this process can be quite infuriating and frustrating, the key to mastery of the ultramicrotome is practice. The more problems encountered at the outset, the better equipped one becomes at finding solutions. Students may initially get lucky and achieve good results on their first attempt but it is unlikely that they will be able to produce consistent quality results unless they encounter some of the problems which are described below. • Inability to cut any sections: a. microtome advance not reset b. specimen/blockface too large c. poor embedding or soft block d. poor knife (not evaluated correctly) or improper clearance angle setting e. water level in trough too low f. vibrations in room g. blockface gets wet due to high water level h. faulty ultramicrotome (check that all locks are tight) • Variations in section thickness from one section to another a. blunt/dull knife edge b. incorrect clearance angle (usually needs to be increased) c. incorrect cutting speed d. soft block e. drafts or temperature changes in the room f. faulty ultramicrotome (check that all locks are tight) • Knife marks a. imperfections or contamination of the knife edge - knife poorly evaluated • Chatter a. resin and tissue hardness are not properly matched b. poorly trimmed block - excessively high/tall, not well supported c. locks not engaged - looseness leads to vibration d. cutting speed may be too high e. clearance angle may be too high f. blockface too large (approaching 1.0 sq. mm)

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• Compression (can use organic vapors of chloroform or heat to flatten sections) a. cutting speed too high b. incorrect clearance angle c. poor knife edge d. soft block • Folded/Wrinkled sections a. poor knife edge b. cutting speed too high c. water level too low d. soft block e. incorrect clearance angle • Sections pulled down back of knife a. water level too high b. cutting speed too slow c. clearance angle too small • Ribbons not straight or not formed a. poorly trimmed block b. two sides of blockface parallel to knife edge are not parallel to each other c. water level in trough not correct d. incorrect cutting speed e. blunt/dull knife • Holes in sections a. poor infiltration of resin into tissue b. air bubbles in resin/tissue - vacuum infiltration is recommended to avoid this • Cannot see sections as they come off the knife edge/cannot achieve proper trough level a. adjust/move illumination b. blockface not at lowest point in cycle c. leaky boat seal It becomes obvious from a glance at these troubleshooting tips that many problems can be solved by manipulating cutting speed and/or knife clearance angle. LKB/Leica has summarized a methodical approach to varying these parameters. They suggest starting at 4˚ and a speed of 2mm/sec. If the results are unsatisfactory, they recommend changing the speed to 1mm/sec, then 5mm/sec. If good results are still not obtained, a change in clearance angle to 1˚ is recommended (don’t forget to retract your knife sufficiently) starting once again at 2mm/sec and moving to 1mm/sec, then 5mm/sec. A final step would involve changing the clearance angle to 7˚ using the identical cutting speeds as previously noted.

46 ELECTRON MICROSCOPY: A HANDBOOK OF TECHNIQUES FOR THE BIOLOGIST

At all steps of ultrathin sectioning, contamination is to be avoided. The most common problems arise when oil is not removed from razor blades used in trimming and glass strips are not adequately cleaned with a non-filming soap and rinsed well enough. Additionally, as glass knives are prepared they become contaminated from oily fingers and trough tape adhesive. Trough fluids must be pure double distilled water and one must be careful not to contaminate the fluid by introducing dirty instruments such as grids, forceps and eyelashes into the boat. If the trough fluid is dirty, section contamination is inevitable. Once sections are collected on grids, they must be stored in a dry, dust free environment such as a grid box. Care should be taken not to drop the grids. Keep a layer of Ross lens tissue under the work area as you handle the grids. If you should happen to drop the grid, at least it will be onto a non-linting paper. Be especially careful when poststaining which is a common source of section contamination. Materials Required for Ultrathin Sectioning The following is a complete list of all the materials required in the EM lab to conduct ultrathin sectioning and all related procedures such as block trimming and glass knifemaking. • Block Trimming a. Properly embedded tissue within resin block b. Small (50ml) beaker containing acetone for the removal of razor blade oil c. Double edged razor blades (cleaned with acetone prior to use) d. Ross lens tissue (used to dry and clean blades before and during use) e. Microtome chuck (collet, etc.) to hold block f. Trimming base plate for supporting the chuck while trimming g. Pliers (to squeeze block out of BEEM capsule) h. Small metric ruler to estimate size of trimmed blockface i. Dissecting microscope with overhead illuminator (such as Bausch & Lomb) • Glass Knifemaking a. Plate glass strips (LKB/Leica recommended) b. Liquid soap (non-filming such as Liquinox) c. Distilled water for final rinsing d. LKB/Leica Knifemaker (7800 series or more recent model) e. Dissecting microscope with overhead illuminator for knife edge evaluation f. Mylar boat tape (or strips of electrical tape which are cut in half lengthwise after attachment to a clean strip of plate glass) g. Nail polish for sealing the boats h. Razor blade for cutting excess boat tape

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• Ultramicrotomy a. Trimmed block (0.25 - 0.5mm on the longest side with two sides parallel) b. Glass fracture knives with attached and sealed boats (2-4 knive are recommended with at least 1/3 to 1/2 quality cutting edge on left side as previously evaluated) c. Ultramicrotome (complete with microscope head, stage, glass knife holder, cold light source, and any required accessories) d. Double distilled water in sterile centrifuge tube (place in test tube rack) e. Clean micropipette or tuberculin syringe to fill boat f. Eyelash to pick up thick sections and to orient thin sections in boat g. Ross lens tissue (to clean and regulate meniscus level in boat) h. Acetone cleaned 200, 300, 400 mesh grids (on clean filter paper in a covered petri dish) i. Jeweler’s forceps with an o-ring lock j. Filter paper in covered petri dish to blot grid dry after section collection k. Grid gripper in covered petri dish for temporary storage of grids (quality grids should later be moved to a grid box for permanent safe storage)

48 ELECTRON MICROSCOPY: A HANDBOOK OF TECHNIQUES FOR THE BIOLOGIST

Chapter 3 - Post-Staining In order to form an image, enhancement of contrast (differences in optical or grain density) is necessary, especially when an ultrathin section is considered. In light microscopy, colored dyes/stains are utilized in order to improve contrast and impart color differences. In the preparation of biological samples for the TEM, electron dense, heavy metal stains are used. The most common staining employed is double post-staining, meaning that two stains are used after the sections are collected on grids. Another technique which is readily found in the literature involves en bloc staining of tissue blocks before the blocks are embedded in the epoxy resin(s). Typically, tissue blocks are stained en bloc using aqueous 0.25% to 4% uranyl acetate solutions for 5-60 minutes just prior to dehydration in ethanol (after osmium tetroxide post-fixation, tissues should receive a buffer wash). Post-staining involves the use of the salts of the heavy metals, lead and uranium. Caution should be used in handling these heavy metal stains since they are extremely toxic, especially in powdered form. Uranyl salts are low-level radiation emitters. Disposal of heavy metals should be done in accordance with local regulations. The usual stains employed are uranyl acetate, followed by lead citrate. Uranyl acetate and lead citrate react will many cellular components including proteins and nucleic acids. If one does not stain with uranyl acetate, the nuclei are most affected with a noticeable lack of contrast due to weak or no staining of the nucleic acids contained within. Both stains are said to be positive stains since they increase the density of the structures under consideration as opposed to the background. In negative staining, the background density is increased relative to the structures (as in the negative staining of virus particles using 1-2% phosphotungstic acid - PTA). The preparation of stain solutions is described below. ✥ Uranyl acetate A 0.5% aqueous solution of uranyl acetate is prepared by dissolving 0.2g of uranyl acetate in 40ml of distilled water in an Erlenmeyer flask (50ml) and gently mixing over a 10 minute period. The solution will be clear and yellow with no notable sediment. The solution is photosensitive and should be stored in the dark at room temperature. The solution may be filtered or centrifuged just prior to use, however, this is usually not necessary so long as you take the solution from the center of the given volume. It should be noted that uranyl acetate solutions can also be prepared in methanol (saturated solution), ethanol or acetone and in volumes less than 40ml. Many investigators will prepare small volumes as needed to reduce the possibility of contamination.

UNIT 1 – PREPARATION OF BIOLOGICAL SAMPLES FOR TEM

49

✥ Lead Citrate There are a number of formulations for lead citrate with the most widely used developed by Reynolds (1963) and Venable and Coggeshall (1965). In the Reynolds formulation, 1.33g of lead nitrate and 1.76g of sodium citrate are added to 30ml of CO2-free, double distilled water in a 50ml Erlenmeyer flask and mixed regularly over a 30 minute period. At this point, 8ml of 1N (4%) NaOH is added with an obvious “clearing” of the solution upon mixing. Finally 12ml of the CO2-free, double distilled water are added with gentle mixing. The final working solution should be placed into sterile centrifuge tubes. The tubes should be filled to capacity to minimize CO2 bearing air from coming in contact with the solution. The tubes containing the solution should be stored in the cold at 4˚C and centrifuged for a least 5 minutes just before use. The final pH of the solution should be checked and ideally be alkaline at 12 or higher. The main problem associated with the use of lead citrate is contamination. Lead citrate readily reacts with atmospheric CO2 to form a crystalline precipitate known as lead carbonate. On your sections, lead carbonate deposits are extremely electron dense leading to large, unsightly dark staining artifacts which obscure detail. One should avoid breathing directly on the lead citrate solutions. As you stain, you should breathe out of the side of your mouth and also minimize the time of lead exposure to the air. The Venable-Coggeshall formulation allows you to prepare smaller quantities of the solution as it is needed. In this recipe, 0.01 to 0.04g of lead citrate are added to 10ml of CO2-free, double distilled water in a centrifuge tube along with 0.1ml of 10N (40%) NaOH and mixed. In order to prepare CO2-free distilled water, it can be autoclaved and sealed right after removal or it can be boiled for at least 10 minutes and then sealed while hot. Allow the water to come to room temperature before solution preparation. ✥ Post-staining Procedure Grids can be post-stained within 15 minutes following section collection, after the grids have air-dried. The following procedure should be used with care taken not to contaminate work areas with toxic heavy metal stains (fig. 5). • Using a squirt bottle, sprinkle water on the table surface. • Take a precut 4" by 4" square of Parafilm and pull along the water surface to produce a smooth, flat sheet without air bubbles. The sheet should be spread smooth with the protective cover (imprinted with the product name) in place as to prevent water from contacting the parafilm surface. Once flattened, the protective cover is removed and the film is covered with a Petri dish lid. • Place 10-15 pellets of NaOH (caution - caustic, causes severe burns) at the top of the parafilm sheet and recover with the petri dish lid. The NaOH should be wetted slightly and will serve to absorb atmospheric carbon dioxide, reducing the possibility of lead carbonate formation and contamination.

50 ELECTRON MICROSCOPY: A HANDBOOK OF TECHNIQUES FOR THE BIOLOGIST

• Using a Pasteur pipette, place as many drops of aqueous Uranyl acetate stain in a horizontal row to correspond with the number of grids to be stained, not to exceed five (5) drops per setup. If one holds the tip of the pipette near the surface of the parafilm and has good control over the pipette bulb, the diameter of the drops can be adjusted. The drops should be just slightly larger in diameter that the 3.0mm grids themselves. The UA drops should be placed in a row just below the NaOH pellets without coming in contact with the NaOH. • Using a different pipette, place an identical row of CO2-free distilled water drops below each drop of the UA without placing them too closely together. Then, skip a row (leaving room for a row of lead citrate) and form two (2) more identical rows of the CO2-free distilled water. • Without direct breathing on the solution or staining setup, lay out the row of lead citrate directly below the first distilled water row using a clean, new pipette. Recover with the petri dish lid. • STAINING: Place a grid section (dull) side down in a drop of UA using jeweler’s forceps and stain for 15 minutes with the petri dish cover in place. From 1 to 5 grids can be stained simultaneously. After 15 minutes the first grid in UA can be carefully picked up on edge and blotted on edge with a filter paper circle. The filter paper circle should be placed between the tines of the forceps which hold the grid. The circle can now be more accurately controlled in its movement to contact the edge of the grid. A small amount of stain will be absorbed onto the filter paper. The blotted grid can be placed into the water drop below the UA drop for rinsing (30-60 seconds). Repeat for each grid as necessary (up to a total of five). As blotting commences, the filter paper circle should be rotated to a clean area and eventually discarded. The tines of the forceps can be cleaned regularly throughout the process by blotting them on a clean filter paper region. After the first water rinse, blot each grid on edge as described and move them into the lead citrate stain for up to 5 minutes (DON’T BREATHE!!). After the lead citrate, the grids can be blotted and moved through two water rinses for approximately 1 minute each. After the final water rinse, the grid is blotted on edge and placed section (dull) side up on a grid gripper or stored on edge in a grid box to dry. After post-staining, grids can be observed using the TEM within 15 minutes, when they have dried. Be extremely careful not to damage the grids during staining. Since the grids are handled quite often during staining, it is common for beginners to mangle their grids. This is not desirable considering all of the work that has come before the staining process. If you encounter a large amount of staining artifact, you can try using a two setup method where one setup contains UA followed by three rows of water (without NaOH pellets) and the other contains NaOH pellets, a row of lead citrate and three rows of water. This setup minimizes lead citrate exposure to the air. Some investigators will use a rinse row of 0.01N NaOH below the lead citrate in this setup, however, some loss of lead staining may result.

UNIT 1 – PREPARATION OF BIOLOGICAL SAMPLES FOR TEM

Fig. 5

Parafilm (4"x4")

NaOH Pellets

Uranyl Acetate (15 min)

Distilled Water (30-60 sec)

Lead Citrate (2-5 min)

Distilled Water (30-60 sec)

Distilled Water (30-60 sec)

Petri Dish

51

52 ELECTRON MICROSCOPY: A HANDBOOK OF TECHNIQUES FOR THE BIOLOGIST

Chapter 4 - Grids & Grid Supports ✥ Grids Grids are typically 3.0mm in diameter and can be made of any non-magnetic metal such as gold, platinum and nickel. The most common metal used in the manufacture of grids is copper. They come in a variety of mesh sizes (referring to the number of bars per inch) and mesh patterns. Common grid mesh sizes are 200, 300, 400, up to 1000. It is recommended that large sections in ribbons be collected on 200 mesh or larger size grids. The ribboning helps support the sections and larger mesh sizes translate into a larger viewing area (less section regions covered by grid bars). Isolated sections should be picked up on 300-400 mesh grids in order that they be supported and not wrap around grid bars. Square mesh grids are routine, however, hex-mesh and other shapes are available. Finder-grids are also available with reference coordinates to help you relocate previously examined sections. Open hole and slot grids, which must be coated (see grid supports below), are useful for serial sectioning. Grids should be cleaned before use by sonicating them for 5 minutes in a small beaker of acetone. After sonication, the excess acetone is poured off and the beaker is placed upside down on a 9cm diameter filter paper circle in the large diameter lid of a petri dish. As the acetone evaporates, the clean grids will drop to the filter paper surface. Use the other half of the petri dish to cover the grids and label the their type and size with a marker. ✥ Grid Supports The use of modern epoxy resins in the embedding of biological samples for the TEM has reduced the need for grid support films. The three-dimensional polymerization of epoxy resins with the addition of the proper curing agents and heat (60˚C) or UV, results in an embedment which is resistant to damage from solvents and heat, including the electron beam. Ultrathin sections are therefore very stable under the electron beam. Embedding media which were used in the past for TEM, such as the methacrylates, were unstable under the primary beam with the resulting ultrathin sections requiring a support. Even with the use of epoxy resins, grid supports may be necessary. A good example involves the process of serial sectioning. Serial sections allow us to follow a particular structure through many ultrathin sections in order to develop an understanding of its threedimensional architecture. Since grid bars could block the ability to view the structure(s) of interest, typical square-mesh or hexagonal-mesh grids are not employed for the collection of serial sections. Instead, an open slot or circle grid is used (see below). The large open area cannot support sections, therefore, a grid support must be prepared.

UNIT 1 – PREPARATION OF BIOLOGICAL SAMPLES FOR TEM

Square Mesh Grid

1234567890 1234567890 1234567890 1234567890 1234567890 1234567890 1234567890 1234567890 1234567890 1234567890 1234567890

Hole/Circle Grid

53

Slot Grid

The most common type of support used today is either plastic - Formvar (polyvinyl formal) or carbon films or a combination of both. Formvar Films Working solutions of formvar in ethylene dichloride (0.25%) can be purchased directly from EM supply companies or you can prepare the solution yourself by placing 0.25g of formvar powder into 100ml of ethylene dichloride and gently mixing. The solution should be clear with no sediment and stored in a brown bottle out of direct sunlight and heat. CAUTION: Ethylene dichloride is flammable and is a potential carcinogen. When ready for use the solution should be transferred to a clean coplin jar. Supplies: 0.25% Formvar solution in Coplin jar clean 400ml plastic beaker (tri-pour) Glass Microscope Slides Ross Lens Tissue Single-edged Razor Blade Liquinox soap (non-filming) Distilled water Grids (to be coated) Jeweler’s Forceps The glass slides are cleaned using Liquinox soap, rinsed in distilled water and allowed to stand on end and air dry. The plastic beaker is then filled completely (about overflowing) with distilled water and returned to the work area. A clean glass slide is dipped slowly into the formvar solution and withdrawn. The end of the slide is placed on Ross lens tissue to remove the excess solution and allowed to air dry for a few minutes (some individuals gently wave the slide to hasten drying). The slide is now scored with a single-edged razor blade just inside the periphery of the glass slide and across the slide into three to four square regions (fig. 6). The slide is picked up (scored side up) and slowly introduced into the distilled water of the plastic beaker at a very shallow angle (20-30˚). Prior to the introduction of the formvar coated slide, the distilled water surface is cleaned by pulling a sheet of Ross lens tissue across the water surface, as was done to clean the water surface of the glass knife boat used in ultramicrotomy. Just before placing the formvar coated slide into the distilled water, one can breathe on the scored surface. “Frosting” the surface

54 ELECTRON MICROSCOPY: A HANDBOOK OF TECHNIQUES FOR THE BIOLOGIST

in this way is believed to facilitate stripping of the formvar off of the glass slide. Under fluorescent lighting, you should observe floating formvar “sections” in the beaker. These floating films should appear of a uniform silver color under fluorescent lighting. If the films are dark gold, purple, etc., the solution must be cut with the addition of ethylene dichloride. If the films are too thin, you can add more powdered formvar to the solution. Formvar films which display a variety of colors (rainbow) may be contaminated and a new solution should be freshly prepared. The greatest difficulty in working with these plastic films involves their stripping from the glass slide when placed into the distilled water. Exceptionally clean slides are not ideal for this purpose. You can also experiment with slides of different manufacturers as some prove better than others. To facilitate formvar stripping, a thin and even coating of oil from the skin (usually best when taken from alongside the nose) can be applied by index finger to one surface of the slide (make sure you know which surface is oil-coated since this is the side you will want to score). One can also use a thin coating of saliva on one side of the slide. Both techniques work well to strip the formvar from the slide and it should be noted that contamination of the formvar does not occur as a result. Once the formvar films are cast on the water surface, the slide can be dropped to the bottom of the beaker. Using the fine forceps, grids can be carefully placed, dull side down, on the floating formvar film. Concentrate the grids at one end of the film without overlapping them. The final technique involves collecting the coated grids, dull side up (with the formvar surface on top). Some individuals use a large container such as a fishtank to submerge the films from above with a glass slide. The slide is swept through the water in a large “U” fashion and is removed with the coated grids on top of the slide. Others use a rapid “slap and flip” technique with a glass slide to collect the coated grids. The simplest method involves placing a clean glass slide or wax coated cardboard strip exactly perpendicular (90˚) to the floating formvar film, in the region without any placed grids and gently submerging the slide using a straight downward force (fig. 7). This film with the coated grids just rolls up on the slide with the grids in the correct orientation (dull side up). The slide is removed from the water and allowed to air dry. Once dry, the grids are picked up from the slide surface and either used immediately or stored for later use. Fig. 6 - Formvar Coating on Glass Slide

Glass Slide

Formvar Coating

Razor Blade Score Lines

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Fig. 7 - Collection of Formvar Coated Grids

Razor Blade Score Lines

Glass Slide

Clean DH2O Surface

Slot Grids (Dull Side Down)

56 ELECTRON MICROSCOPY: A HANDBOOK OF TECHNIQUES FOR THE BIOLOGIST

Carbon Coating Carbon films are strong and extremely stable under the electron beam. Additionally, they can be made very thin and are essentially electron transparent. Carbon coating can be carried out alone or on top of plastic films in order to provide increased stability. Some investigators also “sandwich” various support films around the ultrathin sections (plasticsections-plastic, plastic-sections-carbon). Carbon coating is usually conducted in a high vacuum evaporator. The high vacuum created in the bell jar is required so that air molecules will not interfere with the evaporated carbon and lead to an uneven coating. Two pure carbon rods are prepared so that one is sharpened to a point and the other is flat on end. The rods are locked into the two suitable electrodes in the evaporator with the pointed end of one rod in direct contact with the flat end of the other. Beneath the carbon rods are the grids you wish to coat on a glass slide and a white porcelain plate with a drop of low vacuum oil or immersion oil at its center. The bell jar is evacuated to 10-5 Torr and voltage is applied to the electrodes. At the pointed carbon rod tip, it becomes white hot and carbon is evaporated. The carbon falls to evenly coat the grids. Thickness can be determined by the darkness of the porcelain plate in comparison to the central oil spot. Due to the presence of the oil, the area beneath it will not darken but rather, remain a bright white. The desired color of the indicator plate is described as a light tan/brown which results in a 400-700Å thick coat (fig. 8). Since carbon does not adhere well to naked copper grids, an initial coating of plastic such as formvar is recommended. If desired, the plastic coat can later be dissolved away in chloroform and ethylene dichloride. Carbon coating attachments are also available for low vacuum sputter coaters (see discussion of sputter coating in Unit 2 on SEM Sample Preparation). Carbon coating of SEM samples over metallic coating (gold, etc.) is required when elemental analysis is being conducted.

UNIT 1 – PREPARATION OF BIOLOGICAL SAMPLES FOR TEM

Fig. 8 - Carbon Evaporation Setup

Bell Jar

carbon rods

electrode oil grids

57

58 ELECTRON MICROSCOPY: A HANDBOOK OF TECHNIQUES FOR THE BIOLOGIST

Unit 2 - Preparation of Biological Samples for SEM Since the same high vacuum environment (10-5 Torr) is required in order to generate an electron beam in the conventional SEM as is required for the TEM, the observation of living samples is not usually possible (environmental ESEM’s and differentially pumped, low vacuum SEM’s may permit the observation of living samples). However, because the SEM is utilized primarily in the observation of surface features, the tedium of ultra-thin sectioning, necessary to TEM specimen preparation, is avoided. Depending on the nature of the surface of interest, a sample might be prepared simply by adhering it to an aluminum support or stub and examining it in the SEM. Of course, such a sample would have to be hard and conductive, characteristics rarely found together in biological specimens. Common examples of hard surfaces include chitinous insect and crustacean exoskeletons, lignified cellulose cell walls of plant material/wood, calcium carbonate shells of molluscs/bivalves such as oysters and clams, silicon frustules (shells) of diatoms, and complex calcium and phosphorus hydroxyapatites which comprise part of the non-living matrix of bone. A problem exists in that these and other biological samples are usually insulators which are incapable of emitting an adequate signal when contacted with the SEM primary electron beam. Lack of adequate signal leads to an inferior quality image (low SNR). In addition, there are many occasions when one wishes to examine a soft tissue sample which would readily degrade under the high vacuum operating environment of the SEM, not to mention, be subject to the forces of autolysis and decomposition. Another factor to consider is whether the visualization internal structure using a SEM is desirable or even possible. Since ultra-thin sectioning is not required for SEM samples, they can typically be larger (recall that TEM tissue blocks had to be minced to a thickness no greater than 0.5mm in one dimension to ensure complete infiltration of fixatives and embedding media). When dealing with soft tissues, the tissue pieces should remain somewhat small (~2-3mm3) to allow for complete penetration of the primary fixative and prevention of internal collapse and resultant artifacts in surface morphology (shape). Also of importance is the handling of small samples and microbes. The following chapters shall be divided into the preparation of a variety of samples including hard tissues/structures, soft tissues, internal structures, microbial samples and SEM samples prepared for the TEM for SEM/TEM correlation.

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Chapter 5 - Hard Tissue Preparation As stated earlier, hard tissue samples can include chitinous insect and crustacean exoskeletons, lignified cellulose cell walls of plant material/wood, calcium carbonate shells of molluscs/bivalves such as oysters and clams, silicon frustules (shells) of diatoms, the non-living matrix of bone and teeth. Such samples may have adherent soft tissue and/or surface debris associated with them. This material should be removed using as delicate a technique as possible. Surface lint and dust may be easily removed with compressed air (care should be taken since some compressed air sources contain particulates which could induce surface damage). In addition, surface debris can be rinsed away using distilled water or an isotonic physiologic buffer solution (the need for an isotonic solution is questionable since hard tissues are unlikely to shrink or swell osmotically). In the case of soft tissue adhering to samples such as bone, the soft tissue can be dissolved away by boiling in 28% ammonium hydroxide followed by subsequent rinses in distilled water. CAUTION: Ammonium hydroxide must be opened and used in a fume hood. Due to its extremely high vapor pressure, a bottle of 28% ammonium hydroxide opened in even a well ventilated room will quickly saturate the atmosphere with toxic ammonia. Inhalation could prove to be fatal. Following cleaning, which may not be necessary, hard tissue samples are mounted to a standard 15mm diameter, aluminum specimen mount or stub, using an appropriate adhesive (types of suitable adhesives will be covered later in this unit). Depending on specimen conductivity, or the usual lack of it when dealing with biological samples, and the desired SEM imaging voltage (low ≤ 5kv or high > 5kv to 25kv), the mounted samples will then be given an ultra-thin (100 - 200Å) conductive coating of a metal such a gold, platinum, palladium, or gold-palladium. Since most samples are generally insulators and high voltage is usually selected for maximum signal generation and the resultant high quality images, the norm involves the application of this conductive coating. The techniques and procedures of sputter coating and vacuum evaporation will be covered in detail later in this unit, in conjunction with soft tissue preparation. It should be noted that even though hard samples are inherently more durable than their soft tissue counterparts, care should be exercised in handling these samples so as not to induce artifacts. This is especially true of samples such as small insects which should be gently attached to the specimen stubs using fine forceps. The use of liquid adhesives for small samples should be avoided since the liquid has a tendency to creep up the sides of the specimen and envelope it. This is not a problem if one wishes to study glue!

60 ELECTRON MICROSCOPY: A HANDBOOK OF TECHNIQUES FOR THE BIOLOGIST

Chapter 6 - Soft Tissue Preparation In order to examine soft tissue samples in “as near to life-like condition as possible”, chemical fixation is required. Without chemical fixation, post-mortem autolysis and decomposition would distort even the surface of the specimen of interest. Even with the surface “stabilized” through conductive coating, internal degradation would lead to glaring artifacts of surface morphology. In addition, chemical fixation will permit observation of internal structures using cryo (cold) techniques and allow for the correlative preparation of SEM samples for the TEM via epoxy resin embedment and ultra-thin sectioning. The preparation of soft tissue samples for examination in the SEM begins in an identical manner to the preparation of samples for the TEM (see Unit 1 - Preparation of Biological Samples for TEM). A generalized protocol follows: •Primary Aldehyde Fixation •Buffer Wash •Secondary Osmium Tetroxide Postfixation •Optional Buffer/DH2O Wash •Dehydration Series (Ethanol/Acetone) •Intermediate Fluid Series (Freon TF/113) •Critical Point Drying (in Transitional Fluid - liquid CO2 or Freon 13) •Mounting (adhesive on 15mm dia. aluminum stub) •Conductive Coating (Sputter Coater or Vacuum Evaporator) Firstly, the tissues of interest must be excised and placed into the primary fixative. As for TEM preparation, we typically use a combination method of in situ and immersion fixation. The tissue blocks for SEM preparation can be somewhat larger since the aldehydes (primary fixative) can penetrate through at least 3mm of sample. Tissue blocks are therefore best minced to the dimensions of 2-3mm3. If strips of tissue are cut, they should be no thicker than approximately 2-3mm on one of the sides. Minced tissue blocks can be transferred to vials containing the primary aldehyde fixative and remain for one hour. Once again the best single aldehyde choice is the doubly reactive, cross-linking, glutaraldehyde. As for TEM sample primary fixation, we use a 3% concentration of glutaraldehyde carried in the proper isotonic buffer vehicle (for most soft mammalian tissues, 0.2M phosphate buffer, pH 7.2-7.4 is ideal). The chemical action of aldehydes (stabilization of the cellular protein matrix) and other fixatives along with the purpose of buffer solutions and their preparation are covered in the chapters on TEM specimen preparation. Once the primary fixation is complete, three buffer washes of ten minutes duration each are conducted to remove unbound aldehyde and prevent the undesirable precipitate reaction with the secondary fixative, namely, osmium tetroxide.

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Following the buffer washes, the samples are placed into the secondary fixative, osmium tetroxide. The OsO4 will react mainly with unsaturated lipids and impart an increase in conductivity to the biological sample. The OsO4 is also carried in the same buffer vehicle as for the glutaraldehyde. You will recall that the benefit to osmium tetroxide fixation for TEM was the introduction of an electron dense stain (for phospholipid membranes and other osmophilic structures) for added contrast. In SEM, the generation of a signal from the surface is of paramount concern. Incorporation of the heavy metal osmium to the sample allows for increased surface signal emission when contacted by the primary electron beam of the SEM. Later in this unit, I will discuss how osmium incorporation can be enhanced by the addition of agents such as tannic acid and thiocarbohydrazide (TCH), so much so, that conductive surface coating may be avoided. An optional buffer or distilled water wash of two changes for ten minutes duration each may be performed, although, I have not observed any artifacts arise due to the omission of this step. Next, the tissue must undergo complete dehydration, but not for the same reason as TEM sample dehydration. Residual water in the tissue would cause surface tension collapse as it dried. For this reason, the common process of critical point drying - CPD (or some alternative) is employed in the preparation of samples for the SEM. The CPD process, which is carried out under liquid carbon dioxide, would be ineffective if water remained in the tissue block. A complete description of CPD theory and procedure is found later in this unit. By contrast, you will recall that samples are dehydrated for TEM in order that the epoxy resins infiltrate the tissue blocks (epoxies are not miscible with water). Both acetone and ethanol (EtOH) are common dehydrating agents used. Since shrinkage of tissue blocks and the resultant surface distortion is undesirable for SEM specimens, the dehydration schedule is more gradual for SEM, starting at 30% EtOH. The usual ascending series is 30%, 50%, 70%, 95% EtOH for ten minutes each, followed by 100% EtOH for two changes of ten minutes each with the vials being filled to capacity as usual. The preparation of such a dehydration series using 95% (not 100%) ethanol has been described in the unit on TEM specimen preparation. The final steps in the soft tissue protocol will involve the preparation of tissue blocks for critical point drying which has traditionally been performed in either liquid carbon dioxide or liquid freon (Freon 13). Usually, before the critical point drying process, tissue blocks are passed through an ascending intermediate fluid series of Freon TF/113 since it is miscible with both ethanol and the transitional fluid used in critical point drying. The freon is diluted with 100% ethanol, not water, in order to produce the concentration series. The usual ascending series is 30%, 50%, 70%, 95% Freon TF/113 for ten minutes each, followed by 100% Freon TF/113 for two changes of ten minutes each. As the scientific community has become more aware of the environmental impact of freons on the degradation of the protective ozone layer (which partially shields the earth from harmful UV radiation), alternatives to its use have been discovered and will be considered following the discussion of critical point drying. A common past alternative to Freon TF

62 ELECTRON MICROSCOPY: A HANDBOOK OF TECHNIQUES FOR THE BIOLOGIST

was amyl acetate with its characteristic strong, banana-like odor. It was typically used as an intermediate fluid between ethanol and liquid CO2. Due to its toxicity, it should always be purged out of a critical point dryer into a fume hood. A benefit to its use was that one always knew it had been completely purged from the critical point dryer be the marked absence of its trademark odor. The process of chemical fixation of biological soft tissues have been carried out in a liquid environment. If in the final step, these tissues are simply allowed to air dry, tremendous surface tension distortion will occur leading to obvious surface artifacts. It should be noted that water can exert a surface tension force of over 2,000 psi. In order to prevent this surface tension damage, the technique of critical point drying (CPD) has been utilized since 1968. The principle of CPD involves an understanding of phase (solid-liquid-gas) boundaries, especially, for our purposes, the boundary between liquid and vapor. Every fluid possesses what is known as a critical density (Dc) at which the boundary or interface (the actual liquid surface) between the liquid phase and the vapor phase becomes indistinguishable. At first, when a fluid is introduced to a sealed container, an equilibrium exists between the liquid and vapor phases. In order to attain critical density, this equilibrium must be shifted to the vapor phase through heating and the related increase in pressure. Critical density (Dc) is attained at the critical temperature (Tc) and critical pressure (Pc) of the given fluid. Therefore, to eliminate the interfacial boundary between liquid and vapor and avoid the associated surface tension force, the fluid must be elevated to its critical temperature and pressure. Since the critical temperature and pressure of water is excessive, at 374˚C and 3,184 psi respectively, and would damage delicate biological soft tissues, other liquids typically serve as transitional fluids (so named because they make the transition between liquid and vapor phases). The most common of these is liquid carbon dioxide (LCO2) and liquid freon 13. Due to its higher price and negative impact on the environment, liquid freon has become the less popular of the two transitional fluids. Liquid CO2 has a critical temperature and pressure of 31˚C and 1,073 psi, respectively, compared with liquid freon 13 at 28.9˚C and 561 psi. The liquid CO2 is introduced to the critical point dryer at the high pressure (600 -800 psi) of its storage tank. The storage tank is specially ordered with a siphon tube which takes the liquid CO2 from the bottom of the tank since as the tank is emptied, CO2 gas rises and collects at the top of the tank. CAUTION: Liquid carbon dioxide is under high pressure and is very cold. Use care in handling the tanks and opening the tank valve. The use of a regulator is not required, however, the tank pressure is between 600-800 psi. Make sure the hose between the CPD and the tank is threaded and tightened properly, and is a hose rated for high pressure applications (a minimum burst pressure rating should be printed on the hose - the hose used in this lab is rated at 17,000 psi). In addition, the storage tank

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should be chained or strapped to a wall or a tabletop to prevent it from accidentally falling and rupturing. Under its high pressure, the tank could act as a missile causing serious injury. Finally, CO2 is a colorless, odorless gas and should be vented from the CPD into a fume hood to prevent asphyxiation as it displaces the normal atmosphere. ✥ Critical Point Dryer Operation The CPD is a thick walled metal chamber which can withstand pressure in excess of 2,000 psi. The chamber is constructed of a metal of excellent heat conductive properties such as brass, copper or bronze. The chamber is usually machined to accommodate some type(s) of specimen holder(s) which will retain the tissue blocks throughout the procedure. The chamber also includes three high pressure needle valves, the inlet valve (for the introduction of the LCO2), the drain valve (to drain off the intermediate fluid - freon TF/ 113) and the vent valve (to vent the CO2 gas). Additionally, the CPD chamber must have some provision for heating the transitional fluid. In the more expensive models, there is an electrical heater. In the least expensive models, the entire CPD chamber is simply lowered into a container or bucket of hot water. The main drawback to this method is that residual external water on the unit might contact the tissues as they are removed, rehydrating them. Our Pelco Jumbo Dryer (same as the Polaron Jumbo) is designed with a water jacket just external to the drying chamber. Hot and/or cold water can be routed through the jacket by means of plastic tubing attached to a faucet. A temperature gauge is added in order to monitor the water temperature, although, the only required gauge is a chamber pressure gauge. The Pelco CPD is equipped with a safety valve which is designed to rupture at 2,000 psi. CAUTION: The CPD is a high pressure device which if used improperly can lead to serious injury or death. Follow all manufacturer directions carefully. If equipped with a thick quartz view window (as in the case of the Pelco Jumbo Model), it MUST BE EXAMINED FOR CRACKS BEFORE EACH USE!!! NEVER REMOVE the protective Lexan shield, if so equipped, just outside the view glass - it is there for your safety!! NEVER LOOK DIRECTLY INTO THE VIEW GLASS (use a mirror if you are curious) - if it were to break, the high pressure would produce many high velocity, very sharp projectiles! Ideally, the CPD should be located in a fume hood. For obvious reasons stated above, the CPD is alternatively known as “the BOMB”. In the routine operation of the Pelco CPD, the unit is placed into the fume hood and the high pressure hose is connected tightly to the liquid CO2 tank. The inlet water hose is attached to the faucet and the outlet water hose is run into the hood sink. The drain hose (optional) is also run into the hood sink. The heavy rear door is removed using the steel rod tool. If this is the first CPD run of the day and the bomb is at room temperature, you

64 ELECTRON MICROSCOPY: A HANDBOOK OF TECHNIQUES FOR THE BIOLOGIST

will not have to cool the unit by running cold water through the water jacket. If the bomb has been heated, it will have to be cooled to approximately 20˚C before use. It is often a good idea to perform a “dry run” of the unit to check it for any leaks (valves, window and rear door dowdy seals may require tightening) - it is better to test the unit than to sacrifice good tissue blocks which will ultimately dry down with surface tension collapse in a bomb which cannot maintain pressure! All three needle valves should be closed clockwise. The valve on the liquid CO2 tank can now be opened. Tissue blocks in their second change of 100% freon TF/113 can now be loaded into the CPD boat. Most CPD units come with a device to contain any number of specimen holders. The Pelco unit comes with a three reservoir/channel boat into which some nine specimen baskets, which resemble sewing thimbles, can be loaded. The boat has a large rear stainless steel pin which interfaces with a hole in the rear door of the CPD. At the front of the boat is a spring loaded valve which when inserted into the CPD, opens to allow the intermediate fluid to drain out. Firstly, the CPD boat is filled with freon TF/113. At this point, the tissues are transferred from their vials to the tissue baskets. Sine most tissues will float in 100% freon TF/113, they are easily transferred by pouring the vials contents into the baskets over a sink. CAUTION & REMINDER: All potentially toxic chemicals used in the fixation process should be handled using disposable gloves and wearing goggles, ideally within a fume hood! Once transferred to the baskets, the baskets are quickly placed into one of the three CPD boat channels which are full of the intermediate fluid, freon TF/113, to prevent premature drying down through a liquid interface. Sheets are available (fig. 9) to identify the specific tissue which occupies a specific basket in the boat. Once a channel contains three baskets, a wire mesh cover is placed on top of the baskets to prevent tissue loss in the bomb. The CPD boat can hold nine baskets at a maximum. Once the boat is loaded with tissue baskets, the boat is carefully carried over to the bomb and gently inserted. A slot on the rear underside of the boat must align with a corresponding slug in the rear floor of the CPD chamber. The central hole of the rear door is lined up with the stainless steel pin of the boat and the rear door is threaded clockwise until tight. Use the steel rod tool to tighten the rear door and prevent leaks. Your tissues are now ready to undergo critical point drying. With the tissues properly loaded into the CPD and with all valves closed, the three valves will be used (or as I like to call it, juggled) in order to introduce the liquid CO2 and flush out the freon TF/113 (fig. 10). Initially, the upper inlet and vent valves are opened. Do not be alarmed at the amount of noise coming from these high pressure needle valves. The vent valve must be opened to allow for the escape of gas and permit the required volume of transitional fluid to enter the bomb. You should be able to observe the liquid CO2 enter the bomb and rise to a desired level above the boat. As the liquid CO2 is entering the bomb, the lower drain valve must be opened to flush out the freon TF/113

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under pressure. Fig. 9

At first, a clear fluid will be observed flowing down and out of the drain line, followed by a solid CO2 “snow”. Allow the solid CO2 to exit the drain line for a few seconds, ensuring complete flushing out of the freon TF/113, and then close the drain valve. Using the inlet and vent valves, allow the liquid CO2 to reach a volume just above the top of the boat, then close both valves. At this point, you should not hear any “hissing” which would be typical of a seal or valve leak. The pressure gauge should be holding steady at 600-800 psi (The actual liquid CO2 tank outlet pressure).

66 ELECTRON MICROSCOPY: A HANDBOOK OF TECHNIQUES FOR THE BIOLOGIST

Fig. 10 INLET VALVE

VENT VALVE

WATER INLET

REAR DOOR

VIEW GLASS

SAFETY VALVE

DRAIN VALVE

WATER OUTLET SUPPORT COLUMN

PEDESTAL

Now, hot water can be flowed into the water jacket. The flow rate should be moderate (as viewed directly at the outlet hose) and the water temperature should be approximately 4045˚C. As time passes and the heat is conducted from the water jacket to the CPD chamber, the temperature gauge and pressure gauge will rise accordingly. At approximately, 29˚C and 1,000psi, you will observe a turbulence at the liquid CO2 interface, followed by the

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instantaneous disappearance of the liquid interface at critical temperature and pressure (once again, the use of a mirror is recommended for viewing this process vs. looking directly into the CPD view glass). When this phenomenon occurs, at critical temperature and critical pressure for the given fluid, the density of the liquid phase equals the density of the vapor phase (known as the critical density) and the boundary between the two phases can no longer be distinguished. The tissues within the bomb have been critical point dried without surface tension distortion. Even though the critical temperature and critical pressure for liquid CO2 is 31˚C and 1,073 psi, respectively, the bomb chamber is heated to a somewhat higher level (approximately 40˚C and 1,500 psi) to prevent recondensation of the CO2 vapor. The bomb is held at this level for a few minutes then slowly (approximately 100 psi/min.) vented to the atmosphere using the vent valve. Rapid venting would result in recondensation of the CO2 vapor and wetting of the dried tissue blocks. At atmospheric pressure, the CPD boat is carefully removed to a dry location by slowly unscrewing the rear door of the CPD. The tissue baskets are uncovered and removed to a clean, dry work area. Small (7ml) specimen vials are ideal for supporting the tissue baskets at the work area as long as they are dry internally. The tissue blocks, which are obviously dry and somewhat brittle in appearance, are now ready for adhesive mounting to specimen stubs. ✥ Alternatives to Critical Point Drying Due to current concerns with the effects of chlorofluorocarbons (CFC’s) such as freon on the environment (protective atmospheric ozone), the tendency has been away from the use of intermediate fluids and transitional fluids and critical point drying. Fluorocarbon Drying In an alternative to critical point drying, a solid fluorocarbon such as Peldri II, has yielded good results. In the use of Peldri, this solid fluorocarbon is heated to its melting point and held until use in its liquid state. The tissue blocks are brought through the dehydration series and placed into the liquid Peldri. The mixture is taken off the heat source and allowed to solidify. Once solid, the samples are placed into a low vacuum environment and the solid Peldri sublimates to the vapor state without passage through a liquid interface. Dry tissue blocks are mounted and coated as usual. Organo-Silicon Compounds A more recent alternative to the use of freons as intermediate fluids and transitional fluids for CPD is the use of the organo-silicon compounds, Tetramethyl silane (TMS) and Hexamethyldisilazane (HMDS). Samples are simply placed into these compounds after routine dehydration. Although liquids at room temperature, these agents evaporate from the sample surface without the usual surface tension forces exerted by other liquids. Air drying is therefore performed using these compounds without the surface artifacts produced when using other liquids, such as water. Air drying using these compounds has

68 ELECTRON MICROSCOPY: A HANDBOOK OF TECHNIQUES FOR THE BIOLOGIST

been reported at both room temperature and 60˚C (in an incubator). Adhesive mounting of dry tissue blocks to the typical, 15mm diameter, aluminum specimen stub is a rather simple matter. Firstly, the specimen stubs should be previously cleaned by sonicating them in a beaker of acetone for 5-10 minutes. The stubs are removed and allowed to air dry on a sheet of lint-free cloth. A variety of adhesives have been used for mounting SEM specimens to the stubs. They include liquids such as cyanoacrylates (‘crazy glue’), rubber cement, low resistance contact cement, silver paint (for a conductive contact to the stub), and Pliobond (manufactured by Goodyear - it is easily solubilized in acetone and ideal for removal when taking SEM tissue blocks to resin embedment and sectioning for TEM). The main drawback to liquid adhesives is their tendency to creep up the sides of a small sample and quite possibly cover the surface in the glue. To prevent this, a long time favorite for the attachment of small samples has been double sided scotch tape. Even better, we have found that adhesive transfer tabs, manufactured by the Avery label company, to be ideal for the adhesive mounting of most every SEM sample. These glue tabs are simply lifted off their backing sheet and placed in the center of the stub. Downward pressure is applied as indicated on the surface of the tab, and the tab is removed, leaving a thin layer of adhesive behind. Tissue blocks, or any other sample, are carefully handled with jeweler’s forceps and placed into the adhesive of choice. A stereo-microscope can be used if specific tissue orientation is important to your study. By example, if you wish to examine small intestinal villi, you should image the sample under the stereoscope to ensure that the villi are facing upward versus placing them into the glue. The final step in the preparation of soft biological samples for SEM, not to mention hard samples, is the application of a thin (approximately 100-200Å thick) conductive coat to the surface of interest. This is done since most biological samples are insulators and would yield a poor surface signal (secondary electron emission) when contacted with the primary electron beam. In addition, these samples would readily degrade under the high voltage (>5kv) beam. Aside from high voltage beam damage, conductive coating is required to prevent what is known as charging artifact; a buildup of negative surface charge which serves to repel the primary electron beam and also prevents the loss of secondaries from the surface. Uncoated samples could be imaged under low voltage (≤5.0kv) conditions, however, with a resultant loss of image quality (low SNR). ✥ Conductive Coating Currently, there are two techniques utilized in the modern EM lab for putting a thin conductive coat on a sample. They are Vacuum Evaporation and Sputter Coating. Each method will be described below.

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Vacuum Evaporation In this technique, a high vacuum evaporator is required. This device (see fig. 8) was originally described in the chapter on TEM Grids & Grid Supports (Chapter 4) under the heading of carbon coating. The main difference here is that in place of mounting the flat and pointed carbon rods into the evaporator electrodes, a pointed (bent into a V-shape) piece of tungsten wire is mounted. At the bend/point of the tungsten filament, a short, thin piece of gold, platinum or gold/palladium is wrapped. Directly under the tungsten filament is the rotating mechanical specimen stage with the provision for insertion of specimen stubs. Once the filament and stubs are set up, the bell jar is put in place and the volume is evacuated to high vacuum (10-5 Torr). At this point, the stage motor is energized and the rheostat is advanced until the tungsten burns white hot and the gold, gold/palladium, etc. melts (a noticeable bead of molten metal forms) and then evaporates to thinly and evenly coat the sample under the rotating stage and high vacuum conditions. The bell jar is brought back to atmospheric pressure and the samples removed from the stage. Care should be exercised when removing the samples. Be sure not to touch the embrittled tungsten wire which might break and cause large particulate contaminants to rain down upon your unremoved samples. After use, the evaporator bell jar environment must be restored to high vacuum – all high vacuum systems should always be maintained at high vacuum. CAUTION: The process of evaporating these metals should be observed through a piece of thick, dark glass. The intense brightness could permanently damage the retinas of your eyes.

Sputter Coater The sputter coater operates at low vacuum (10-2 Torr) using a small rotary pump to evacuate a Pyrex cylinder. The system is typically maintained at atmospheric pressure. Operation of a modern sputter coater, such as the Denton Desk II in this lab, is quite simple and somewhat automated. The one requirement for the sputter coater is a tank of ultrapure (at least 99.9% pure) argon gas. Such tanks of inert gas are under high pressure (approximately 2,500 psi when full) and must be regulated down (this tank must also be chained to a table or wall near the coater unit). A single or double stage regulator is required to bring the outlet pressure to 5 psi. A plastic or tygon tube connects the tank/ regulator to the sputter coater. Before use of the coater, the argon tank valve must be opened and the regulator pressure knob advanced to 5 psi. If there is a valve after the regulator as is our case, it must be fully opened (all valves typically open clockwise unless otherwise noted). You should understand the argon is not yet flowing into the sputter coater chamber. It is flowing down the plastic tubing and is stopped at an automatic solenoid valve. The sputter coater has a hinged head (fig. 11) into which runs a high voltage (1-3kv) cable. This cable interfaces with a thin gold (or platinum, palladium, etc.) foil target which serves as the source of the conductive heavy metal. A replacement gold target currently costs approximately $400.00.

70 ELECTRON MICROSCOPY: A HANDBOOK OF TECHNIQUES FOR THE BIOLOGIST

Once the stubs with mounted samples are placed on the platform/stage under the sputterhead, the cylinder is evacuated to low vacuum conditions (50 millitorr) while the ultrapure argon gas is introduced. A negatively charged high voltage field (1-3kv) is applied to the gold source cathode/target. This negative field ionizes the argon atoms to positive ions as per the equation below: Ar˚ ➞ Ar+ + eDue to electrostatic attraction, the positive argon ions are accelerated to the negative gold source cathode. The argon ions randomly strike the gold target forcibly causing the ejection of gold atoms from the target at various angles. These gold atoms build up an even conductive coat on the surface of the samples. A permanent magnet attracts stray electrons which result from the ionization of the argon atoms, and sends them to ground. These energetic electrons could interact destructively with the surface of the sample if not removed. It should be noted that sputter coating can be conducted under atmospheric conditions, however, ionization of many normal atmospheric components, even water, results in the formation of a number of highly reactive ions/radicals which would readily damage the surface of our delicate biological samples. Argon is chosen since it is an inert, stable or noble gas. Once samples have been coated, the chamber is returned to atmospheric pressure and the stubs are carefully removed. NOTE: After samples have been conductive coated, they should be stored in a desiccator toprevent absorption of atmospheric water vapor. If desiccator chambers are not available, small desiccators can be fashioned out of Petri dishes. The lower rim should receive a thin layer of vacuum grease or petroleum jelly. Small holes can be punched or burned into large BEEM capsules which are then filled with silica gel and capped. The capsules (1-2 per dish) are placed into the Petri dish. They need to be checked at least weekly as the silica gel hydrates (you will observe a color change from deep blue to pink to clear as the silica gel hydrates). Hydrated silica gel can be collected and baked in an oven at 350˚F to drive out the water for reuse.

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Fig. 11

HIGH VOLTAGE CABLE

SPUTTERHEAD

GOLD TARGET CATHODE

PERMANENT MAGNET

SPECIMEN STAGE

Argon Inlet

To Rotary Pump

Denton Desk II Operation This modern sputter coater can coat samples reliably in under five minutes (compared with at least 30 minutes for the vacuum evaporator). It has LED readouts for pressure (in millitorr), current (in milliamps) and time (countup or countdown). It can be run in manual or auto timed modes as desired. Generally, a test run is desirable to establish normal operating parameters such as pressure and current before introducing your valuable sample stubs. The routine operation follows: • Prepare argon tank and regulator (5 psi outlet pressure) as described previously. • Switch on main circuit breaker on rear of machine. • Open sputterhead and place stubs on top of stage. • Close sputterhead making sure it seats properly on the seal - vacuum grease is usually not necessary.

72 ELECTRON MICROSCOPY: A HANDBOOK OF TECHNIQUES FOR THE BIOLOGIST

• Switch on MAINS (red switch) on the right side of the machine - the rotary pump will come on. • Observes pressure gauge and allow to drop below 100 millitorr (when mains are first energized, the OFF button flashes, then glows steady - it MUST glow steady to proceed to the next step). You may need to adjust the pressure using the pressure control knob near the pressure LED. • Depress the SPUTTER button - you will hear a “pop” which is the opening of the solenoid valve to admit the argon gas. At this point, the SPUTTER knob light should be glowing steady. This step may fail at first attempt. In that case, allow the pressure to once again drop below 100 millitorr and try again - you may need to adjust the pressure knob. • Adjust the pressure using the pressure control knob to 50 millitorr. • For manual timing, depress the manual start button & for automatic timing depress the timed start button - in either case high voltage is applied to the gold target and a plasma field is observable as a bluish ring just below the target. The manual timer counts up and the auto timer counts down from whatever you preset the time for (up to a maximum of 999 sec). • Using the current control knob, quickly regulate the current to 45 milliamps - if the plasma field weakens and begins arcing/flashing, the pressure needs to be increased using the pressure control knob. • Once the desired time has elapsed, press stop for manual timing (auto timing will auto shut off the high voltage after the preset time has elapsed). • Turn off the MAINS switch - the solenoid valve will shut and the vent valve will automatically open to restore atmospheric pressure. • Open sputterhead and remove stubs to a desiccator for storage. • Repeat as needed for coating of each stub (the stage accommodates about six 15 mm diameter stubs). As noted earlier, a manual test run with no stubs present is desirable to preset the vacuum and current at optimum levels. Maintenance of the sputter coater is rather simple as well. Each year, the rotary pump oil and mist filter should be changed. When the gold foil target breaks through continued usage, it is easily replaced by removing the four thumbscrews which hold the meshwork cage in place. Four screws are removed from the retaining ring which holds the circular foil in place. The old foil is removed and the new foil put in place using the retaining ring and meshwork cage. On occasion, the pyrex cylinder should be cleaned of sputtered gold using 95% ethanol. If heavily coated, a good metal polish such as Wenol or Pol, followed by 95% EtOH works well.

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✥ Comparison of TEM and SEM Soft Tissue Protocols When one compares the preparation of soft tissue for the SEM and the TEM, a number of obvious differences becomes apparent. The following table summarizes those similarities and differences which were detailed in the chapters on SEM and TEM specimen preparation: TEM Protocol

SEM Protocol

3% Glutaraldehyde (Phosphate buffer) – 1hr.

same

Buffer Wash – 3 X 10 min.

same

1% OsO4 (Phosphate buffer) – 1hr.

same

Optional Buffer/DH2O Wash – 2 X 10 min.

same

70%, 95%, 100% EtOH – 2 X 10 min. each

30%, 50%, 70%, 95% EtOH – 1 X 10 min. each 100% EtOH – 2 X 10 min. 30%, 50%, 70%, 95% Freon TF – 1 X 10 min. ea 100% Freon TF/113 – 2 X 10 min. Critical Point Dry in Liquid CO2 (31˚C, 1,073 psi)

Propylene Oxide (P.O.) – 3 X10 min. each 1 Part P.O. : 1 Part Epoxy Resin – 1hr. Pure Resin – Vacuum Infiltrate – 1hr. Embed in BEEM Capsules Cure @ 60˚C – 48hrs. Ultra-thin Section

Sectioning Not Required

Mount Sections on Grids

Mount Tissue Blocks on Stubs

Heavy Metal Staining

Conductive Coating

74 ELECTRON MICROSCOPY: A HANDBOOK OF TECHNIQUES FOR THE BIOLOGIST

Fixation Schedule SEM Mammalian Soft Tissue Protocol - In-situ and Immersion fixation - 2-3mm3 tissue blocks Initial Fixation at 4˚C - Ascending to Room Temperature

Schedule

Duration

3% Glutaraldehyde (0.2 M phosphate buffer, pH 7.4)

1 hour

Buffer Wash

3 x 10 minutes each

1% OsO4 (buffered as above)

1 hour

Buffer or DH2O Wash (optional)

2 x 10 minutes each

30% Ethanol 50% Ethanol 70% Ethanol 95% Ethanol 100% Ethanol (fill vials completely)

1 x 10 minutes each 1 x 10 minutes each 1 x 10 minutes each 1 x 10 minutes each 2 x 10 minutes each

Optional LN2 Cryofracture - between 1st & 2nd 100% EtOH - see description in text! 30% Freon TF/113 50% Freon TF/113 70% Freon TF/113 95% Freon TF/113 100% Freon TF/113

1 x 10 minutes each 1 x 10 minutes each 1 x 10 minutes each 1 x 10 minutes each 2 x 10 minutes each

Critical Point Dry - in LCO2 Adhesive Mount on Aluminum Stubs Sputter Coat (Gold) - 50mT, 45mA

30-45 seconds

UNIT 2 – PREPARATION OF BIOLOGICAL SAMPLES FOR SEM

Fixation Schedule Worksheet SEM Mammalian Soft Tissue Protocol - In-situ and Immersion fixation - 2-3mm3 tissue blocks Initial Fixation at 4˚C - Ascending to Room Temperature

Schedule 3% Glutaraldehyde (0.2 M phosphate buffer, pH 7.4) Buffer Wash 1% OsO4 (buffered as above) Buffer or DH2O Wash (optional) 30% Ethanol 50% Ethanol 70% Ethanol 95% Ethanol 100% Ethanol (fill vials completely) 100% Ethanol (fill vials completely) Optional LN2 Cryofracture - between 1st & 2nd 100% EtOH - see description in text! 30% Freon TF/113 50% Freon TF/113 70% Freon TF/113 95% Freon TF/113 100% Freon TF/113 100% Freon TF/113 Critical Point Dry - in LCO2 Adhesive Mount on Aluminum Stubs Sputter Coat (Gold) - 50mT, 45mA

Time In

Time Out

75

76 ELECTRON MICROSCOPY: A HANDBOOK OF TECHNIQUES FOR THE BIOLOGIST

Chapter 7 - Alternative SEM Specimen Preparation A number of alternatives and enhancements to the “typical” biological soft tissue protocol described above, are available. The following are the most common alternate procedures. ✥ Uncoated Specimens Conductive samples require no coating, which even at 100Å thickness can obscure surface structures. Conductive samples should typically be imaged at high voltages (20-25kv) to take advantage of the increased signal to noise ratio (SNR) and yield a higher quality image. In order to view the non-conductive biological sample in an uncoated state, a low voltage beam (≤5.0kv) is required so as to minimize beam damage to the sample. Unfortunately, the use of a low voltage beam will drastically reduce the generation of the secondary electron signal from the surface and produce a poor image (low SNR). It is interesting to note however that even though the uncoated, low voltage image is of inferior quality in comparison to an identical coated sample imaged at high voltage, the amount information which is hidden through conductive surface coating is significant. In terms of uncoated biological samples, investigators have prepared them in a variety of ways for introduction to the SEM. In the simplest example, fresh specimens have been utilized. Such samples would have to be quite resilient in order to hold up under the high vacuum environment. Organisms with external shells and exoskeletons, such as insects, are good examples. In addition, cellulose containing botanical samples, including pollen grains, might be observed “fresh”. One should avoid samples which contain abundant water since dehydration surface tension artifacts will occur in addition to unacceptably long pump down times for the SEM. An alternative to ambient temperature fresh specimens is the use of a cryogen in order to freeze the sample prior to insertion in the SEM. Common cryogens employed include liquid freon 13 and liquid nitrogen. The sample, which is cleaned in water or buffer as necessary, is mounted onto a stub and plunged into liquid nitrogen. In order to maintain its frozen state, the SEM must be fitted with a cold stage which is held at liquid nitrogen temperature. It should be mentioned that this technique differs from freeze drying which is conducted under vacuum conditions with samples that are typically primary fixed in buffered aldehyde and cryo-protected in glycerol or dimethyl sulfoxide (DMSO) before being plunged into a liquid/solid cryogen “slush” and “dried” in a freeze drying unit under vacuum. The cryoprotectant will minimize damage induced through the formation of ice crystals in the extreme cold. In another alternative to uncoated specimen preparation, there exist a number of osmium enhancement agents which serve to concentrate additional osmium tetroxide into the soft tissue sample rendering it conductive, without the need for surface conductive coating.

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Examples of such osmium enhancement agents include Tannic Acid and Thiocarbohydrazide (TCH). Thiocarbohydrazide has been used successfully in this and many other labs in a protocol known as OTO and the alternative, OTOTO, with “O” indicating the osmium tetroxide solution and “T” indicating the TCH solution. In the OTO procedure, the typical primary aldehyde fix, buffer washes and osmium tetroxide (“O”) postfix is conducted. This is followed by a distilled water wash, then, 1% aqueous TCH (“T”) followed by 1% aqueous osmium tetroxide (“O”). This is followed with a distilled water wash and continuation of the usual protocol dehydration series. The OTOTO procedure adds an additional 1% aqueous TCH and 1% aqueous osmium tetroxide step to the OTO procedure. Critical point drying is conducted for soft tissues followed by adhesive mounting, but without the need for conductive coating. Samples can be imaged under high voltage conditions with good stability and signal generation. ✥ Cryofracture Technique Since the SEM is limited to the imaging of surface features, techniques have been developed that allow the viewing of internal structures. One of the most useful of these for the internal imaging of biological/cellular structures is cryofracture. In this technique, a tissue block is immersed in liquid nitrogen (LN2) for a few seconds and then fractured on a chilled brass or copper plate/disk using a pre-chilled, single-edged razor blade. A basic procedure which has yielded excellent results in this lab involves performing the cryofracture step between the two 100% ethanol changes of the dehydration series. The vial, maximally full of 100% EtOH and containing the tissue blocks are brought to the cryofracture station along with forceps, another vial filled with 100% EtOH (and labelled with the tissue type and a reference to cryofractured samples - for example, kidney (c), with the (c) designating samples which have been cryofractured), and a new acetone cleaned, single-edged razor blade. The cryofracture station (fig. 12) consists of the lower portion of a Petri dish, into which is placed a copper or brass (excellent heat conductors) disk and the clean razor blade. When ready, the Petri dish will be filled with LN2 to a point even with the top surface of the copper disk. If your lab has the large LN2 tanks with cryo hoses, the Petri dishes can be filled directly from the hose or by transferring a small amount of LN2 to a small dewar vessel for pouring into the dish. In this lab, we maintain a 10 liter dewar of LN2 which has an integral dipper for transferring small volumes of LN2 to the Petri dish. CAUTION: Liquid Nitrogen is extremely cold at -150˚C. It can cause instantaneous frostbite injury. Goggles must be worn to prevent eye contact and cryogloves are recommended. It should be noted that prolonged contact with the cryogen due to the Leidenfrost effect is more damaging to your tissues than a momentary surface contact with exposed skin. You would have a much greater chance of frostbite damage if the LN2 were to be trapped against the skin in a pair of gloves. NEVER touch anything which has been cooled with LN2!!

78 ELECTRON MICROSCOPY: A HANDBOOK OF TECHNIQUES FOR THE BIOLOGIST

The next steps should be performed quickly yet carefully as the liquid nitrogen evaporates rapidly. A tissue block is selected from the 100% EtOH vial using the jeweler’s forceps and submerged into the LN2 in the Petri dish. The block is held for a few seconds and then placed on the surface of the now chilled copper disk. The cold razor blade is used to fracture the tissue block using a deliberate vertical, downward force. The fractured pieces are then transferred to the empty 100% EtOH, cryo labelled vial using the cold forceps. These tissue blocks can be processed the rest of the way as usual. In alternative protocols to the above, a cryoprotectant such as glycerol or DMSO can be utilized. In a protocol devised by Tanaka (1981), DMSO was used as described below: Following the osmium tetroxide postfixation, samples are transferred to 25% DMSO (DMSO is diluted with buffer), then 50% DMSO for 30 minutes each. The samples are then cryofractured as described above, except that they are put into 50% DMSO after the cryofracture has been completed (not 100% EtOH). A buffer wash is performed followed by the usual dehydration series. Using this protocol, Tanaka was able to view well preserved internal cellular ultrastructure such as mitochondrial cristae and Golgi membranes. Fig. 12

FORCEPS & TISSUE BLOCK PETRI DISH

COPPER DISK

LIQUID NITROGEN

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✥ Microbial Specimen Preparation Microbes such as bacteria, protozoa and others present a unique challenge to preparation for observation under the SEM. The main problem involves our inability to observe these organisms with the unaided eye. How can we be sure that samples are present in the specimen vials? How can we avoid sucking them up with our pipettes while decanting off solutions? The most important developments in the field of microbial specimen preparation for SEM have involved the design and utilization of a number of containment strategies for the organisms of interest. A variety of commercial and homemade sample holders have been used over the years. Keeping containment in mind, it is obviously critical to have some idea as to the dimensions of the organism you wish to fix. In the past, both nucleopore and millipore filters, along with polycarbonate membranes, have been used for such specimens as bacterial cells (~0.25 - 0.75µm diameter). Suspensions of these cells can be concentrated on the filters by passage through a syringe/ filter holder unit which is commercially available. The filters can be removed and passed through the various fixatives and into the CPD. After drying, via CPD or air dried through the organo-silicon compounds described earlier (TMS or HMDS), the filters may be cut to fit on the stub surface (conversely, large stubs may be utilized) and glued and conductive coated. As an alternate, bacteria can be attached to a polylysine coated cover slip which is then passed through the various fixation, drying, mounting and coating steps. Since you will mainly working with single cells, the various protocol durations can be reduced considerably. Protozoa are considerably larger than bacterial cells and are contained in a variety of ways for fixation. Many individuals have modified large and small BEEM capsules as microbial containment vessels. A hole is cut into the cap and fine mesh filter is put in place after loading the cells. The loaded BEEM capsule is placed into a vial for passage through the various fixatives. Diffusion of fixation agents occur across this microbial barrier. A convenient choice of a diffusion barrier is Nitex mesh which is available in a number of mesh sizes. For most protozoa, a 40µm mesh is adequate. Nitex mesh, also known as bolting cloth, is a nylon material which is purchased by the yard. Preparation of Nitex Bags In this lab, a 40µm mesh Nitex bag is fashioned to contain our protozoan cells. Approximately 2" squares of Nitex are cut out and folded in half. An old pair of forceps is used to hold the free edges of the folded cloth together with about 1.0mm of the edges extending beyond the forceps. A bacterial transfer loop is heated in a flame and run along the tightly held edges of the Nitex cloth. The excess 1.0mm edge is cut off while simultaneously sealing the free edges of the cloth. The cloth now has two open ends, one of which is sealed in the identical manner by rotating the cloth square 90˚ and holding one of the two open edges in the forceps. The edge is sealed and the bag now has one open end into which the protozoa are loaded. Nitex bags should be prepared in advance of the fixation. In addition, a number of micropipettes should be made before the actual time of

80 ELECTRON MICROSCOPY: A HANDBOOK OF TECHNIQUES FOR THE BIOLOGIST

fixation by pulling standard Pasteur pipettes in a flame. The micropipettes prevent the transfer of large volumes of culture media and fixatives during the fix process. Microbial Protocol As mentioned earlier, due to the size of the typically single cells, the duration of fixation steps is greatly reduced as compared with large tissue blocks. For protozoa, a Quick Fix method is employed which involves mixing buffered Glutaraldehyde and aqueous Osmium Tetroxide. From the chapter on TEM specimen fixation, you will remember that glutaraldehyde and osmium tetroxide react to form an undesirable precipitate. It is permissible to mix these two fixatives in this Quick Fix since the samples will be removed (approximately 15 min.) before the precipitate reaction has become a problem leading to the formation of surface artifact. The Quick Fix is carried out in the wells of transparent glass blocks, under a stereomicroscope. The stereoscope is placed under and at the front of a fume hood to reduce the chance of vapor contact with investigator eyes. Multiple wells of a white porcelain plate are filled with buffer solution for transfer into after the quick fix. The initial solutions required are: • 3% Glutaraldehyde (in 0.1M phosphate buffer, pH 6.8 - match to culture medium) • 0.1M phosphate buffer, pH 6.8 (for washing) • 4% aqueous OsO4 (purchased in sealed ampoules) A good log phase (high density growth) protozoa culture is desirable. Using a 1.0mm serological pipette, 0.5ml of the culture is transferred to a microfuge tube and placed into a microcentrifuge head. A tube of water is marked and placed in the head opposite the culture tube for the purpose of balance. The microfuge timer is advanced to provide a burst of speed but without engaging the timer to even 1min. duration (which may be too forceful and could damage your cells at 18,000 x g). When the head has stopped, the culture tube is removed and the cells should be concentrated at the bottom. The Quick Fix solution should be mixed just before the culture tube is spun down. It is prepared by mixing 2 drops of the buffered 3% Glutaraldehyde to 1 drop of the aqueous 4% OsO4 in the glass depression well which sits on the stage of a stereomicroscope, under the hood. If a fume hood is unavailable, a fan should blow across the microscope stage to prevent fumes from coming up into the face of the investigator. Do not contaminate solutions by using the same pipette for mixing! Once the Quick Fix is mixed, a micropipette bulb is depressed and inserted gently to the bottom of the culture tube. The bulb is released and a large number of cells should enter the micropipette. The contents of the micropipette are “shot” into the quick fix and allowed to remain for 15 minutes and no longer! The use of white photographic mounting board under the glass well block will enable you to easily observe the fixed and darkening cells under the stereoscope. As they fix, the cells will concentrate at the center bottom of the well as they become impregnated with the high density heavy metal, osmium. As cells in the first well fix, other wells in the block may be prepared as the number of samples you require dictates.

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Once 15 minutes has elapsed, a fresh, clean micropipette is used to transfer fixed cells to the buffer solution in the wells of a white porcelain plate (white is ideal since dark fixed cells will stand out in contrast against the white porcelain background). Once again, pressure is held on the micropipette bulb as the tip is advanced into the bottom of the fixation well. You must view this under the stereoscope so as to carefully guide the tip to the area of concentrated cells. You must attempt to avoid transferring excessive fixation mixture to the buffer and prevent a precipitation reaction! With some practice, you will readily suck up the fixed cells from the bottom of the well with a minimum of fixative. Depending on the number of cells initially transferred into the fixation well, you may perform a number of transfers out of that well and into a number of buffer wells using the same micropipette. At this point you should concentrate on getting all of your fixed cell out of the quick fix mixture wells and into the buffer wells. The buffer well plate can now be taken out of the hood to the lab tabletop and placed on the stage of another stereoscope. Specimen vials are filled with buffer solution and a previously prepared Nitex bag is inserted into the vial and buffer solution so that the open end sticks slightly out of the vial. A slipknot lasso of white (no pigments which may come out in the EtOH or freon series) sewing thread is looped over the open end of the Nitex bag. A new, clean micropipette is used to transfer at least 20 to 40 cells into the open Nitex bag. The cells are shot down into the lower portion of the bag submerged in the buffer solution to prevent premature drying down and surface tension collapse. The lasso thread is tightened in order to seal the opening and the remainder of the typical soft tissue protocol continues with a reduction in the duration of each step (see below). To prevent damage by pipettes to the Nitex bags and escape of its cellular contents, solutions are decanted off by pouring them out. New solutions are pipetted in from above the bags. It is unlikely that the cells will dry down between protocol steps since the bags retain adequate moisture. Once through the intermediate fluid series, the bags are pulled out of the vials and laid into the freon TF/113 filled channels of the CPD boat. The wire mesh covers are placed over each channel for protection and the CPD run progresses as usual. While the CPD run occurs, clean stubs can be prepared with glue transfer tab adhesive coatings. Upon removal from the CPD boat, the cells within the Nitex bags are dry and very delicate. Holding the bag upright (thread sealed end UP), the bag is lightly tapped on the side to concentrate cells at the bottom. The thread is pulled off or the top of the bag is cut off. At the now open end, an adhesive prepared stub is inverted and placed at the opening of the bag. The bag is inverted and lightly tapped again. Dry cells should sprinkle down on the adhesive surface and stick. This can be confirmed with a stereo microscope view of the surface of the stub. The stub can now be conductive coated as usual (consider this a relatively flat surface and coat for 30 sec.). In addition to the above “sprinkling” procedure, some cells may be trapped in the Nitex mesh. Once again, this can be observed under the stereoscope. In this case, the Nitex mesh can be opened and glued directly to an adhesive coated stub. The excess Nitex can be trimmed at the edge of the stub and the stub can be sputter coated. It should be noted that there are many variations to the above microbial techniques in the literature. These Nitex bags have proven very successful in this lab.

82 ELECTRON MICROSCOPY: A HANDBOOK OF TECHNIQUES FOR THE BIOLOGIST

In addition, we are usually not concerned about cell orientation on the stub surface. For some investigations, this orientation is critical. Once on the surface of a stub, cells may be repositioned using a microneedle which is prepared by pulling a capillary tube in a flame and allowing the end to seal. It is best if the cells do not contact adhesive prior to repositioning by keeping them in the Nitex mesh as opposed to sprinkling them out randomly onto an adhesive coated surface. The microneedle may be placed into a glue such as Pliobond so that the cells adhere to the needle for specific orientation and final placement (dorsal vs. ventral, etc.). The entire microbial protocol is listed below and a blank schedule sheet is provided on the following page: • Quick Fix - 15 min. • Buffer Wash - 2 X 5-10 min. • 30%, 50%, 70%, 95% EtOH - 5 min. each • 100% EtOH - 2 X 5 min. • 30%, 50%, 70%, 95% Freon TF/113 - 5 min. each • 100% Freon TF/113 - 2 X 5 min. • Critical Point Dry (or alternative) • Adhesive Mount on Stubs • Sputter Coat with Gold (30 sec. @ 50mT and 45mA) • Store in Desiccator Microbes make very interesting and rewarding samples for examination under the SEM with preparation of living samples being conducted in less than half the time required for bulk tissue samples. ✥ Correlative SEM to TEM Sample Preparation It is often desirable to correlate images generated by a variety of instrumentation, such as LM, SEM and TEM, since their resolution ranges overlap. Sample blocks prepared for the SEM can be taken and prepared for conventional TEM. It is important that the soft tissue blocks be small (no larger than 1.0mm3) at the outset of SEM preparation for reasons of fixative penetration. Once a sample has been viewed and photographed using the SEM, it can be gently removed using acetone (Pliobond glue is readily dissolved using acetone and is a good SEM adhesive choice if one is contemplating this procedure). The tissue blocks are transferred to propylene oxide (3 changes for 10 minutes each) and embedded in epoxy resin using conventional techniques (see Unit 1).

UNIT 2 – PREPARATION OF BIOLOGICAL SAMPLES FOR SEM

Microbial Fixation Schedule Worksheet SEM Microbial Protocol - Immersion fixation of Single Cells Initial Fixation at 4˚C - Ascending to Room Temperature

Schedule Quick Fix (2 parts 3% Glutaraldehyde in buffer to 1 part 4% aq. OsO4) Buffer Wash Buffer Wash 30% Ethanol 50% Ethanol 70% Ethanol 95% Ethanol 100% Ethanol (fill vials completely) 100% Ethanol (fill vials completely) 30% Freon TF/113 50% Freon TF/113 70% Freon TF/113 95% Freon TF/113 100% Freon TF/113 100% Freon TF/113 Critical Point Dry - in LCO2 Adhesive Mount on Aluminum Stubs Sputter Coat (Gold) - 50mT, 45mA

Time In

Time Out

83

84 ELECTRON MICROSCOPY: A HANDBOOK OF TECHNIQUES FOR THE BIOLOGIST

Unit 3 - Black & White Photographic Principles in Electron Optics The black and white electron photomicrograph is the final technical product of the electron microscopist. Since the TEM and/or SEM image disappears when the filament is deactivated, it becomes a necessity to capture a permanent image for the purpose of detailed analysis. Silver-based black & white photography has satisfied this requirement for many years. Even though many EM labs have converted to digital image capture, storage and analysis/manipulation through the use of personal computers (PC’s), most electron microscopists still rely on photographic techniques in the course of their studies. From earlier discussions on electron optical principles you should recall that in order to achieve the highest possible resolution, voltage stabilization circuitry is employed which results in an electron source with a single constant wavelength. This monochromatic source reduces the effect of chromatic aberration which would limit resolving power. The constant electron wavelength means that “color” electron images are impossible (unless they are pseudo-colored using a computer or hand colored - Dr. Martin’s dyes yield excellent results in hand coloring of EM images when applied directly on photographic prints, especially SEM). The quality of the final photomicrograph is dependent on all of the tedious steps that preceded its production. An individual experienced in EM techniques need only look at a micrograph to determine flaws in the technique of the individual, whether the flaws resulted from poor tissue fixation, tissue embedment (TEM), ultrathin sectioning (TEM), post-staining for TEM/conductive coating for SEM, EM alignment and operation or poor photographic technique. Careful attention to detail at each stage in the process will usually produce quality results. It should be noted that the resultant photomicrograph will typically show more detail that the TEM or SEM viewing screen image. This is due to the fine-grained characteristics of most EM films in comparison to course-grained TEM fluorescent view screens and low resolution, 625 line, SEM monitors/CRT’s. Black & white photography involves a two step process: 1. Negative production (exposure and processing) 2. Enlargement Printing (exposure and processing of photographic paper using a negative in an enlarger)

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Chapter 8 - Film and Paper Composition Modern films are composed of four (4) important layers (fig. 13). The upper layer is a protective anti-scratch layer which is usually dull/non-reflective in appearance. Below this is the most vital layer to the process of photography, the emulsion layer. In this layer, silver halide (such as silver bromide) crystals/grains are suspended in a gelatin matrix some 25.0µm thick. In this layer, the photochemical reactions required to produce a permanent, high resolution image take place. Under the emulsion layer is a support layer commonly composed of cellulose acetate (commercially known as “estar”). The final layer is known as the anti-halation layer which prevents photon/electron backscatter to the emulsion layer. This layer is shiny in appearance, therefore, if you examine a sheet of film you should notice a dull/emulsion side and a shiny side. Most manufacturers (Kodak) will cut a small notch in the film to allow one to determine the emulsion side of the film even in total darkness - the emulsion side is up when the film is held with the notch in the upper right corner (fig. 14). The emulsion side of photographic paper is easy to determine since it is highly glossy under the bright yellow or amber safelight used in printing. Fig. 13 Film Composition:

anti-scratch - dull side Emulsion (silver bromide/gelatin) Support/Base (cellulose acetate) anti-halation - shiny side

Fig. 14 Film Notching Guide:

emulsion side is up

86 ELECTRON MICROSCOPY: A HANDBOOK OF TECHNIQUES FOR THE BIOLOGIST

✥ Emulsion The effect of electrons is similar to that of photons on light sensitive emulsions. As stated earlier, the emulsion layer is the active site of the photographic process wherein a chemical reaction occurs. The emulsion layer is approximately 25.0µm thick and is composed of usually uniformly sized silver halide (such as silver bromide - Ag+Br-) crystals suspended in a gelatin matrix. Depending on the film, the crystals can range in size from 0.1 to 10µm in diameter. This crystal diameter relates to the film speed as shall be seen. For film, this emulsion layer is coated on a cellulose acetate support or base. For paper, the emulsion is coated on cellulose-based paper (Kodak Kodabromide paper) or coated on a resin (Kodak RC paper). When an energy source such as electrons or photons encounter a crystal, the crystal is disturbed/rearranged in various regions. The crystal is said to be in an activated state, however, an actual image cannot be seen at this point. A latent image exists which theoretically can persist for years. This latent image must be converted to an actual image through processing of the film or paper. The chemical reaction that occurs when electrons/ photons encounter a crystal is reduction, the gain of electrons. Relative to a gain of electrons, the reduction reaction can be summarized as follows: Ag+ + e- ➙ Ag˚ In this case, silver ions are reduced to silver atoms. The main difference between electron versus photon effects on the emulsion is that electrons are 100% quantum efficient. This means that a single electron is capable of activating a single silver halide crystal whereas, it takes 10 to 100 photons to activate a single crystal. Therefore, no true film speed exists with respect to electron effects on light sensitive emulsions. Since this is the case, the electron microscopist can take advantage by using very fine-grained films for the highest resolution. This is true only when electrons bombard the film to form the image as in the case of the TEM. In SEM, photons from a high resolution photo-CRT are activating the emulsion. It should be indicated that a single silver halide/bromide crystal is composed of a large number of individual ionic bonded Ag+Br- molecules. During the reduction reaction, the crystalline lattice is rearranged yielding the latent image. ✥ Film Speed (ISO/ASA) The film speed simply refers to the diameter of the silver halide crystals suspended in the emulsion layer. Fast films (400, 1000) have large crystal diameters. These large crystals are more likely to encounter and react with photons under conditions of low source (photon) density. If you have limited light conditions (indoors without a flash) you will need a fast film in order to record an image. The resultant enlargement prints are very grainy and of low resolution due to the large crystal size. Slow films (25,100) require an adequate photon concentration since they are very fine-grained. The fine-grained films

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result in high resolution prints even at high magnifications. Since electrons are 100% quantum efficient relative to the silver crystals, we can take advantage of the slow, fine-grained, blue-sensitive films available, such as Kodak Electron Image Film 4489 with an estar base and an ASA of 15. This 3.5" by 4.25" sheet film permits excellent enlargements of high resolution. As it is blue-sensitive, the same safe light used in enlargement printing, yellow or amber, can be used when handling this film. Light sensitive emulsions, even when used to capture electron generated images, are typically sensitive to certain wavelengths/colors of the visible spectrum. Safelights can be used to take advantage of this fact and allow the individual to handle films and papers under some lighting conditions. When choosing the appropriate safelight filter, the wavelength selected must be below the threshold energy necessary to activate the silver halide crystals of the emulsion. If the film is blue-sensitive, a yellow or amber OA or OC safelight filter is appropriate. If the film is yellow-sensitive, a yellow safelight filter would provide the necessary energy to activate the emulsion. In this case, a red filter could be used. Bear in mind that along with the appropriate filter, a low 10-15watt bulb is necessary at a minimum distance of 4 to 5 feet. ✥ Supports (Base) Film In the past, emulsions were coated onto breakable glass plates. The advantage of glass is that it contains no water and requires no desiccation prior to insertion into the TEM. This is not important in the SEM since images are taken off the high resolution recording CRT at atmospheric pressure. The modern support used for film is cellulose acetate or “estar”. Although the estar plates are easier to handle and not subject to breakage, they must be handled carefully to avoid dust, fingerprints and scratches. The cellulose acetate contains water and must be desiccated prior to insertion in the TEM. Most modern TEM’s have a plate dryer which is evacuated by a rotary pump. Paper Paper for enlargement printing is manufactured by coating an emulsion layer on either cellulose paper (Kodak Kodabromide) or resin (resin-coated or RC paper). Since the processing chemicals are absorbed by the cellulose paper, it requires longer processing times, especially the final wash in running water of up to 60 minutes. With RC paper, only the emulsion is affected resulting in shorter processing times. By example, the final wash time is reduced to 4 minutes. For drying, the RC paper should be treated for removal of excess water using a sponge or squeegee and allowed to air dry. By contrast, the cellulose paper should be dried in a heated drum or plate dryer, allowing the print surface to contact the shiny ferrotype surface for a high gloss final print. Papers are designated by both letter and number combinations. The letters refer to the

88 ELECTRON MICROSCOPY: A HANDBOOK OF TECHNIQUES FOR THE BIOLOGIST

type of print surface such as matte or glossy. For scientific publications, the glossy surface is most desirable and is noted by the letter “F”. The numbers (usually 1 through 5) indicate the paper grade or contrast level of the paper. Paper grades will be covered in more detail under the topic of enlargement printing. ✥ Routine Photographic Films and Papers Used For TEM and SEM Films TEM: Kodak Electron Image Film 4489 (3.25 x 4", ASA=15) - OA or OC safelight. SEM: Kodak Commercial Film 4127 (4 x 5", ASA=50) - A1 (red) safelight. Polaroid Type 55 P/N Film (4 x 5" positive and negative, ASA=50) - no safelight required. Paper For use in the enlargement printing of any of the negatives listed above: Kodabromide F, 1-5 (fiber based, single or double weight) - OA or OC safelight Kodak Polycontrast F - RC (resin coated, requires a Polycontrast Filter Kit) - OA or OC safelight

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Chapter 9 - Processing (Films and Papers) Processing involves a series of steps that will transform the latent image, produced via exposure of the film or paper, into an actual image made up of dark, light, and gray (halftones) regions, namely, contrast. The image will also be fixed and preserved on the film or paper by these procedures which include: Development, Stop Bath, Fixation, Washing and Drying. The ideal way to process negatives involves loading the exposed film into the appropriate sized open frames/holders and pass them through the required solutions which are stored in tanks with floating lids (fig. 15). Papers are usually processed in various sized trays with washing carried out in a unit which has an attachment for circulating water, such as a large tub which houses a rotating drum (fig. 16). It is important that the films and papers be agitated throughout the entire process. Agitation reduces the chance that air bells (bubbles) on the surface will prevent necessary contact with the various processing solutions. Figs. 15 & 16 – Film & Print Processing Stations Illustrations

Negative Processing Station - Tanks

Developer (D-19) 1:2

Stop Bath Water

Rapid Fixer straight

1.5 min.

2-3 min.

4 min.

H

C

Running Water Wash 20-30 min.

NOTE: Times and dilutions indicated pertain to Kodak 4489 TEM Film

Print Processing Station - Trays

Developer (Dektol) 1:2

Stop Bath with Indicator

1-2 min.

5-30 sec.

Rapid Fixer 1:1

H

Running Water Wash

2 min.

NOTE: Times and dilutions indicated pertain to Kodak RC Paper

C

4 min.

90 ELECTRON MICROSCOPY: A HANDBOOK OF TECHNIQUES FOR THE BIOLOGIST

✥ Development Development is the most critical step in processing. Under-development produces an image of poor, “muddy” contrast and low grain density (the image will appear too light). Conversely, over-development will result in “fogging” and produce a dark image (high grain density) of poor contrast. In development, the latent image on the film or print is essentially soaked in an “electron bath”, specifically, a reducing agent which will give up electrons to the silver atoms (which were initially reduced from silver ions during exposure to the source electrons or photons). During this “electron soak”, reduction of silver atoms to black metallic silver occurs. The latent image is transformed from activated crystals to actual black grains which yield contrast and a true, observable image. This reduction to black metallic silver begins at a “nucleus” in the activated crystal and spreads. Evidence of this can be seen by examination of these silver grains under the TEM as is done in TEM autoradiography. At TEM resolution, the grains appear elongated and worm-like versus their circular, dot-like appearance under the light microscope. In order to prevent inactivated crystals from being reduced, leading to fogging, development must proceed under very specific times and temperatures. The main components of a developing solution are as follows: Hydroquinone - the reducing agent. It requires an alkaline environment (pH 10-11) in order to donate electrons. Borax or Sodium metaborate - provides the required alkaline pH 10-11. Sodium sulfite - preservative, prevents air oxidation of developer, lengthens shelf life. Potassium bromide - slows development, serves as a competitor of the reduction reaction, allows for control over the development process. Typical developers include D-72 (Dektol) which is used primarily for papers and D-19, a high contrast developer used for films in scientific applications and x-rays. These developers are available in powdered form and are mixed into one gallon of water at 90120˚F. A good working solution has a characteristic odor, no precipitate/sediment, is clear with a slight yellowish color, and is slippery/slimy to the touch due to the alkaline pH. A stock solution in a tightly capped bottle has a shelf life exceeding one month. Most working developer solutions are diluted from the initial stock solution. Development times and temperatures must be carefully monitored to ensure that only activated silver grains develop. Development parameters for common TEM/SEM films and papers are as follows: Film TEM Film 4489 SEM Film 4127 SEM Polaroid 55

Developer Dilution Time Temperature D-19 1:2 4 min. 20˚C D-19 1:1 4 min. 20˚C Instant processing - no developer required

UNIT 3 – BLACK & WHITE PHOTOGRAPHIC PRINCIPLES IN EM

Paper Kodabromide Polycontrast RC

Dektol Dektol

1:2 1:2

1-2 min. 1-2 min.

91

20˚C 20˚C

One should be careful not to pull a developing print out of the solution too early. Under an OA/OC safelight, the image appears much darker than under normal white light conditions. Since you actually observe the image develop, the tendency for the novice is to move the print out of the developer before the recommended times as listed above. If the print is too dark after the minimum time has elapsed, adjustments to exposure time or fstop will be necessary. ✥ Stop Bath As indicated by its title, the stop bath stops the development process. Typically, the stop bath is a 1% acetic acid solution which contains the pH indicator, Bromthymol Blue. Kodak markets this solution as Indicator Stop Bath; it is mixed with water in a typical dilution of 1:62 (16ml per liter of water). The purpose of the acetic acid is to neutralize the alkaline development solution (unless the pH is 10-11, hydroquinone will not act as a reducing agent). The bromthymol blue is used to gauge the acid-base neutralizing capacity of the stop bath. At a pH below 7.0, this indicator appears yellow and above pH 7.0, it appears blue. Under the OA/OC safelight, the solution will darken to indicate that it is no longer effective in stopping development and should be replenished. If it is changed often, water can also serve as an effective stop bath and is recommended for the TEM films (Kodak 4489) since the production of a mottled appearance on the films has been reported with the use of indicator stop bath. The water in the film development tank should be changed after each group of films (a quantity 8 to 10, in their holders) has been passed through. Kodak 4127 film can make use of the indicator stop bath, however, if the same tanks are used for SEM and TEM processing, water will suffice. For paper, the use of indicator stop bath mixed into a development tray of water is recommended. Stop Bath parameters for common TEM/SEM films and papers are as follows: Film TEM Film 4489 SEM Film 4127 SEM Polaroid 55

Stop Bath Time Water 1.5min. Water or Indicator 30 sec. Instant processing - no stop bath required

Paper Kodabromide Polycontrast RC

Indicator Stop Bath Indicator Stop Bath

30 sec. 30 sec.

Temperature 20˚C 20˚C

20˚C 20˚C

Almost continuous agitation in the stop bath is recommended due to the short time duration required.

92 ELECTRON MICROSCOPY: A HANDBOOK OF TECHNIQUES FOR THE BIOLOGIST

✥ Fixer Photographic fixer, not to be confused with the fixation of biological samples, serves three important purposes. Firstly, and most importantly, it contains a component which releases inactive silver halide from the emulsion layer. This component, commonly known as Hypo, is actually Sodium or Ammonium thiosulfate (or thiocyanate) which reacts with the inactive, ionic bonded silver halide to yield soluble silver thiosulfate/thiocyanate. In other words, the hypo allows for the inactive crystals remaining in the emulsion layer to dissolve into the fixer solution. In large photographic processing labs, the fixer may be collected and sold back to the film manufacturer who extracts the silver halide to be used in the manufacture of new film. The second ingredient is potassium alum which hardens the gelatin matrix of the emulsion to protect and preserve the image. This component is often referred to as the hardener. The final ingredient of fixer is an acid, usually acetic or sulfuric, which aids in the hardening process and also can neutralize any developer carried over from the stop bath. Fixer is sold in powdered form (Kodak Fixer) or liquid form (Kodak Rapid Fixer) which uses the ammonium thiosulfate based hypo. When preparing the stock solution, water temperature is critical. If not adhered to, the final solution will appear cloudy and may not clear upon standing. The stock solution is usually prepared in one gallon capacity bottles and used straight for films, and in the case of rapid fixer, diluted 1:1 for papers. To prepare a gallon of stock rapid fixer for films, approximately one-half gallon of water at 16-27˚C is added to the bottle. The entire contents of solution A is slowly poured into the water, followed by solution B (caution: strong acid) added in small volumes followed by agitation. Water is finally added to bring the total volume to one gallon, followed by mixing. The final solution has a pungent odor (acidic), is clear and colorless and has a long shelf life (up to three months or more). To check the effectiveness of the fixer solution, hypocheck, a 4% potassium iodide solution can be purchased or prepared by mixing 4.0g of KI into 96ml of distilled water and dispensing into dropper bottles. To test the fixer, two drops of the hypocheck are added to the fixer solution. If a milky white precipitate appears, the solution is exhausted and should be discarded. During fixation, materials should be agitated and the normal white room lights can be restored half way through the process (i.e. if a 4 minute fix is called for, lights can be turned on after 2 minutes). For Polaroid negatives (Type 55 P/N), the company recommends the use of a sodium sulfite solution in order to clear the film, for 4 minutes duration. Straight fixer can also be used with satisfactory results.

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Fixation parameters for common TEM/SEM films and papers are as follows: Film TEM Film 4489 TEM Film 4489 SEM Film 4127 SEM Film 4127

Fixation Fixer Rapid Fixer Fixer RapidFixer

Dilution straight straight straight straight

Time 8 min. 2-3 min. 8 min. 3-4 min.

Temperature 20˚C 20˚C 20˚C 20˚C

SEM Polaroid 55

Fixer

straight

4 min.

20˚C

Paper Kodabromide Kodabromide Polycontrast RC

Fixer Rapid Fixer Rapid Fixer

straight 1:1 1:1

8min. 8min. 2 min.

20˚C 20˚C 20˚C

✥ Washing The purpose of washing the processed photographic materials in water is to remove any of the processing chemicals from these materials. If the chemicals are not washed off and out of films and papers, especially the fixer, these materials will brown with the passage of time. It is important to adhere to the specific times and temperatures for washing, especially for fiber-based papers. Over-washing a fiber-based paper may result in creasing and possible tears as the paper is dried. Colder temperatures will require longer wash times with the ideal temperature at 20˚C. Washing should always be done in a circulating water bath with constantly running water. Films can be left in overflowing tanks and papers can be placed in specially designed paper washers, such as large tub washers with rotating drums and upper and lower drains. Washing parameters for common TEM/SEM films and papers are as follows: Film TEM Film 4489 SEM Film 4127 SEM Polaroid 55

Wash Water Water Water

Time 20-30 min. 20-30 min. 10 min.

Temperature 20˚C 20˚C 20˚C

Paper Kodabromide Polycontrast RC

Water Water

30-60 min. 4 min.

20˚C 20˚C

94 ELECTRON MICROSCOPY: A HANDBOOK OF TECHNIQUES FOR THE BIOLOGIST

✥ Drying Drying of photographic materials, especially negatives, should be done in as dust-free an environment as possible. After washing, films should be dipped into a 1:200 solution of Photoflo (Kodak), which is a wetting agent to promote even drying and the absence of water spots on the negatives. Films are then hung on some type of “clothesline” to air dry. After films are dry, they should be protected in glassine envelopes. Kodabromide, fiber-based papers, are dried in a heated flat plate or drum dryer with a shiny ferrotype surface. With the print face-up, a roller squeegee is used to remove excess wash water. The print is then positioned so that the printed surface contacts the hot ferrotype surface which yields the desired glossy surface (with F type papers). The paper is rolled into the drum dryer with typically a canvas overlay. In the plate dryer, a canvas covering is stretched over the back of the print(s). After about five minutes, the drum can be unrolled (or the canvas cover pulled back) and the prints will simply separate themselves from the ferrotype surface. The ferrotype should be routinely be cleaned with the appropriate ferrotype polish (available at photographic supply stores). Resin-coated (RC) papers are simply wiped with a soft photographic sponge or squeegee and allowed to air dry with F papers yielding a glossy surface.

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Chapter 10 - Negative Handling and Exposure (TEM & SEM) In general, unexposed negatives (film) for TEM and SEM must be loaded into some appropriate light-tight carriers for insertion into the “camera” of the microscopes. As discussed earlier, films are to be handled carefully under the appropriate safelight conditions. The film sheets should be held on their edges to avoid the transfer of oily contaminants from the skin and the resultant area of poor exposure and processing. The following section will describe the loading and exposure of films for the TEM and SEM generally, and the Hitachi HS-8 TEM and the Hitachi S-2400 SEM, specifically. ✥ TEM - Hitachi HS-8 In the case of the TEM, films must be loaded directly into the vacuum environment of the microscope, usually in a camera chamber located below the viewing screen. When the screen is pulled out of the way, the imaging electrons will now contact the photographic emulsion and expose it, producing a latent image. Water-containing, estar based films must be desiccated prior to loading and insertion into the TEM camera chamber. Otherwise, you can expect lengthy pump-down times and contamination/corrosion of internal TEM components. As stated earlier, modern TEM’s will usually come with a plate dryer. Films should be dried at low vacuum for at least one hour prior to loading. The modern TEM will usually have a light-tight storage/loading box (for storage of multiple unexposed film plates in the microscope) and a light-tight receiver box (for collection of exposed plates) that will be placed into the TEM camera chamber. Within these boxes, the film plates are held, often in flat metal plate holders. For the Hitachi HS8, 18 film plates can be loaded into a single storage/load box. A single sheet of film, emulsion side up (notch in the upper right corner), is placed into the metal holder and a rectangular metal frame is added to keep it in place. It is important that the flat side of the frame be placed down against the film and the beveled side face up – otherwise, the film holder might get stuck in the motorized camera mechanism. The lower edge of the frame is pushed into the spring arrangement at the base of the metal holder, while the upper edge of the frame is guided and held in the two, L-shaped, upper catches. The loaded plate holders are then stacked into the storage/load box, whose entire front sliding door is removed, with the plate holder central notch pointing out. Once again, the box holds 18 plates at the maximum. Do not overfill the storage box! Once full, the storage box can be put into the desiccator/plate dryer for a minimum of one hour at low vacuum. A desiccated storage box can then be transferred into the TEM camera chamber. In order to do this, the TEM must be on and at high vacuum. The camera airlock must be closed and air admitted into the camera chamber through the camera door. The cover for the load box site is lifted straight up and off. If still in place, the old storage box is removed from the TEM and the new box put in its place. The arrow on the front door of the box and the “F” (for front) must face the front of the TEM upon loading into the TEM camera chamber. When in place, the front and rear doors of the box are slightly raised

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so that the single lowest plate holder can be advanced forward under the TEM view screen by the motorized plate drive. Before placement of the storage box into the TEM, the black blanking plate should be removed since they have become stuck in our particular scope in the past - this necessitates performing the insertion process under safelight or dark conditions! A full receiver box should be removed and replaced with an empty one. These receiver boxes can be found behind the front camera door. They have a front handle and a hinged top lid which should be opened only under safelight conditions, when you are ready for processing. When the two camera chamber doors are replaced, the chamber is rough pumped using the rotary pump (CAMERA button), followed by deselecting the CAMERA button (usually for PLATE) and opening the camera airlock. ✥ HS-8 Camera System and Film Exposure The camera system and film exposure mechanism of the Hitachi HS-8, as for other modern TEM’s, is essentially automatic. As the TEM operator, you must ensure a number of conditions are met prior to activating the automated camera system for the best possible image capture. For example, if your image is under or over focused, the auto camera system will provide an excellent exposure of an out of focus image. This brief section will provide a step by step procedure for capturing a properly exposed image of high resolution and optimal contrast. Initially, it is important to understand the basic workings of the TEM camera system. Within the lower region of the TEM column (observable through the small, lower circular viewport of the HS-8) four CdS photocells are located in the peripheral electron beam path. When energized, these photocells will begin to collect electrons (or photons) up to a certain sensitivity set point. At this set point, a metallic shutter mechanism is set in motion which effectively blocks the electron beam from contacting the film emulsion. The exposed film can be moved into a light-tight receiver box and later removed for processing. All of these events will occur in the high vacuum of the TEM column. As stated earlier, films for TEM must be desiccated prior to their insertion into the microscope. The actual procedure of high quality image capture for the HS-8 follows: 1. Scan your grid at low magnification (1,000-2,100 X) and locate a section and area of interest. You will need to magnify, center, adjust brightness and focus in order to locate an area worthy of an exposure. Avoid sections and areas containing obvious contamination and sectioning artifacts. 2. Determine the magnification you will be exposing the film at. Important: if you change the magnification you will have to perform all of the steps which follow for the new mag setting! 3. Focus the image using high brightness and the coarse and fine objective lens focus controls. The screen should be tilted and the 10X binoculars utilized for this process - the

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binocs should be focused on the screen crosshair. If a small hole exists in the vicinity of the area of interest, it is ideal as a focusing aid. The presence of interference fringes inside or outside the hole indicates a condition of underfocus or overfocus, respectively. Note: to the human eye, the underfocus image appears to be in best focus - this is the point of maximum contrast/granularity, also known as defocus contrast. The human eye responds to the high contrast as the most desirable image. True focus is achieved when no interference fringe is observed in conjunction with the hole - although not as high in contrast, true focus yields the highest resolution. 4. Frame the area of interest using the stage (X,Y) controls - note the small central framing box on the main view screen. Note: the view screen must be in its flat, not angled, orientation for this framing process since the film will lie flat beneath the screen. 5. Activate the Exposure Meter by depressing the button in the right auxiliary drawer. In order for the camera and auto exposure mechanism to function properly, the camera airlock must be opened - a small square status indicator will light, a box imprinted with the #0, on the left main panel. 6. Using both the Brightness knob (condenser lens current) and the electric deflection coil Bright X, Y, spread out the beam to the screen periphery. This step is critical since the photocells are located in a peripheral position. If the beam intensity is central, the photocells will not be contacted directly and begin to collect the electron energy necessary to close the shutter. An overexposure will result. The photocells must also be evenly illuminated for optimum performance of the automated system. This is accomplished using the Bright X,Y in conjunction with the Brightness control. 7. Under “normal” conditions, the photocell sensitivity knob (the large black knob on the right main panel - labelled Exposure Meter) should be set to #5. By normal, I mean that none of the photocells are blocked by a grid crossbar(s) - this situation would cause the three exposed photocells to do the work of four leading to an overexposure as the film is contacted by the electron beam for an excessive time period. In an alternative situation, a section may be slightly torn away from a grid bar edge and directly over a photocell. The torn area will allow maximum electrons to contact this photocell and close the shutter prematurely - the result is an underexposure. Under these abnormal conditions, the photocell sensitivity knob would have to be adjusted from the #5 position. As you go up to #6, #7, the sensitivity decreases for exposures taken with an intruding gridbar. The sensitivity increases as you go down for exposures taken with tears in the resin. 8. At this point, an automatic exposure could be attempted, however, you may be wasting a tremendous amount of film if you do not perform this simple exposure time check before loading a piece of film under the view screen. To test the exposure time, lift the chrome plated shutter lever to the right side of the view screen - lift steadily and completely back. As you pull the lever back, you will hear the click of the microswitch with sets the auto

98 ELECTRON MICROSCOPY: A HANDBOOK OF TECHNIQUES FOR THE BIOLOGIST

exposure system electronics into motion. At the sound of the microswitch, begin to time in seconds until the shutter swings into position to block the beam (observable through the lower view port) and the red LED comes on (above the photocell sensitivity knob). The ideal exposure time at sensitivity #5 is between 2-3 seconds! If this time is not achieved, adjust the intensity with the Brightness knob and try again until you get two identical, consecutive results. Obviously, this test should be done without film under the screen. 9. You are now ready to perform an actual film exposure. With the screen flat, press the black FEED button on the right main panel. The motorized plate drive will feed one piece of film from the load box and move it under the main view screen. The illuminated status box #0 goes dark and box #1 lights to indicate the new position of the film. At this point, pull back the shutter lever and time the exposure (2-3 seconds). Wait for the closure of the shutter and the red LED activation. After the exposure, lower the view screen flat by pushing the shutter lever fully forward. The FEED button is depressed again and the exposed film plate is advanced into the open receiver box (the #1 light goes dark and #0 is once again illuminated). Exposed TEM films can eventually be removed from the microscope by closing the camera airlock - this action also closes the light-tight receiver box internally. The films can be processed as described earlier in this chapter. ✥ SEM - Hitachi S-2400 Since all photographic exposures in SEM come off of an external, high resolution (2,000 line) CRT, there is no need to desiccate films. In order to use the Polaroid Type 55 P/N film, a 4x5" Polaroid camera back is required. It is locked into the camera box of the SEM photo-CRT with two sliding clips. The silver lever on the Polaroid back is swung into the “L” (load) position and the film is inserted carefully into the slot. It is pushed all the way to the back until a positive “click” is heard. The film is pulled out until it stops. The film is then exposed as described later. After exposure, the film is pushed all the way back in and the silver lever swung to the “R” (release) position (which is where it should be normally left when not in use). The film is pulled all the way out of the camera back and allowed to automatically process for the time recommended (20-25 sec. at room temperature for type 55 film). The film is pulled apart at the tabs and the positive print removed and coated as necessary. Negatives are detached at the perforation and cleared in fixer or sodium sulfite solution, washed and air dried. Rollers of the Polaroid back should be regularly checked and cleaned of processing chemicals, especially if streaks appear on the prints/negatives - cleaning is done using water on a non-linting cloth. If Kodak sheet film, 4127, is used, it must be loaded into the light-tight, dual cassette film holders. These two-sided holders use a sliding blind to expose a loaded sheet of film in the camera box of the photo-CRT. The cassettes are loaded under red safelight conditions making sure that the films (emulsion side up) are slid into the lower film channel and not the upper blind channel. The blind must be properly slid into the slot in the hinged

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endpiece and the metal lock put in place. When ready for an exposure, the loaded cassette is placed into the camera box of the photo-CRT and locked into place with the Polaroid back side clips. The lower blind is pulled out to expose the film in the box. The automatic exposure for the SEM is conducted and the lower blind is slid fully back into place and locked. The cassette is then flipped over (if you forget to do this, a double exposure will result) and another exposure is prepared. The exposed films can then be carried into the darkroom for processing. Dry negatives are then evaluated (for focus, quality) and enlargement printed. The S-2400 has a video output jack (RS-170) which allow for the connection of external monitors, video-recorders and thermal video printers. The addition of a frame grabber and computer allows for digital image capture, archiving and analysis (using programs such as NIH Image - a public domain program available on the internet) as well. A thermal video print (12¢) is an excellent way to determine preliminary image quality, focus, astigmatism, etc. before committing to the more expensive films ($1.00-2.00). For the best quality video prints you can either take them using the slowest viewing scan pass (setting of 3-4) or perform a photo pass (without film loaded in the camera box), allowing the pass to complete and the image to “freeze” on the viewing CRT. The frozen photo image is of the highest quality for these video prints. It should be understood that the video printer is capturing images at the lower resolution of the viewing CRT (625 lines) and not the photoCRT (2,000 lines). Image capture using a VCR should also be performed under conditions of slowest or photo scan rates for the highest quality images. ✥ S-2400 Camera System and Film Exposure As for the modern TEM, a modern SEM possesses an automated camera and exposure system. The SEM differs in that exposures are taken off of a high resolution CRT at atmospheric pressure. There is no need to desiccate films prior to their usage in the SEM. There are also a number of conditions which must be satisfied for high quality image production with the SEM. The following is a step by step method for producing these quality images: 1. Scan stub at low magnification in order to locate tissue blocks, then specific areas of interest. Once an area of interest, such as a kidney Glomerulus, is located, a final magnification must be decided upon (for example, 5,000X). 2. If possible, raise the magnification by a factor of 10X higher than the desired magnification (in the above example, raise the mag to approximately 50,000X). At this point, align the final aperture using the focus wobbler switch “APER ALIGN” in the subpanel - ideally you should observe a central pulsing of the image indicative of proper alignment (typically use TV scan rate A). 3. Stigmate (STIGMATOR X,Y) the image using a high contrast image (CONTRAST/ BRIGHTNESS controls) and a REDUCED AREA/RAPID scan rate. The best images to stigmate on are spherical in nature. Reduce or eliminate image stretching. The

100 ELECTRON MICROSCOPY: A HANDBOOK OF TECHNIQUES FOR THE BIOLOGIST

STIGMATOR MONITOR control in the subpanel can also aid in image astigmatism correction. 4. Once the image is stigmated, the magnification is reduced to the original desired value (5,000X) and the image is framed on the viewing CRT (its entire area is the frame box). 5. Using REDUCED AREA/RAPID scan, the image is critically focused at the chosen mag using the MANUAL FOCUS controls. 6. The final image BRIGHTNESS & CONTRAST is adjusted using either AUTO (ABC) or MANUAL features. If ABC is used, select the auto mode (LED will light) and depress the ABC button. ABC will appear on the viewing CRT and the SEM will make an audible sound when the auto function is completed. This can be performed at any scan rate, however, I prefer a slow scan of 3 or 4. In addition, the first attempt is rarely optimal. Always allow three to four attempts of pushing the ABC button for best results (two identical brightness/contrast results are desired). Although ABC saves time in the print darkroom since all of your negatives will be of identical exposure characteristics, there are times when it just will not yield quality images. This is especially true when a single object, such as a Paramecium, stands out against the stub background. ABC will simply average the areas of high and low contrast and brightness and a muddy, washed-out image will result. In order to emphasize the dark background and the raised, bright object, a MANUAL brightness and contrast adjustment must be performed. Initially, select the LINE ANALYSIS button (press ONCE) and use the manual brightness and contrast controls to keep the moving waveform within the bounds of the central bright, band-like area which exists horizontally across the viewing CRT. Now select scan rate #2 and adjust for the highest quality image in conjunction with the line analysis data (moving waveform should be predominantly within this central brighter area for optimum photographic brightness and contrast). 7. Once brightness and contrast is adjusted, select slow scan #3 or #4 and view the final image at high resolution - a video print is ideal at this point to critique your image before using the film. 8. In order to expose a piece of film (either Kodak sheet film or Polaroids), it must be loaded/slid into the light-tight camera box (which faces the high resolution photo-CRT) and locked into place with the side clips. The film is exposed, face down in the camera box as described earlier. 9. The exposure speed is checked (usually set on #2 - 80 sec. scan pass) and the photo START button is depressed. A scan line will slowly move down the viewing CRT as it does on the photo CRT - “painting” the image one line at a time on the piece of exposed film. At the end of the photo run, the SEM will make an audible sound and the final image will digitally FREEZE on the viewing CRT (the image will remain even if the filament current

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is turned off). To eliminate the frozen image, select ANY other scan rate (A/B, 1/2, etc.) Kodak films can now be transported to the darkroom for processing. Polaroids can be auto processed and the final print received in approximately 20-30 seconds. Type 55 P/N prints will need to be coated, while the negative will need to be cleared in a fixer or sodium sulfite bath as described earlier. Polaroid Type 53 films are coaterless and do not yield a negative. Its 400 speed (ASA/ISO) will also require that you stop down the lens in the photo-CRT camera box. The other SEM films discussed thus far have a speed of 50.

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Chapter 11 - Enlargement Printing Enlargement printing is the final step in the production of an electron micrograph. The enlarger is used to expose a sheet of photographic paper to light which passes through a negative. Areas on the negative with high grain density (i.e. dark/black areas) will interfere with the passage of light to the photographic paper. This corresponding area on the photographic paper will receive little or no exposure and therefore appear white after processing. Conversely, areas of low grain density on the negative will allow the passage of light to the paper thus exposing it. Subsequent to processing, these areas will appear dark/ black. The result is a positive print or electron photomicrograph. Since the enlarger lens can be positioned at various heights above the photographic paper (usually held in an easel), a large range of magnifications/enlargements (or even reductions) of the original negative are possible. Enlargers used for the printing of EM negatives must be able to accommodate large format negatives up to 4" X 5". In terms of enlarger design, the main unit possesses a 110v lamp housing followed by a condenser lens assembly. A variable condenser assembly is common which must be adjusted depending on the size dimensions of the negative. Next, there must be some provision for the insertion of the negative holder which contains the negative, below the condenser. Appropriate size negative holders are required which typically sandwich the negative between two metal plates. The negative should be loaded with the emulsion/dull side down (shiny side up). If in doubt as to the dull vs. the shiny side of the negative, use the notching guide. Below the negative you will find the enlarger lens assembly with a provision for removing and changing it. The focal length of the lens should be printed on it somewhere, along with the clickstop apertures or f-stops. Rotation of the f-stop ring will allow you to select different apertures. In deciding which lens focal length to use, measure the negative on the diagonal in millimeters. The enlarger lens focal length should be equal to or slightly larger than the negative diagonal measurement in millimeters. For 35mm negatives, a 50mm lens is used. For both TEM and SEM negatives, a 135mm focal length lens is a good choice. In the main unit of the enlarger, you might also find a site for the insertion of filters (such as polycontrast filters). If this is not available on the enlarger, a filter kit and filter holder which attaches to the enlarger lens is available for purchase at photo suppliers. A solid support platform of wood or metal and a rack and pinion gear system (motor or hand driven) to move the main unit closer to or away from the support base/platform completes the basic enlarger design. Additional useful equipment would include an easel (adjustable to a variety of paper sizes), a grain magnifier for fine focussing, and an automatic timer for convenience in timing exposures. ✥ Photographic Paper Grades Photographic paper is available in a variety of different surfaces/textures and contrast grades. For scientific work, the glossy surface (designated “F”) is the standard. Paper grades, usually numbered 1 through 5, refer to the contrast level of the paper. Number 1 paper is soft or low contrast and number 5 is hard or high contrast. The ideal negative can

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be described as possessing black blacks, white whites and a range of grays or halftones. This ideal negative should be printed on #3 paper. If the negative is high in contrast, it should be printed on low contrast, #2 or #1 paper to compensate. Similarly, a low contrast, muddy gray negative should be printed on a high contrast, #4 or #5 paper. These different paper grades can be purchased separately or a single paper, known as Polycontrast, with a range of paper grades is available. Polycontrast paper is designed with two emulsions; a high contrast purple sensitive emulsion and a low contrast yellow sensitive emulsion. These emulsions are selected for by the use of various purple and yellow shaded polycontrast filters. The advantage is that only one type of paper is purchased and the paper grade ranges are finer, with half-steps available (e.g. 4.5, 3.5, etc.). ✥ Process of Enlargement Printing Firstly, the darkroom must be setup. Trays of Dektol (1:2), indicator stop bath, and rapid fixer (1:1) are prepared and OA/OC safelight(s) activated. All paper grades or Polycontrast paper should be available in a light-tight paper safe. Wash water can be flowed into the print washer (which ideally is located near the fixing bath for ease of transfer. The easel should be set for the appropriate size paper (i.e. 8" X 10"). The variable condenser should be set to the negative dimensions and the proper focal length lens inserted and adjusted to the largest aperture (smallest f-stop). The negative is carefully loaded into the negative holder emulsion side down. Dust and fine hairs can be removed using a camel hair blower brush. Be careful not to scratch the negative or transfer fingerprints to its surface. The negative holder is inserted into the enlarger unit and the light for focusing is turned on (if a timer is used, it will have some provision for energizing the light for focusing and then turning it off for automatic timed exposures). With the room lights off, the negative is focused on a sheet of white paper in the easel using the enlarger focusing knob. In order to fill the frame of the easel, it may be necessary to raise or lower the enlarger lens. You may want a high magnification print of a selected area, in which case, you will have to raise the enlarger and frame the area of interest by moving the easel. The level of magnification can be determined later either by measuring the same objects on the negative and print in millimeters and by dividing the print object length by the negative object length or by placing a clear metric ruler in the negative carrier and comparing it to a metric ruler on the easel. The image must be framed and focused (preferably using the grain magnifier) with the aperture wide open to ensure the maximum available light. After framing and focusing, the lens is stopped down since the quality area of the lens is not at the periphery. The enlarger light is turned off. At this point, a test print must be made to determine proper exposure time and contrast level. One method is to expose one-sixth of a sheet of photographic paper using a rectangular cardboard cover. The enlarger light is turned on and the paper exposed for 10 seconds. The enlarger light is turned off and the second onesixth of the paper is uncovered by sliding back the cardboard mask. This area is exposed for 10 seconds, and so on, until the entire piece of paper is exposed for a final 10 seconds. Of course, the first exposed strip will have been cumulatively exposed for 60 sec., the next,

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50 sec., and so forth. The other method is to use a Kodak Projection Print Scale which is a pie wheel with wedges of increasing density. Each wedge is numbered in seconds of exposure time. To use it, the Print Scale is placed on top of the test piece of photographic paper and a 60 second exposure is done. The test print is processed, as discussed earlier in the chapter, and each wedge judged for the optimal exposure time which can be read directly off the wedge. It should be noted that initial test prints should be done using #3 paper grade unless you have experience judging negative contrast levels. The test print also serves to determine whether the contrast level of the print is adequate or if a change of paper grade is in order. Changes of paper grade require a modification in exposure time which will be covered later. If the contrast of the test print is satisfactory, the correct exposure time must be determined. Exposure times between 15-25 seconds are ideal. Long exposure times may introduce vibration effects to your focus while short exposures lack control over consistent duplications. If necessary and if possible, the f-stop should be adjusted to allow for a final exposure time of 15-25 seconds, based on the results of the test print. Once the proper exposure time is determined, the timer is set and a fresh sheet of paper is loaded into the easel (be careful that you have not moved the position of the easel). The paper is exposed for the correct time followed by processing, washing and drying. ✥ Enlargement Printing Variables Three common variables in enlargement printing are increases/decreases in magnification, contrast/paper grade and lens aperture diameter (f-stop). When exposure time has been determined for a certain enlargement and the magnification is then increased, the exposure time will also have to be increased since there is a greater spread of the light. Less available light means having to increase exposure time for a print comparable in quality to the lower magnification print (relative to grain density and contrast). Reduction in magnification requires a reduction in exposure time. The following calculation provides an approximate change in exposure time when changing magnification:

T2 = T 1

(M2 + 1)2 –––––––––– (M1 + 1)2

where: T2 = the new exposure time T1 = the old exposure time M2 = the new magnification M1 = the old magnification Relative to changes in paper grade, an increase in paper grade, say from #3 to #4, will require an increase in exposure time. A decrease in paper grade will require a decrease in

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exposure time. A calculation for approximating such changes in exposure time relative to changes in paper grade can be made since each paper grade has a reference printing index. The print index for each paper grade is listed below: PAPER GRADE PRINT INDEX 1 5000 2 3200 3 2000 4 1250 5 1000 The calculation based on these print indices follows: I2 T2 = T1 ––––– I1 where: T2 = the new exposure time T1 = the old exposure time I2 = the new paper grade print index I1 = the old paper grade print index In terms of a change in lens aperture or f-stop, one must firstly understand that the higher the f-number, the smaller the diameter of the lens opening or aperture. Therefore, the f8 opening is larger than the next stop, f11. To be specific, the next stop up is one-half the diameter of the stop below it and will admit only one-half as much light. By example, an exposure of 60 sec. at f11 is equivalent to 30 sec. at f8 and a 15 sec. exposure at f5.6. A test print indicating 60 sec. at f11 could be brought into ideal exposure time by opening the lens to f5.6 and exposing for 15 seconds. Conversely, a test print that indicates 5 sec at f5.6 could be stopped down to f11 and exposed for an equivalent 20 seconds. ✥ Printing Tricks If attention to detail is given throughout all stages of EM sample preparation, microscope operation and photography, resorting to darkroom “tricks” will not be necessary. The following are intended to yield a more uniform and professional quality print. When areas of a negative are extremely dark, the result on the print will be a stark white area, such as the unexposed paper appears. To provide some background grain density, a process known as intensification can be used. While in the developer solution, the surface of the white areas are rubbed with a couple of fingers. Body heat causes the activation and development of some crystals in the area. Burning-In involves negative regions which are again too dense/dark which produces

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washed out print areas. These areas can be burned-in by exposing the entire print for the ideal exposure time (as determined by test print) and then adding some additional exposure time to the excessively white print region while masking the rest of the print surface. You can purchase cardboard masks for this purpose or you can simply use your hands at differing distances from the lens to form a suitable mask. While the additional exposure time elapses, it is imperative that you keep the mask or your hands moving in order to prevent a darker outline of the burned-in area. Dodging-Out is similar to burning-in except that some regions on the processed print are excessively dense/dark due to a low grain density negative area. In this case, the ideal print exposure time is determined by test print along with the area of concern. During the course of the ideal exposure time, this low density negative region is masked at few second intervals throughout the exposure. Do not forget to keep that mask moving using shaky lateral motions.