Preparatory techniques, including cryotechnology

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PREPARATORY TECHNIQUES, INCLUDING CRYOTECHNOLOGY ANN LEFURGEY† JOHN D. SHELBURNE‡ PETER INGRAM*

ANN LEFU RGE Y ET AL.

PR EPA RATORY TEC HNIQUES

†Veterans Affairs Medical Center and Department of Cell Biology/‡Pathology

Duke University Medical Center Durham, North Carolina *Research Triangle Institute and Department of Pathology Duke University Medical Center Research Triangle Park, North Carolina

INTRODUCTION Analytical electron microscopy (AEM) instrumentation now offers the ability to correlate microscopic anatomy of human tissues with elemental chemical composition. This ability is a signi~cant improvement over older bulk methods of chemical analysis that necessarily disrupt histomorphology. The quality and validity of the morphochemical correlation obtained by AEM are, however, highly dependent on the appropriate selection of specimen preparative techniques for the particular problem under consideration. The route employed to bring any biological tissue from its in vivo state to its in situ form in the microscope is potentially fraught with numerous pitfalls affecting both structural and chemical integrity. Often, a paradoxical trade-off exists between the two. Well-established EM preparative techniques that achieve excellent structural preservation (e.g., chemical ~xation, osmi~cation, staining) often sacri~ce chemical integrity. However, techniques suited for chemical preservation (e.g., freeze-~xation) may disrupt subcellular architecture or yield poor ultrastructural images. The microscopy literature has expanded in the past decade with much effort to delineate and resolve the dif~culties associated with various preparative techniques of biological specimens. Several excellent reviews of this work have discussed the evolving available methodologies, their advantages and disadvantages. It is beyond the scope of this chapter to review this complex and vast subject in detail. Instead, the major focus will be to review several concepts that are especially relevant in biomedical applications and to outline techniques particuBiomedical Applications of Microprobe Analysis Copyright © 1999 by Academic Press. All rights of reproduction in any form reserved.

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larly useful in general surgical pathology. Subsequent chapters in this book provide additional detailed information on specimen preparation. As a guide to the literature, the reader is referred to several books with excellent discussions of preparatory techniques for X-ray microanalysis in research,1–14 general discussions of ~xation,10,11,15–19 medically oriented reviews,20–30 and references with discussions of artifacts.10,11,31–40 In general, no one technique has assumed universal acceptance for all studies in biology and medicine. Instead, the preparative methodology must be thoughtfully tailored to the speci~c problem under study. As discussed by Echlin,10,41 Coleman and Terepka42 have outlined excellent criteria for optimum specimen preparation for biological X-ray microanalysis. These are: (1) the normal structural relationships of the specimen should be adequately preserved so that it is possible to accurately identify the analyzed structures; (2) the amount and (3) the chemical identity of material lost from, or gained by, the sample must be known and should be minimized; (4) the amount of elemental redistribution and translocation within the sample must be known and should be minimized; and (5) if chemicals must be used, they should not mask the elements being analyzed. In order to achieve these goals, it is usually necessary to prepare the sample by freezing. Only in this manner can the numerous artifacts well known to occur with aqueous chemical ~xation be avoided. It is important to remember that the act of freezing a sample is itself a form of ~xation. After all, the term ~xation simply means to hold the tissue as it was at a moment in time in vivo. With aqueous chemical ~xation, while the protein shell of a tissue may be well preserved, electrolytes are translocated because of activity gradients set up by the introduction of the ~xative. Thus only by freezing is there preservation of all elements and molecules in the sample. Nevertheless, it is clear that freezing techniques are tedious and dif~cult. Numerous artifacts are caused by improper freezing, by cryoultramicrotomy, and by freeze-drying. For many problems in medicine, freezing technology is unnecessary. For example, in the study of asbestos ~bers in lung tissue, results can be obtained readily from formaldehyde-~xed tissues with probably no signi~cant alteration in the composition of the ~bers. In general, one should employ the easiest preparative method that will suf~ce to answer the questions being posed. In problems of human pathology, much of the microanalytical work to date has relied on relatively unsophisticated techniques of specimen preparation. These methods require only minor modi~cations from routine histologic processing for light or electron microscopy. Because most clinical cases in which AEM has been used have involved foreign material identi~cation and localization, usually nondiffusible, inorganic xenobiotics, these simple techniques have usually been adequate. Again, simple preparative techniques are generally superior to more elaborate ones if they are suf~cient to resolve the problem under study. Undoubtedly, greater numbers of pathologists in the future will address more dif~cult questions in human pathophysiology, using microanalytical techniques to describe the in vivo distribution of soluble endogenous elements and molecules. Such work necessarily calls for signi~cant technical improvements in and wider adoption of more elaborate methods of tissue preparation, such as cryotechniques.

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This chapter outlines the major preparative techniques employed to date for qualitative energy dispersive X-ray (EDX) analysis in human pathology and enumerates some common artifacts and contaminants that can hamper proper data interpretation. Additional details are provided in subsequent chapters on speci~c organs and problems.

CHEMICAL FIXATION Artifacts Aqueous chemical ~xatives such as buffered formaldehyde and glutaraldehyde are commonly used in medical centers for ~xing biopsy and autopsy tissues. For many of the medical questions raised in this book, aqueous chemical ~xation is not only adequate, it is preferred, as it is generally much easier than freeze ~xation. Also, chemically ~xed tissues pose little or no infectious hazard to workers. However, wet chemical preparative techniques are notorious for causing losses, redistributions, and additions of diffusible substances. All wet processing steps, particularly ~xation, have been well demonstrated to cause signi~cant leaching of endogenous diffusible elements. Organic compounds may also be lost during dehydration and embedding steps. Diffusible elements may then redistribute themselves down concentration gradients, become bound preferentially to other tissue structures, and may thereby eventually appear concentrated in artifactual locations. The formation of osmium–calcium precipitates in conventionally EM processed tissue is an example. Moreover, phase transformations of endogenous substances can occur during wet techniques; thus amorphous calcium phosphate may become crystalline hydroxyapatite during conventional processing.3 Finally, exposure of the tissue to a variety of histologic solutions necessarily causes the introduction of extraneous elements and contaminants into the specimens. Mercurial ~xatives such as Zenker’s formalin add mercury and chromium, cacodylate-buffered glutaraldehyde introduces arsenic and sodium, and osmium tetroxide post~xation, of course, deposits osmium. Moreover, numerous common stains contain a host of heavy elements as normal constituents. Examples include iron in Prussian blue, aluminum in hematoxylin, bromine in eosin, and copper in luxol fast blue and in Alcian blue, as well as silver, lead, and uranium staining. For example, a low concentration of aluminum (Z ⫽ 13, Ka ⫽ 1.49 KeV) cannot be detected reliably in a conventional paraf~n section stained with hematoxylin and eosin not only because hematoxylin may add aluminum, but also because the eosin may add bromine (Z ⫽ 33, Ka,b ⫽ 1.48 KeV). Contaminants from the histology laboratory may also occur from poorly washed glassware, exposure to dust particles, and the use of tap water. Urban tap water often has readily detectable aluminum as well as numerous other elements. We therefore recommend the use of distilled and deionized water for the preparation of all appropriate chemicals and ~xatives and all cleaning of glassware. During the course of our work, for example, we have found lead as an occasional contaminant in stained paraf~n sections from biopsies obtained from outside hospitals. The addition of such exogenous elements to the specimen can severely

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compromise the AEM study, both in terms of imaging and X-ray data interpretation. Table 1 of Chapter 1 reviews several common peak overlap problems. Histologic stains can also remove elements; we have found that calcium and phosphorous are leached from undecalci~ed bone sections stained for aluminum.43,44

Conventional Histologic Sections Despite these many potential artifacts, probably the most common type of medical microprobe analysis involves the use of routine histologic sections. Formalin-~xed, paraf~n-embedded tissue sections have been shown to be particularly helpful in cases where high-resolution ultrastructural elemental localization is deemed unnecessary, thereby avoiding the need for more dif~cult EM tissue processing.25,45 In particular, paraf~n-embedded tissue has been demonstrated to be suf~cient for cases involving inorganic particulates or insoluble, tightly bound exogenous metals. The sections are cut at 5–10 lm and are picked up onto spectroscopically pure, ultrasmooth carbon planchets (Fig. 1). Paraf~n tissue sections adhere readily to the carbon substrate after heating at 60⬚C for 30 min, followed by

FIGURE 1. (A,B) Two methods of mounting conventional histologic sections on a carbon support. Because plastic coverslips are insulators, carbon paste should be as close as possible to the section in order to minimize charging.

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deparaf~nation in xylene and air drying. The sections are subsequently rotary carbon coated by vapor deposition to minimize electron beam charging; for sections 5 lm thick or less, carbon coating has been shown to be usually super_uous and may be omitted.22 Also, the thinner the section, the less likely buried particulates will be missed with backscattered electron imaging (BEI). This preparative method permits the microscopist to use routinely ~xed, paraf~n-embedded tissues for investigating unknown particles discovered by light microscopy (LM). If the particles appear by LM as numerous and relatively homogenously distributed in the section, one may simply apply this technique to a serial unstained section cut from the paraf~n block. In some instances, the paraf~n block may not be available or the particles in question are quite small and few, requiring LM localization prior to scanning electron microscope (SEM) study. In these cases, tissue section transfer methods have been used successfully to lift the section off the glass slide onto a carbon planchet.46,47 If photomicrographs are taken of particles of interest prior to the section transfer, speci~c (inorganic) inclusions can be localized and probed with precision to determine their composition.47 The major drawback of this technique is that sometimes the crystal of interest may be lost during the transfer. In some rare instances, one may discover the presence of foreign material during the analytical SEM study that had not been appreciated by the prior LM examination. In these cases one would ideally like to reexamine the section by LM to locate and de~ne the histological correlate of the microanalytical ~nding. In rarer cases yet, one may wish to undertake a more sophisticated mode of microanalysis of the pathologic tissue, such as ion microscopy (IM). In both of these situations, the following technique of specimen preparation is extremely useful, as it enables correlative light, electron, and ion microscopy of a single histologic section.48,49 In brief, sections are cut onto transparent chlorine-free polyester coverslips (Thermanox, Miles Scienti~c, Napierville, IL). These coverslips permit excellent LM, as well as analytical electron and ion microscopy. Light microscopic examination of the specimen is possible both before and after microanalytical studies (Figs. 1 and 2). As this technique is somewhat more tedious and time-consuming than those using serial unstained or lifted stained sections, it should be reserved for cases where precise correlative microscopy is needed and the other techniques do not suf~ce. Unfortunately, polarizing light microscopy is dif~cult or impossible with plastic coverslips, as they are anisotropic. For polarizing particles the tissue section transfer method described earlier is better. An overview of these procedures is given in Fig. 3 as a _ow diagram. The various possible strategies are indicated with arrows. Figures 4 and 5 and Fig. 14 of Chapter 1 illustrate these different approaches. Figure 14 of Chapter 1 illustrates a case of talc granulomata. In this case, talc particles were numerous and relatively homogeneous by light microscopy. In order to probe these particles, it was only necessary to cut serial sections onto carbon supports. Talc particles were then imaged easily and quickly with the backscattered electron detector and probed for elemental composition. In Fig. 4, a pathologist had identi~ed polarizing particles in giant cells in a patient with pulmonary sarcoidosis. In order to probe these particular crystals and avoid the numerous aluminum silicates, ubiquitous in human lung, it was necessary to photograph the cell in question

A

B

FIGURE 2. Schematic diagram of the recommended technique for optimal high- and low-resolution microscopy of histologic sections on plastic coverslips. (A) For high-resolution light microscopy (e.g., using a 40 or 100⫻ objective lens), the plastic-mounted histologic section is sandwiched between a glass coverslip and a glass slide with oil on both sides of the plastic coverslip. It is important to note that the tissue should be facing up as shown. (B) For low-resolution light microscopy (e.g., using a 20⫻ or lower power objective lens), it is only necessary to place the plastic-mounted section on a glass slide, section side up. No oil or coverslip is needed, thus potentially minimizing artifacts.

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FIGURE 3. Techniques for analyzing conventional histologic sections are reviewed in this _ow diagram. Details are given in the text.

and then lift this section from the glass slide and place it onto a carbon support. Using the previously recorded light photomicrographs as a guide, it was then a simple matter to identify and probe the crystals in question. Figure 5 illustrates the most elegant but most tedious approach. These plastic coverslips permit correlative light, electron, and ion microscopy. Once a conventional histologic section is examined on a carbon support, it is dif~cult or impossible to remove it

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A

B

C

FIGURE 4. (A) Light photomicrograph. Pulmonary sarcoidosis. A giant cell (arrow) is examined with partially crossed polars in order to enhance the cytoplasmic crystals. (Conventional paraf~n section stained with H&E, ⫻ 220) (B) The identical section was lifted from the glass slide using a plastic spray (28) and is examined here with backscattered electron imaging. The same giant cell is indicated with an arrow (⫻ 258). (C) A higher magni~cation view of the same giant cell examined with backscattered electron imaging (⫻ 1065).

intact. Thus future bright-~eld transmitted light microscopy studies on the same section are not possible. Plastic coverslips solve this problem.

Epon Sections Numerous studies have used sections of Epon-embedded tissues that have been prepared by conventional or modi~ed EM processing,21,50–71 (Chapter 12). Depending on the tissue and the level of ultrastructural detail desired, osmi~cation

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FIGURE 5. Same region of a single histologic section imaged by light microscopy (A), pre-ion microscopy backscattered electron imaging (B, tilt ⫽ 30⬚), ion microscopy imaging of barium (C), and post-ion microscopy backscattered electron imaging (D, tilt ⫽ 0⬚). This hematoxylin and eosin-stained paraf~n section is of an abscess that contains numerous crystals of barium sulfate. Arrows indicate identical particles of BaSO4.

and staining with lead or uranium may be omitted, as each of these steps introduces metals into the tissue that can interfere with both imaging and analysis (Table 1 of Chapter 1). EDX can be performed on thin sections mounted on grids of copper, nickel, titanium, steel, beryllium, carbon, or nylon, followed by carbon coating to minimize charging. The specimens are then analyzed by a conventional TEM or STEM equipped with an EDX detector. There are several disadvantages of this approach. These include the small sample size (compared to paraf~n sections), the usual presence of heavy metal stains, and confounding peaks from the grids except when beryllium or carbon is used. Unfortunately, there is often a presumption that routine electron microscopy preparative procedures are somehow “better” than paraf~n processing for microprobe analysis. In fact, for most microprobe studies in surgical pathology, the reverse is true.

Undecalcified Bone Microprobe studies on mineralized bone necessarily employ the techniques well described in several references.72–74 In brief, the bone is ~xed in alcohol and embedded in a resin such as methacrylate. The point is that the bone tissue is not decalci~ed as is conventional practice. This type of sample is ideal for microprobe studies on aluminum deposition, as is noted in Chapter 3.

Digestion Techniques Digestion preparative techniques for AEM of inorganic particulates in tissue are reviewed in Chapters 7. These techniques involve an alkaline digestion step

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to remove the organic matrix, followed by a concentration step using ~ltration. The cellulose acetate ~lter containing the particulates is then transferred to a formvar- and carbon-coated TEM grid and dissolved slowly by acetone vapor, resulting in the deposition of the inorganic residue directly onto the grid. Both selected area electron diffraction (SAED) and EDX can then be performed on the particulates to yield complementary data for accurate identi~cation. Replication techniques75 are an alternative to digestion and afford better correlation with the histology. The ~lter can also be mounted on a carbon disk coated with a suitable conducting ~lm and examined by SEM. While digestion techniques are usually employed on formalin-~xed tissues, this is merely for convenience. Freeze-~xed tissues can of course also be treated in a like manner. The disadvantages of digestion techniques are obvious to morphologists. However, the major advantage is that particle burdens can be quantitated accurately. Instead of searching tediously for dozens of particles with BEI, the investigator readily images thousands of particles by SEI and/or BEI and all are on the “surface.” As discussed in Chapter 1 (Fig. 13) and Chapter 7, automatic imaging and counting systems exist to facilitate this type of analysis.

CRYOFIXATION Although freeze ~xation and cryoultramicrotomy are, in principle, ideal methods for the ultrastructural microanalysis of diffusible substances, they currently pose substantial technical dif~culties, especially for ultrastructural work. Ice crystal damage, thawing, and ice recrystallization artifacts present real hazards to freeze ~xation, cryosectioning, freeze drying, and section transfer. Such phenomena can result in the gross disruption of morphology and the serious translocations of endogenous diffusible elements. Likewise, contamination during tissue transfer steps, freeze drying, and carbon coating is always a potential threat to chemical integrity. Moreover, the techniques used for ultrastructural preservation are laborious and require a high degree of technical expertise and sophisticated equipment. Gilkey and Staehelin76 and others77–82 have reviewed cryotechniques in detail. Some of the most successful ultrastructural cryostudies have been conducted by Sommer et al.,83 as described in detail in Chapter 9. The technique of freeze substitution, i.e., the replacement of frozen water with an organic solvent at low temperature and subsequent resin embedment, has been revitalized.84–88 Although this has the disadvantage that there is invariably some loss or redistribution of localized elements due to the procedure, it does allow for the production of thin sections for the easier identi~cation of conventional morphology. Similarly, cryoprotectants are used occasionally to improve the depth of tissue preservation free of ice crystal damage during freezing.8,81,82 Again, however, there will inevitably be some translocation of physiologically labile elements, as well as potentially toxic effects from some cryoprotectants. Such procedures should be used with caution in conjunction with quantitative microprobe analysis. At the light microscopic or cellular level of resolution, however, the techniques used for conventional cryostat microtomy are comparatively simple and may suf~ce to

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answer many important questions in physiology and pathology.89–95 Chapters 9, 11, 13, and 15 provide many examples. In general (with the exception of high-pressure freezing96), the faster the rate of freezing, the smaller the size of ice crystals formed in the tissue. Ideally, freeze ~xation is so rapid that no ice crystals form, i.e., vitri~cation. In current practice, vitri~cation of water may only be achieved with ideal samples in a research setting,97 and even then there is controversy. With biopsies, the general goal is to keep ice crystal size smaller than the probe size. In this manner, translocations caused by ice crystal damage become relatively unimportant. The easiest freezing techniques involve the rapid plunging of small samples into liquid nitrogen-cooled cryogens such as liquid propane, various freons, or ethane.98–100 We prefer ethane as it is less explosive than propane and provides greater ease in cryosectioning. Direct immersion into liquid nitrogen itself is not recommended. Nitrogen gas bubbles insulate the tissue, producing a fairly slow rate of cooling, hence larger ice crystals. Surgical pathologists commonly drip liquid cryogens (e.g., freons) directly onto the tissue. While adequate for diagnostic work, this technique is not recommended for cryo~xation as ice crystal damage is usually severe. Also, electrolytes in the mounting medium may interfere with a microprobe study. Newer technologies for cryostat freezing and section collection show promise for electron probe cases, as the freezing process is a rapid method involving controlled dropping of a cooled metal mirror heat extractor onto the specimen.101,102 These methods have been shown to produce cryostat sections with vastly improved structural preservation and of a quality suitable for immunolocalization of c-myc protein103–105 and bone morphogenetic proteins.106 Although the methodology is complex, some of the best results to date have been obtained by “slamming” tissues against a liquid helium-cooled, highly polished copper surface.83,107–109 It has been argued that the vitri~cation of tissue water can be achieved with this approach.97 Outstanding ultrastructural and X-ray microanalytical results have also been achieved with high-pressure freezing devices.80,110 Vitri~cation has also been demonstrated to occur with this method.36,96,111,112 Other methods and devices that are portable and relatively low in cost may be adapted more readily for use in the surgical or autopsy suite, although the freezing obtained may not always achieve vitri~cation quality as with metal mirror helium or high-pressure cryo~xation. Of special note is the cryogun device developed by F. Lightfoot;113,114 this portable metal mirror instrument is liquid nitrogen cooled and can achieve excellent preservation to depths of 15–30 lm.115 Other cryo~xation techniques useful in a wide variety of clinical and/or research settings include cryopunching,117 cryojet freezing,118–120 cryoplunging,121–131 clamp freezing,132–136 and a cryoneedle technique.137–139 In all cases, the quality of freezing should be monitored independently with parallel aliquots of tissue processed by freeze-substitution and/or freeze-fracture analysis.83,99,140 After freeze ~xation, the tissue can be either microtomed or examined whole. For ultrastructural studies, the frozen tissue should be maintained under ⫺140⬚C to prevent recrystallization. Storage of frozen tissues prior to further processing simply involves immersion under liquid nitrogen (⫺196⬚C).81 During the 1990s, important advances in instrumentation and techniques for

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cryoultramicrotomy and cryotransfer have made maintaining these low temperatures throughout all subsequent steps of microtomy and/or transfer much less dif~cult,141–151 although low-temperature cryopreparation for high-resolution elemental analysis and imaging is still challenging.10,81,82,152,153 Microtomy is accomplished with either a cryoultramicrotome for ultrastructural studies81,99,141–151 or a conventional cryostat for low-resolution (cellular) work.89,90,92,93 Sections or whole mounts are usually examined after freeze drying. During freeze drying it is important that all water is removed from the tissue with minimum disruption of the anatomy. In practice, this translates into very slow warming rates. Devices for optimum freeze drying have become available commercially at moderate cost.154 We have sometimes used the electron microscope ~lm dryer for this purpose and found it satisfactory; one simply transfers the hydrated sample to a precooled heavy copper block and places this rapidly into the ~lm dryer and leaves it to warm up to ambient temperature for about 24 hr. A conventional carbon evaporator can also be used in a like manner for freeze drying cryosections. The major dif~culty with these relatively inexpensive freeze-drying devices is silicon and sulfur contamination from pump oils. These can be minimized by suitable trapping of the pumping lines. For quantitative analyses, however, it is strongly recommended that totally dry pumped systems (such as cryopumps) be used.154,155 Cold stages can permit the examination of non-freeze-dried, i.e., frozen-hydrated, sections or bulk samples.156–170 Unfortunately, the presence of water (ice) reduces both the X-ray signal and the visualization of morphology greatly. In order to increase the X-ray signal, one can increase the beam current, but this may melt the ice, thus destroying the morphology. Despite these problems, cold stages cooled by liquid nitrogen or liquid helium can minimize (but not eliminate) mass loss and can be effective in reducing mass gain (contamination) in the area being probed. With room temperature microprobe analysis, elemental losses can be very signi~cant, especially for volatile elements such as mercury, vanadium, and sulfur. Use of low-beam currents can minimize these losses but may not provide enough X-ray signal. In a research setting, Marshall and Xu169 have demonstrated that it is possible to use the oxygen X-ray peak (Ka⫽ 0.52 keV) to quantitate the water content in frozen bulk specimens. Leapman, Andrews, and colleagues171–174 have accomplished a similar result using electron energy loss spectroscopy (EELS) at high spatial resolution on ultrathin cryosections. From a physiological standpoint, it is important to obtain independent measurements of local water content, as well as elemental content, at the subcellular level. Thus while the examination of frozen hydrated samples on a cold stage is the theoretical ideal, there are numerous technical problems that preclude this approach for routine work. In fact, even the comparatively easier freeze-dried cryostat section is still rarely employed for routine clinical work. In addition to the problems inherent in producing cryostat sections, freeze-dried cryosections are not _at due to the differential water content of various intracellular compartments. Therefore the quantitation of electrolytes (in terms of wet weight concentration) in these sections is dif~cult.3,9,77,82,171–178 Qualitative data on physiologic ion concentrations are currently of little practical clinical value. However, as quantitative EDX has become more readily accepted, freeze-~xed

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biopsies of tissues such as the kidney,175,176 the heart,179–186 the myometrium,92,93 and specimens from patients with cystic ~brosis (Chapter 11) are now being probed to study normal and abnormal electrolyte shifts.

FLUIDS Extremely accurate techniques have been described for quantitative elemental analysis of biological _uids using AEM.187–192 In all such methods, the analysis of microvolume (100 pl) samples can be performed after a series of steps designed to produce uniform evaporation and evenly distributed, small (⬍1 lm) crystals. These analytical methods have proven extremely useful for microdroplet studies in basic renal physiology, as well as in the study of other biological _uids. Relatively few studies to date, however, have analyzed normal or pathologic human _uid.193–198 A notable example of such studies are Quinton’s investigations of cystic ~brosis (CF) sweat ducts.196–198 Quinton and associates have collected sequential sweat droplets under water-saturated mineral oil from single sweat duct ostia in vivo and analyzed these droplets by electron probe microanalysis. By combining these results with electrophysiological studies both in vivo and in vitro, Quinton has well characterized the ion transport properties of CF sweat ducts. For further discussion of microprobe analysis in cystic ~brosis, see Chapter 11. This approach holds great potential for further investigations in human pathology.

WHOLE MOUNTS A variety of normal and pathologic human tissues have been analyzed by air-dried, whole mount preparations. Hair (Chapter 13), ~ngernails (Chapter 11), urinary stones (Chapter 10), and suspected environmental pollutants can be mounted directly onto spectroscopically pure carbon planchets using conductive carbon paste.199,200 Erythrocytes, platelets, and sperm cells have likewise been analyzed as whole mounts, on TEM grids or SEM mounts, requiring no adhesive material.20,23,55,194,201–204 Such simple techniques have often been shown to be adequate when compared to other techniques such as wet chemical or frozen preparations. Similarly, droplets of pharmaceutical agents containing labeled molecules can be placed on specimen supports for determination of the detectability of the label, e.g., element, in question. Such analyses have been integral components of experimental design in several studies where subsequently the labeled molecules have been detected within cryoprepared cells and tissues by X-ray microanalysis and imaging.130,131,205–208

COATING Most biological samples must be coated in order to avoid charging. We recommend carbon coating of freeze-~xed, freeze-dried cryosections on carbon-

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coated or formvar grids in order to stabilize the sections under the electron beam and to minimize possible rehydration during transfer to the microscope. We have found that the most generally useful type of coating is with carbon. Because of its low atomic number, no characteristic peak for carbon is seen with a standard beryllium window EDX detector; Bremsstrahlung radiation, however, is present as a broad continuum. We apply the carbon coating using a rotary device so that it is applied evenly to all features of the sample. Abraham22 has pointed out that even a large sample such as a 5-lm histologic section can be studied with no coating whatsoever if it is put on a carbon support dike. While we agree with this observation and have obtained the same result, we have found that unless one is very careful, histology laboratory sections are often thicker than 5 lm and thus will indeed charge at least in certain regions of the sample. This is especially true when dealing with solid tissues such as liver, as opposed to more porous tissues such as lung. In general we would advise rotary carbon coating for histologic sections so that there is no problem in this regard. Obviously, it is extremely inef~cient to put a sample into the SEM and to begin to collect data only to ~nd that charging is occurring in some regions. This necessitates removing the sample from the instrument, coating it, and putting it back in with the loss of at least an hour. In general, metal coats, such as those used routinely in SEM laboratories with standard sputter coaters that deposit gold or platinum, are to be avoided. These do have advantages over carbon in that better resolution for SEM is obtained, and they are extremely easy to operate. Nevertheless, the peaks from the metal may mask peaks of interest in the sample, and the metal coat can sometimes absorb X rays from lower atomic number elements in the sample. Also, the entire sample becomes highly backscatter positive, thus compromising this method of detection. Despite these problems, workers who are interested in asbestos ~bers will often use a thin gold coat and are still able to detect the elemental composition of the ~ber with suf~cient qualitative accuracy that the various asbestos ~ber types can be identi~ed. Thus for some studies a metal coat can be tolerated. Charging due to ice is a problem for frozen-hydrated sections or bulk samples. A carbon coat is usually necessary for analysis. Marshall158,159 has advocated the use of a beryllium coat for frozen-hydrated tissues. Because beryllium peaks are also not observed using standard EDX detectors with beryllium windows, this approach is comparable to carbon coating. Caution must be exercised, however, as beryllium vapor is highly toxic and can produce pulmonary granulomas.

SPECIMEN SUPPORT For specimen support in an SEM, most workers use standard carbon planchets, carbon disks, or stubs. These are readily available from several vendors. In our experience, these are usually clean if one speci~es spectroscopically pure material. Nevertheless, Abraham22 points out that it is prudent to check each stub prior to using it for critical work; some stubs that are allegedly “spectroscopically pure” may still have contaminating particles on the surface. Obviously

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this problem varies from vendor to vendor and possibly from lot to lot. Accordingly, caution is always indicated. If one does not have time to check the stub before putting the sample on it, an acceptable alternative is to be certain to study a region of the stub not covered by the sample prior to spending too much time on the sample itself. As noted in Chapter 1, spectral data should always be obtained from that part of the specimen support devoid of tissue, and it should also be examined microscopically as a ~nal check on its purity. For transmission electron microscopy, so-called “low background” grids made of beryllium can be obtained. As noted earlier, these have the advantage of showing no characteristic X-ray peak in the conventional EDX detector and of being metal. Many workers, however, have found that it is preferable to use nickel or other metal grids as the characteristic peak obtained from the grid can be used in quantitative studies. The peak-to-background ratio from the grid can be obtained by probing areas away from the sample. Copper is usually best avoided for biological studies, as the copper La peak (0.93 keV) line may interfere with the Ka peak (1.04 keV) for sodium. Low background grid holders made of carbon or beryllium are generally used, although, again, aluminum or some other metal can be used in order to unambiguously identify (and quantify) the background X-ray signal. However, this will cause obvious problems in certain clinical situations where the element to be identi~ed is the same material as the grid or its holder! Clinicians and pathologists are urged to avoid the study of histologic sections on glass slides. Although this is tempting, as the sections are normally placed on glass, we have found that the composition of glasses obtained in most histology laboratories can vary from vendor to vendor and even within a single slide. Also, in medical work, silicon is often of interest. Obviously, silicon in the slide can interfere. Elements of physiologic signi~cance such as calcium and sodium are also often present in glass. Thus the background obtained from a glass substrate is usually unacceptable for most types of biological studies. Furthermore, glass is an excellent insulator, and charging is a serious problem when it is the substrate. If it is necessary to have a sample on a transparent support, we recommend the use of chlorine-free plastic coverslips as noted previously. These can be studied by light microscopy, but provide no signal to the standard EDX detector. Nevertheless, these are, like glass, an excellent insulator. A carbon coating must be applied in order to prevent charging.

COMPROMISES For many types of problems it is necessary to reach a compromise between the theoretical ideal of freeze ~xation and the excellent morphologic data obtained with chemical ~xation and/or conventional embedding. Figure 6 outlines the common methods, including interactions between cryo methods and ambient temperature methods. Thus workers interested in calcium deposits have successfully employed regimens that involve freeze ~xation, freeze drying, vacuum embedding with a plastic resin, and subsequent polymerization. In these studies, care must be taken during the microtomy of plastic resins not to _oat the section onto a water bath. Not only can contaminants in the water bath be

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added to the section during this procedure, but it is also clear that electrolytes are lost from the section into the water bath.99 Thus these sections should be picked up dry, directly onto a specimen support such as an EM grid. Needless to say, it is not easy to pick up thin, dry plastic sections. We have found that most of them tend to wrinkle and to fold. Nevertheless, with practice, and with many samples, it is usually possible to ~nd a few good areas on a few grids. Ingram and Ingram209 have proposed the use of low chloride resins as a way of avoiding the chlorine peak present in standard epoxy resins. Low-temperature embedding resins (e.g., Lowicryl) are used most commonly by lowering the sample temperature following chemical ~xation and during dehydration, but some investigations have combined freeze drying with embedding in these resins.210–213 This technique eliminates all chemical ~xation and exposure to organic solvents as the samples are quick frozen, dried in vacuo, and in~ltrated directly in pure resin. Freezing and freeze drying followed by embedding in heat-polymerized resins has also been utilized extensively by Ingram and Ingram214 for X-ray microanalysis of various animal cell types. The combination of low-temperature resins and freeze drying may minimize artifactual translocations and should prove to be useful in clinical applications. However, appropriate controls and standards must be analyzed in parallel to demonstrate not only that translocation is prevented, but also that no interferences arise from elements in the embedding resins. The precipitation techniques noted in Fig. 6 have been reviewed by Morgan2; these techniques are still in use today.215 Dotted lines indicate that these are alternative processing paths, not ~rst choices. In general, ambient temperature techniques are useful as they are technically easier than cryo methods, provide better morphology, and may serve as a quality control monitor, e.g., freeze-substituted tissues can determine ice crystal damage.83,91

FIGURE 6. Summary of methods for the microprobe analysis of biological material (after Morgan7). Dashed lines indicate alternative processing paths.

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Many workers have advocated doing biological X-ray microanalysis in several different ways on the same tissue in order to maximize the information obtained from different techniques and to minimize the artifacts while minimizing the substantial dif~culties of the most elaborate techniques, such as cryoultramicrotomy. With this approach, the investigator not only accomplishes conventional light and electron microscopic studies, but also then studies the tissue using, say, compromise techniques such as freeze-~xed, freeze-dried tissues vacuum embedded in plastic. By slowly assembling data from several different approaches, it may be possible to eventually build up an accurate picture of the true in vivo situation.99 A useful general rule is that if one anticipates future need for any sort of chemical or microanalytical studies, it is always wise to freeze a portion of the tissue and to hold another portion of tissue in an appropriate buffered formalin or glutaraldehyde.216,217 By doing so, one can subsequently tailor an individualized technique of specimen preparation to the problem under consideration, especially if artifacts such as peak overlaps occur with routinely processed tissue. Even if it is not possible to snap-freeze a rare or unusual clinical specimen, if there is excess tissue not needed for ordinary diagnostic work, clinicians and pathologists are urged to store an aliquot in an ordinary freezer. Sealable clear plastic bags minimize contamination and dehydration. Roomans and Wroblewski216 have shown that even though ice crystal damage will of course be severe, this simple, inexpensive procedure will preserve electrolytes for low-resolution (i.e., cellular) analysis.

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