Cryof ixation of Vascular Endothe1ium ROGER C. WAGNER AND S. BRIAN ANDREWS School of Life and Health Sciences, University of Delaware, Newark, Delaware 19716 (R.C.W.); Laboratory of Neurobiology, National Institutes of Health, Bethesda, Maryland 20205 (S.B.A.)


Cryofixation, Endothelium, Blood Vessels, Ultrastructure

Cryofixation refers to the immobilization of tissue components by the rapid reABSTRACT moval of heat from the specimen, so that the structure is interred and stabilized in a natural embedding medium, namely, frozen (amorphous or microcrystalline) tissue water. Cryofixation is now often used as a complement to the more traditional fixation methods, especially when the cell structure is delicate or dynamic and may be inaccurately preserved by the slow selective action of chemical fixatives. Vascular endothelial cells are specialized for transcellular transport and for the regulation of blood flow and composition. The dynamic and labile subcellular organization of these cells, presumably reflecting these functional specializations, makes them ideal candidates for cryofixation. Several different types of endothelial cells were directly frozen at temperatures below 20 degrees Kelvin by pressing them against a liquid-helium-cooled block. These samples were subsequently processed for structural analysis by freeze-substitution. Detailed rationales, designs, and protocols are described for both freezing and freeze-substitution. Electron micrographs of cryofixed arterial and venous capillaries (rete mirabile of the American eel), iliac vein (rabbit), and cultured endothelium from the iliac vein (human) reveal that the organization of the characteristic intracellular membrane system of endothelial vesicles is qualitatively similar to that seen in chemically fixed endothelium, especially with regard to the interconnection of clusters of individual vesicles to form elaborate networks. The luminal and abluminal networks are not in communication, a t least not in static images. Quantitatively, however, most directly frozen endothelial cells have far fewer vesicular profiles than comparable glutaraldehydefixed cells. The differences can be explained by presuming that the rapid action of cryofixation (approximately 1msec) gives a more accurate picture of the vesicular network because it captures the transient structure of labile or dynamic membranes.

INTRODUCTION Living tissue cells exhibit dynamic behavior at all levels of organization. This is known primarily by light microscopic observation of tissues intravitally and cells in culture. Numerous cellular components, however, are observable only by electron microscopy and can be ascribed to have dynamic properties only by inference. For instance, displacement of labeled membrane systems at timed intervals prior to fixation implies movement in living tissue. Such assertions rely on the assumptions that chemical fixatives not only preserve structures with a high degree of fidelity but also immobilize them rapidly enough so that discrete timed displacements can be measured. The effectiveness with which chemical fixatives accomplish this depends upon their speed of penetration into cells and the nature of their chemical reactivity with various macromolecular components of the cell. Glutaraldehyde can penetrate individual cells as rapidly as 0.01 sec (Hopewood, 1967) but for 30-40 sec following application, Brownian and saltatory motions of intracellular components still persist (Buckley, 1973). Also, membrane lipids interact poorly with glutaraldehyde and may remain mobile for periods sufficient to allow fusion of adjacent membranes long after


other cell components have been stabilized (Johnson and Rash, 1981). It is not likely, therefore, that rapid motion and interaction of cell components can be captured in mid-process by chemical cross-linking with glutaraldehyde. Many recent investigations have employed rapid freezing or cryofixation as an alternate means of immobilizing cellular structures. Under the best freezing conditions this essentially accomplishes several important objectives: the immobilization of cell membrane systems and other macromolecular assemblies in milliseconds (Van Harreveld and Crowell, 1964) the retention of pools of soluble molecules and ions in their natural location (Somlyo et al., 1977; Gupta and Hall, 1981) and reduction of other potential sources of artifacts due to chemical fixation (Plattner and Bachman, 1982; Robards and Sleyter, 1985; Sitte et al., 1986; Sitte et al., 1987). Rapid cooling also impedes intrinsic tissue reactions which might lead to artifacts. Thus

Received July 15, 1989; accepted in revised form September 26, 1990. Address reprint requests to Dr.Roger C. Wagner, School of Life and Hea1t.h Sciences, University of Delaware, Newark, DE 19716.


cryofixation increases confidence that ultrastructure is preserved close to the state as it exists under physiological conditions. Vascular endothelial cells are highly specialized structurally and biochemically to perform specific functions according to the vessels in which they are found and the tissue which these vessels serve (Simionescu and Simionescu, 1988). They have come to be regarded as dynamic cellular units rather than a static boundary between the blood and the tissues. They are best regarded as dynamic cellular units rather than a static boundary between the blood and the tissues. Current concepts of structure-function relationships in endothelial cells have been largely derived from ultrastructural analysis of chemically fixed tissues. Due to temporal and physical uncertainties associated with chemical fixation, some of these concepts may be in error or at least misrepresented by ultrastructure. For example, elucidation of the manner in which the membranes of the endothelial vesicular interact and the role they play in transendothelial transport (Wagner and Casley-Smith, 1980) requires that these membranes be fixed in their natural state and that discrete timed tracer experiments be assured by rapid immobilization. It is also not known whether chemical fixation affects the structure of interendothelial junctions and if their conduction of materials across the vessel wall is modulated by physiological control mechanisms. We review here techniques which may be applied for the preparation and cryofixation of the endothelium of blood capillaries and veins in excised tissues and of cultured endothelial cells in monolayers. Protocols for freeze substitution and analysis of thin sections of these frozen tissues are also described. The results are discussed according to their impact on endothelial cell structure-function relationships.

METHODS AND MATERIALS Impact Freezing on a Polished Metal Surface There are several excellent reviews on cryofixation (Plattner and Bachman, 1982; Robards and Sleyter, 1985; Sitte et al., 1987). This article is mainly intended to supplement these with detailed information specific to the direct freezing of endothelium. Nevertheless, a few fundamental concepts should be re-emphasized. The object of cryofixation is to immobilize and stabilize the components of tissue in a natural embedding medium such as solid water. However, the physical properties of water show us that ice can exist in three forms: hexagonal crystalline, cubic crystalline, of amorphous (also “vitreous”) (Mackenzie, 1981; Dubochet et al., 1982). Because the growth of ice crystals damages the delicate membrane systems of cells and promotes phase separations leading to shrinking and swelling of water-filled compartments, only amorphous ice is an appropriate embedding medium. Unfortunately for the experimental biologist, ice is exceedingly difficult to freeze in the amorphous form. In the absence of cryoprotection, vitrification requires the removal of heat from the specimen a t rates exceeding lo4 degrees K/sec (Costello and Corless, 1978; Bald, 1984; Mackenzie, 1981). Tissue damage due to crystalline ice formation can be greatly reduced by pretreatment of


the specimen with cryoprotectants such as glycerol or DMSO but this only introduces additional variables which may have unique artifactual consequences. The best solution is to vitrify the specimen, a demanding task for any freezing apparatus. Numerous technological approaches have been developed to maximize cooling rates of various types of specimens (Steinbrecht and Zierold, 1987). One of the most efficacious of these is rapid-freezing against a cold, mirror-surfaced block of metal, which takes advantage of the large heat capacity and the high thermal conductivity of metal. Essentially a freshly prepared specimen is pressed against a highly polished, flat metal-mirror surface previously cooled with either liquid nitrogen (LN,, 77 degrees K) or liquid helium (LHe, 4 degrees K). A LN,-based device for this purpose was originally demonstrated by Van Harreveld and Crowell (1964). Shortly thereafter, beginning with Van Harreveld’s stay a t the NIH in Richardson’s laboratory and continuing for the next decade, several investigators made significant progress in engineering an instrument for routine and reliable direct freezing. The progress is evident by comparing an early prototype of a freezing machine (Fig. l a ) to one that had evolved by the 1980s (Fig. lb). The LN,-based device shown in Fig. l a was built by Heuser and Reese using many of the parts from Van Harreveld’s and Richardson’s early models; the machine worked well enough to provide excellent results on directly frozen synapses (Heuser et al., 1976), but was somewhat unreliable. The demand for better and more reliable freezing led to the development of the classical HeuserlReese freezing machine, which operated a t temperatures approaching 20 degrees K by using liquid helium as the cryogen (Heuser et al., 1979);the machine shown in Figure l b is a fully functional example of such a device. Several models of freezing machines are now commercially available, including but not limited to the Med. Vac. “Cryopress,” the Biorad E7200 “Slammer,” the Reichert MM80, the Life Cell CF100, and Pelco’s “Gentleman Jim.” Only the first two of these can utilize LHe, which would appear to be the cryogen of choice because of its lower temperature, but which is also substantially more expensive and less available. The advantage of LHe may relate primarily to the depth of good freezing-an advantage that varies from essential t o nonexistent, depending upon the preparation. In any case, excellent results have been demonstrated with LN,-based machines (for example, Sitte et al., 1986). General Comments on Specimen Preparation. Freezing efficiency depends upon several characteristics of the tissue sample such as chemical composition, water content (cells or organelles with a higher content of unbound water will exhibit more rapid crystal growth), and size (more specifically surface area t o volume ratio). Cultured cells in monolayers are very thin, have a high surface area to volume ratio, and if frozen en face will exhibit good freezing throughout. However, for excised tissues of appreciable thickness, even under ideal conditions, only a region extending about 30 pm into the tissue from the freezing surface can be frozen



Fig. 1. Evolution of machines for direct freezing against a metal block. a: Archival photograph of a simple, liquid-nitrogen-based device developed at the NIH during the period 1964-1974 by Van Harreveld, Richardson, Heuser, and Reese. A sample mounted on a n aluminum planchette and attached to the cork head (H) by friction was allowed to drop by gravity onto a cold silver block which was contained a t the bottom of a long brass tube, which in turn was immersed in liquid nitrogen within a standard Dewar. To maintain a dry atmosphere within the brass tube, helium gas from a lecture bottle was passed through a heat exchanger inside the Dewar and then through the tube. The drop rod was fitted with a spring (S) for shock adsorption. After freezing, the head was recovered by disconnecting it from the drop rod at the clamp (D). The machine worked well on occasion,

but was prone to pre-freezing the sample during passage through the long brass tube. b Photograph illustrating the fundamental design concepts of a liquid helium freezing machine. The essential components include a quick-mounting sample head (H), a precision-aligned drop rod (R),a cold block chamber which holds the pure copper block in a dry environment rC,, a liquid helium transfer line (Tj, and antibounce devices, including a catch electromagnet (M) and spring 6). This photograph shows a freezing machine which is a direct descendent of the original Heuser Keese prototype that was used to obtain the micrographs in Heuser et al. (1979,. This machine with minor modifications, is still in use in the Laboratory of Neurobrology, NIH, as are others evolved from the same design.


Fig.2. Exploded view of a freezing sample planchette assembly. A polycarbonate ring and filter paper disc are centered and adhered to an aluminum disc. The disc has tabs for securing it to the freezing head. Prior to freezing, a gelatin cushion disc and the sample to be frozen are placed upon the disc assembly. The combined thickness of the filter paper, cushion, and tissue sample should be only slightly higher than the thickness of the plastic ring.

with a quality sufficient for ultrastructural analysis. At greater depths, ice crystal damage precludes meaningful morphology. For most tissues, this means that peripheral connective tissue must be eliminated and the interior laid bare by slicing so that the cells of interest form the freezing surface and will come to lie well within the zone of good freezing. The amount of time that elapses between excision of the tissue from the animal and freezing must be as short as possible in order to minimize postmortem effects on the cells. Mechanical damage to the specimen during excision should also be avoided. Slicing with a sharp razor blade on a smooth teflon surface appears to minimize tissue tearing and provides a smooth flat surface to freeze. The sliced tissue must be prepared and mounted in such a way that it presents a smooth surface parallel to the freezing block. Sample planchettes are prepared prior to tissue excision and freezing. These consist of an aluminum disk with tabs at opposite sides (available from Biorad), a small circle of Whatman #1 filter paper centered on the disc, and a flat polycarbonate ring, the same diameter as the disc (Fig. 2). Several sample planchettes should be prepared by adhering the filter paper and polycarbonate rings to the aluminum disc with epoxy and Photomount (3M) and curing the resin overnight at 60°C with a flat weight pressing down on the assemblies. Gelatin discs are used in order to cushion the impact of the tissue on the freezing block. These are prepared by pouring warm 5 2 0 % gelatin in buffer Ringer’s (op-


tionally prefixed with 0.1%glutaraldehyde and subsequently washed extensively) into a petri dish to a depth no greater than 1mm and allowing it to solidify. Small discs the same diameter of the filter paper are punched out with a rubber stopper punch or with a blade and kept on ice prior to freezing. Before tissue sample application, a sample planchette is labeled on the back with pencil and attached to the freezing head by bending the aluminum tabs over the beveled edge of the head. The filter paper is wetted with buffer Ringer’s, a gelatin disc laid on top, and the sample applied to the top of the cushion. The amount of residual fluid remaining on the freezing surface of the sample will subtract from the depth to which the underlying tissue is well-frozen. However, too little fluid will result in desiccation of the surface due to exposure to the ambient air with resulting serious artifacts. Touching the tissue surface with a small wick of filter paper just prior to freezing appears to remove just the right amount of fluid. The combined thickness of the cushion and tissue sample should be only slightly higher than the thickness of the plastic ring and represent a volume that will not exceed that of the cavity contained by the plastic ring after impact (Fig. 2). Operation of a Liquid Helium Freezing Machine. Several preparations must be made in order to assure a fully functional freezing device before tissue samples are prepared. Liquid helium (25 liters in a LN,-jacketed Dewar) must be available and should be used as quickly as possible since a full Dewar will not keep for more than a few days. Gaseous N, must be available in order to pressurize the LHe Dewar. The freezing surface of the copper block must be polished to a perfectly flat mirrored surface prior to cooling. Scratches, oxidized copper, or other irregularities will result in areas of poor freezing in the sample. Polishing is best accomplished on a flat glass surface using a fine abrasive such as Wehnol or jewelers rouge. A figure eight motion will assure that the polished surface remains flat. Abrasive should be wiped off using lens paper and the residue removed by sonication in acetone or by soaking in tetrahydrofuran. The polished surface should be examined and critiqued using a loupe or dissecting microscope. The copper block should be stored under Freon in a closed container to prevent oxidation of the surface. The polished surface should never be touched as copper is a soft metal and will scratch easily and because oil from one’s fingers will contaminate the surface. To assemble the apparatus, a Dewar of LHe must be available. There are two configurations for conveniently sized Dewars: older, LN,-insulated, 25-1. Gardner-type Dewars and 30-1. Super Insulated Dewars. The latter are rapidly replacing the former since they are safer, better insulated, and lighter and require no maintenance of the insulation. The Super Insulated Dewar retains LHe for at least two weeks whereas the older type must be used within a few days of filling. The freezing machine is assembled by positioning the Dewar under the working platform and aligning the withdrawal port on the top of the Dewar with a hole in the platform (Fig. 4). A vacuum-insulated transfer tube is inserted into the Dewar with the tip just off the



Fig. 3. A cutaway view of the freezing block assembly of the MedVac Cryopress. The tissue sample-attached to a planchette by a compressible gelatin pad as described in Fig. 2-is secured to the telescoping freezing head, inverted, and attached to the plunger rod. The rod drops by gravity, pressing the sample onto the cold surface of the freezing block. The impact is cushioned by a retracting, spring-loaded shaft in the sample head, by the gelatin pad, and also by shock ab-

sorbers at the corners of the cryoblock chamber at the last moment. A catch electromagnet firmly holds the freezing stage to the block after contact, preventing bouncing of the specimen. A floating seal between the transfer tube and the cold block chamber accommodates the movement of the chamber, relative to the transfer tube, that occurs on impact.

Fig. 5. Preparation of a sample of rete mirabile for freezing. A 1 mm slice is taken from the central region of an excised rete parallel to Fig. 4. A schematic of the liquid helium Dewar and transfer line assembly. A LN,-jacketed, LHe Dewar is tapped with a vacuum-jacketed transfer line. This is secured with a small length of rubber hose between the Dewar spigot and a flange on the line. Gaseous nitrogen is administered through the flange which pressurizes the liquid helium chamber and pushes LHe up through the line and onto the bottom of the cryoblock.

the long axis ofthe capillary bed. This is placed upon a gelatin c ~ h ion on a sample planchette. This orientation assures that a maximum amount of endothelium (at three Capillary diameters) will come to lie within the zone of good freezing.



Fig. 7. After slicing off the embedded filter paper, the block should be trimmed to an oblong rhomboid face which includes only the freezing surface of the sample. Thick sections can then be taken, areas of tissue selected, and the face trimmed smaller for thin sectioning.

Fig. 6. Several freeze-substituted samples can be embedded in the bottom of an aluminum pan with the aluminum discs juxtaposed to the bottom of the pan. The polymerized disc containing the samples is removed and cut into rectangular blocks. These are split or cut across the tissue sample providing two faces for sectioning. The two halves should be of appropriate dimensions for securing in a microtome chuck.

bottom of the helium container. A gaseous helium line is attached to the Dewar through a fitting which may be an integral part of the Dewar or may be on the connecting flange. During a freezing operation, the copper block will be placed in the cryoblock chamber and pressure from the gaseous helium tank (at 6 oz. sq. in.) will be used to spray liquid helium out the transfer line and onto the bottom of the block (Figs. 2,3).Cooling of the block is monitored by a thermocouple or similar device. During cooling, condensation and freezing of moisture from the air onto the cold copper block is prevented by an insulating shutter lid which covers the cryoblock mount. This swings away just prior to impact via an electronically operated trip device which is coordinated with the fall of the freezing head to the block. In order to minimize degradative changes in the tissue, the freezing block should be cooled and a completed planchette assembly attached to the freezing head prior to tissue excision. In this case, it is possible in optimal circumstances to freeze samples with intervals of 20 sec between excision and freezing. After application of the tissue sample and adjustment of the amount of fluid on the sample surface, the planchettehead assembly is quickly inverted and attached to a plunger rod by a bayonet mount (Fig. 3). The plunger rod is released by a trip solenoid and drops by gravity from a height of 18 inches onto the freezing block surface. The drop must be plumb so that the tissue surface meets the surface of the freezing block perfectly parallel since an angled impact will result in a tilted zone of freezing. This is facilitated by drop rod bearings which guide the fall. The impact is cushioned by a spring-loaded shaft in the sample block which retracts as the sample meets the freezing block. Impact is

also dampened by four matched shock absorbers at the four corners of the cryoblock holder (Fig. 3). Following impact of the specimen stage on the cold metal surface any bouncing, however small the amplitude, will seriously reduce the quality of freezing. Bounce suppression is implemented by a strong catch electromagnet on the freezing platform which firmly holds the freezing stage to the platform after contact. Immediately after impact, the electromagnet is released, the sample block removed, and the aluminum planchette containing the frozen specimen plucked off with a forceps while immersed in liquid nitrogen. Cryofixation of Capillary Endothelium In Situ Blood capillaries pervade all tissues except cartilage but are present a t various densities per unit volume of tissue. The limited depth of good freezing preservation requires special considerations in the preparation of vascularized tissues. After excision, it is necessary to trim the tissue in such a way that capillaries will be positioned well within the zone of good freezing. This task becomes easier if the tissue has a relatively high density of capillaries. This is also advantageous in that it permits inspection of numerous capillaries within a single section. Rete systems are ideal for this purpose since they are characterized by closed-packed parallel arrays of blood capillaries involved in countercurrent exchange. The retia mirabile of the eel are paired capillary organs located on the dorso-lateral surface of the swimbladder. The consist of parallel bundles of capillary segments (10 per rete) which function in countercurrent exchange of gases and solutes (Wagner et al., 1987). Each rete is actually two separate but intercalated portal systems: an arterial portal system consisting of thickwalled continuous capillaries and a venous portal system consisting of thin-walled, fenestrated capillaries. We have utilized the rete mirabile of the eel swimbladder to investigate differences between chemically fixed and cryofixed capillary endothelium (Wagner and Andrews, 1985). Preparation and Cryofixation of Capillaries of the Rete Mirabile of the Eel Swimbladder. American eels (Anguilla rostrata) were anesthetized by add-


Fig. 8.


ing tricaine methane sulfonate (MS 222) to the water (500 mglliter) for 30-60 mins. The retia mirabile (red bodies) were excised, cleaned of adhering connective tissue, and sliced with a sharp razor blade on a smooth teflon surface into disc-like portions (1 mm thick) parallel to the long axis of the capillary segments (Fig. 5). Some of the slices were placed in 0.5 M Tyrodes-cacodylate buffer (pH 7.4) containing 2% glutaraldehyde and some in buffer alone. These were allowed to incubate for 30 min at room temperature prior to freezing. Other slices were frozen immediately after excision from the animal with no more than 5 min, elapsing between disruption of the blood supply and freezing. Freezing was accomplished with a metal-mirror freezing device as discussed above. Frozen samples were immediately transferred to LN, for storage before processing.

Preparation and Cryofixation of Large Blood Vessel Endothelial Cells In Situ The endothelial lining of large blood vessels forms an attenuated single layer of cells which, if positioned correctly with respect to the freezing block, can be frozen well throughout its depth. A relatively large vessel is advantageous in that when a portion of its wall is cut out, the radius of curvature will assure that a relatively flat endothelialized surface will be frozen. Adult male albino rabbits were anesthetized with phenobarbital and the iliac vein removed. A 5 mm segment of the vein was cut longitudinally and the exposed intima washed thoroughly with Tyrodes buffer ringers (pH 7.4) but never allowed extended exposure to the air. A smaller portion of the wall of the vein about 1mm by 3 mm was cut out so that the long axis of the endothelial cells was parallel to the long axis of the sample. This permitted one to know the orientation of the endothelial cells in the final embedded specimen. The excised portion of venous wall was laid upon a gelatin cushion on a freezing planchette, the surface wicked with filter paper, and the sample frozen on a Med-Vac Cryopress.


ment avoids the difficulties in sectioning or removing a hard substrate such as plastic or glass. Cells in culture are particularly susceptible to desiccation and should be kept wet or immersed in media or buffer. Endothelial cells were isolated from human iliac veins from brain-dead, heart-beating, cadaver renal donors and were cultured by the method of Jarrell et al. (1984). Human amniotic membrane, taken from fresh placentae, was prepared and immobilized in plastic capsules according to the method of Williams et al. (1985) and seeded with dissociated endothelial cells having undergone 16-25 doublings in culture. Prior to freezing, samples of the amnion with endothelial cells attached were washed with Medium 199E buffer, small portions cut out with a scissors, and placed cell side up on a 1 mm gelatin cushion on an aluminum freezing planchette. The surface of the specimen was touched with a filter paper wick and frozen as above.

Post Examination of Frozen Samples Selection of the most favorable frozen samples was performed by examination with a dissecting microscope while the specimens were still immersed in LN2. Well-frozen samples exhibited a shiny, unbroken surface when illuminated from an angle with a fiber optic light source. A dull, milky surface or one with surface irregularities usually indicate a poorly frozen sample and were discarded. Freeze-Substitution of Cryofured Specimens Freeze-substitution is a process of dehydration in which water is carefully removed from a frozen specimen by replacement with an organic solvent (Van Harreveld and Crowell, 1964; Sitte et al., 1986).It must be carried out at a temperature low enough to prevent secondary ice crystal growth. An organic solvent which is liquid a t this low temperature dissolves the ice in the specimen and gradually replaces it. Chemical fixatives dissolved in the solvent stabilize structure preserved and immobilized by cryofixation. After substitution, Preparation and Cryofixation of Human the temperature can then be raised without risk since Endothelial Cells In Culture water is absent and structure has been fixed in statu Monolayers of cells in culture can also be positioned nascendi. Infiltration with embedding resin is then in such a way that all the cells in a sample will come to performed at room temperature followed by heat polylie well within the zone of good freezing preparation. merization. The resulting plastic embedded specimen We have found that culturing of cells in monolayers on can then be sectioned, stained, and observed by TEM as a soft substrate such as the amnionic membrane per- would a chemically fixed specimen. The lowest temperature at which substitution can be mits selection of an area which can be cut to the requisite size for freezing and which upon final embed- performed is mitigated by the melting point of the solvent used. Substitution is usually carried out with acetone (M.P. -94.4"C) a t -80°C in a low temperature freezer. Slow diffusion rates at low temperatures require that the substitution must be allowed to occur Fig. 8. Survey view of a section taken through a cryofixed sample over extended periods (Ornberg and Reese, 1981; Humof the capillary bed of the rete mirabile at a right angle to the freezing be1 and Muller, 1986). Some form of agitation to the surface (upper right). Cross-sectional profiles of thin-walled fenestrated and thick-walledcontinuous capillaries exhibit good freezing to sample vials should be provided for by a rotary device a depth of about three capillary diameters (16 pm). At greater depths, within the freezer but with a driving mechanism exice crystal reticulationbecomes evident in the interstitial spaces, cap- ternal to the freezing compartment. illary lumenae, and endothelial and red cell nuclei. Compression is The frozen samples are freeze-substituted while still evident at the freezing surface and depths below 16 pm. Impact-reattached to the aluminum planchette. This facilitates lated artifacts include protrusions of red cells (large arrow) and large gaps in the walls of fenestrated capillaries (small arrows). x 7,600. transfer of the specimens between solutions and they Scale marker = 1.3 pm. can be labeled on the back prior to freezing with pencil



Figs. 9-12.


so that particular experimental samples may be identified. At no time should the specimen be picked up with the forceps touching the freezing surface since this will damage the zone of good freezing. Precautions should be taken to avoid the introduction of moisture a t any step in the substitution process since the water will crystalize at low temperature and damage the specimen. All beakers, bottles, syringe barrels and needles, scintillation vials, and other containers used in preparing the solutions should be predried in a hot (60°C)oven. Plastic, translucent scintillation vials with wide mouths and screw caps should be used (one sample per vial) for substitutions at the lower temperatures and glass vials may be used at -20°C and above. Freeze Substitution Protocol. The following protocol, originally used by Bridgman and Reese (19841, was used to freeze substitute rapidly frozen samples of rete mirabile, rabbit vein endothelium, and human endothelial cell monolayers in culture: 1. On the first day, the low temperature freezer is precooled to -80°C (3-4 h). A solution of 10%acrolein (v/v) and 0.2%tannic acid (w/v) in acetone is prepared at a volume sufficient for 5-10 ml per vial. It is critical to avoid water in this solution. Pure reagent-grade acetone (Gold Label-Aldrich Chemical Co.)is drawn with a syringe and long needle through a rubber septum from a fresh bottle. Tannic acid is preweighed and dried in a warm oven before use. Acrolein is dispensed by volume with a dry syringe and needle. After dispensing of this solution into the vials, the bottom of the vials are immersed in liquid nitrogen until the solution is frozen. The vials are then fully immersed in LN, so that it covers the frozen solvent. Metal tongs or forceps are used to handle the vials under LN2 but H,O should not be allowed to condense on the surfaces and contaminate the solution. The tissue discs are transferred to the vials under LN, by grasping only the edges of the disc. The caps are placed loosely on the vials so that the LN, can evaporate. The samples are freeze-substituted overnight a t -80°C (about 14 h). During this time, the LN2 on the top of the frozen solvent melts letting the tissue discs descend to the bottom of the vials.

Figs. 9-12. Higher magnification micrographs of areas in Fig. 8 showing greater detail of freezing effects. Fig. 9. Good freezing is evident in both cellular and extracellular compartments from the freezing surface to a depth of about 10-12 pm. x 20,000. Scale marker = 0.5 pm. Fig. 10. Ice crystal reticulation begins to appear in the interstitial spaces from 12-16 pm and connective tissue cells (arrow) show pronounced reticulation of cytoplasm and nucleoplasm in this region.The endothelial cytoplasm and pericyte processes exhibit little or no ice crystal damage. x 20,000. Scale marker = 0.5 pm. Fig. 11. At 16-20 pm depth, endothelial nuclei begin to show a fine reticulation and ice crystals in the interstitium are more pronounced. x 20,000. Scale marker = 0.5 pm. Fig. 12. Endothelial cytoplasm and vesicular profiles begin to show ice crystal damage beyond 20 pm and residual protein in the capillary lumenae also exhibit effects of ice crystal growth. x 20,000. Scale marker = 0.5 pm.


2. On the second day the freezer is allowed to passively warm t o -50°C which takes 2-3 hrs. The tissue discs are then transferred to fresh acetone (at 5 mlhial) and washed 2-3 times in acetone. This can be done in a fume hood by inserting the vials into crushed dry ice shavings up to the liquid level to maintain temperature. A solution of 1.0% Osmium Tetroxide in acetone is prepared. OSO, crystals can be detached from the side of the vials by immersion in LN, and dispensed (2,250 mg vials) into 50 ml acetone. The tissue discs are added to 5 ml OS04 in each sample vial and returned to the freezer which is set to -20°C and substituted overnight. 3. On day three, the tissue discs are washed three times in acetone at -20°C. This temperature can be approximated by placing the vials on a petri dish on dry ice shavings. A solution of 10% glutaraldehyde in 50% methanol and 50% acetone is prepared. A stock solution of 20% anhydrous glutaraldehyde is diluted with 100%methanol and 100%acetone a t a ratio of 1:l. About 5-10 ml of the 10% glutaraldehyde solution is added to each vial and the temperature is reset to 0°C. The tissue discs should be allowed to equilibrate for at least 6 h. The tissue samples are then washed in anhydrous methanol (Aldrich Chemical Co.) and held a t 0°C for 30-40 minutes each wash. This is necessary to avoid uranyl acetate precipitation in the subsequent treatment. The tissue may detach from the aluminum planchette at this point and will require more careful handling with forceps. Labeling of vials should be done faithfully since the sample marking on the aluminum disc is lost. The tissue discs are then transferred to methanol in acetone (1:l)for 1h and then to a solution of 0.5% uranyl acetate in acetone and substituted overnight at 0°C. The uranyl acetate solution may require sonication in a sonifier bath in order for the uranyl acetate to completely dissolve. 4. On the fourth day, the samples are washed 3 times in acetone while permitting them to warm to room temperature. The samples are then placed in 50% acetone, 50% propylene oxide for 1-2 h, 100%propylene oxide for 1-2 h, and 50% British araldite, 50% propylene oxide for 1-2 h or overnight. 5. On day five, the samples are transferred to 70% araldite, 30% propylene oxide, and then 95% araldite, 5% propylene oxide for 5-6 h each step. 6. On the sixth day, the samples are transferred to 100% araldite with two changes of 5-6 h each. The araldite should be degassed in a continuous vacuum but not in an oven. The samples are polymerized in 100%araldite in shallow metal pans (2-3 samples per pan) at 60°C in a vacuum oven for 24-48 h (Fig. 6). It may be necessary after 1-2 h to reorient the tissue discs which have been displaced upwards by outgassing bubbles. The tissue discs or aluminum planchette should ideally lie juxtaposed to the bottom of the pan with just enough plastic on the top to completely cover the freezing surface of the sample. The pans should be level to avoid a plastic disc whose thickness is not uniform throughout.

Figs. 13 and 14.



Preparation of Embedded Tissue for (Fig. 8), ice crystals are not observable for at least 20 Trimming and Sectioning um from the freezing surface. Good freezing is evident in all tissues at high magUpon polymerization, the plastic dish containing nification to a depth of about 10-12 um (Fig. 9). At several embedded specimens can be easily removed 12-16 um depth, reticulation due to ice crystals begins from the metal pan (Fig. 6). If they have been posito appear in the interstitial spaces between the capiltioned flush with the bottom surface, any retained alularies but the endothelial cell cytoplasm and pericyte minum planchettes can be plucked off the plastic with a forceps. Each sample can be isolated by cutting the processes exhibit little or no ice crystal damage (Fig. plastic disc into sections with a jewelers saw and, de- 10). Connective tissue cells between capillaries in this pending on the density of osmium deposition, the entire region exhibit pronounced reticulation of the nucleosample can be transilluminated with a bright fiber op- plasm as well as the cytoplasm. At 16-20 um depth, tic light source to determine how the sample should be endothelial nuclei begin to show a fine reticulation and sectioned. Two cutting faces can be exposed by scoring ice crystals in the interstitium are more severe (Fig. across the sample, placing a single-edged razor blade 11). Endothelial cytoplasm and vesicular profiles begin across the score, and tapping with a hammer. This to show ice crystal damage beyond 20 um and residual cracks the block in two and leaves a smooth face on two protein in the capillary lumenae also exhibit effects of sides. Alternatively, the sample can be sawed in half ice crystal growth (Fig. 12). Nucleated red blood cells also exhibit ice crystal but this removes a millimeter or so of plastic. Enough damage at depths greater than 20 um (Fig. 8). Red empty plastic should be retained on the opposite end of the block from the sectioning face for insertion into the blood cells often exhibit protrusions or herniations at right angles to the freezing surface (Fig. 8). This may appropriate microtome chuck. A perfectly smooth block face when examined with a be due to a shock wave of impact freezing. At lower dissecting scope indicates homogeneity of the sample depths, thin-walled, fenestrated capillary segments and possible good freezing and is ideal for subsequent have large gaps or discontinuities in their wall which trimming. The filter paper disc should first be sliced off may also be a consequence of impact freezing (Fig. 8). the block face by a cut parallel t o the plane of the Venous Endothelial Cells In Situ sample. The block face should be an oblong rhomboid and include only the freezing surface of the sample The endothelial lining of frozen segments of the (Fig. 7). Thick sections may be taken initially, stained venous wall exhibits an irregular pattern which conwith methylene blue, and examined with a light mi- forms to the scalloped contour of the underlying elastic croscope in order to locate areas of interest. Freezing lamina (Fig. 13). This is probably a consequence of condamage observable at the light level can be used to traction of the wall after excision and the lack of a eliminate the sample or portions thereof for thin sec- pressurized lumen. The endothelial cells appear welltioning. frozen throughout their depth. Endothelial cytoplasmic membrane systems such as golgi, endoplasmic reticulum, and Weibel-Palade bodies as well as microtubules RESULTS appear well-preserved (Fig. 14). The endothelial nuclei Capillary Endothelium In Situ are devoid of ice crystal reticulation and the nuclear Sections taken transversely through the capillary envelope appears normal. More distal to the freezing bed and at right angles to the freezing surface (Fig. 8) surface, the vascular smooth muscle exhibits proexhibit profiles of thick-walled continuous capillaries nounced ice crystal reticulation even a t low magnificaand thin-walled, fenestrated capillaries. Tissue just tion (Fig. 13). In areas on the freezing surface where subjacent to the freezing surface exhibiting some com- overlying fluid is nearly absent, desiccation of fluid pression and disruption probably due to slicing is also buffer media results in concentration of solutes with evident. Little or no residual fluid is present on this resulting more intense staining. surface but appreciable tissue desiccation has not resulted. Deeper in the tissue open capillary lumenae Human Venous Endothelial Cells In Culture indicate little tissue compression. At low magnification Cultured endothelial cells exhibit flattened, attenuated profiles characteristic of their condition in cultured monolayers (Fig. 15). They appear well frozen throughout their depth with no evidence of ice crystal reticulation in either the cytoplasm of nucleoplasm. InFig. 13. Survey view of the endothelial lining of a cryofixed segment of the venous wall. Residual fluid covers the surface of the tracellular membrane systems including rough ER, orendothelium at the freezing surface (upper left) but some desiccation ganelles, and microfilament bundles are well-preis evident in the thinnest areas (arrow). The endothelial lining ap- served (Figs. 16, 17). Cisternae of rough ER appear pears well frozen throughout its depth and the nucleoplasm appears dilated and filled with material (Fig. 16). Surface memdevoid of ice crystals. Deeper in the tissue (lower right), however, vascular smooth muscle exhibits extensive ice crystal reticulation. brane and attached caveolae and vesicular profiles are also well-preserved and without evidence of distortion x 18,000. Scale marker = 0.6 pm. due to freezing damage (Fig. 17). The endothelial cell Fig. 14. Higher magnificationof a portion of an endothelial cell in proper does not lie closely juxtaposed to the underlying Fig. 13. Endothelial cytoplasmic membrane systems such as golgi, ER cisternae, and Weibel-Paladebodies (arrow)appear well-preserved as amniotic material but rests lightly on filamentous bunwell as cytoplasmic microtubules. x 41,400. Scale marker = 0.2 pm. dles arising from the surface (Fig. 15). ~


Figs. 15-17.


DISCUSSION The ultimate goal of ultrastructural analysis is to observe cell structure in a condition close to that in the living state. Noncoagulative chemical fixatives have been a boon for this purpose since they cross-link biological macromolecules instead of precipitating them. However, without suitable controls it cannot be known how rapidly or how well chemical fixatives fix or what postfixation changes might occur. As questions concerning structure-function relationships in cells become more sophisticated and more detail is resolved by electron microscopy, this uncertainty takes on an increasing importance. Cryofixation has not only provided a means of rapidly immobilizing cell structure but also serves as a suitable control for chemical fixation. Comparisons of cryofixed and chemically fixed endothelial cells reveal that glutaraldehyde preserves cell structure very well in a qualitative sense. Cellular membrane systems, organelles, and formed cytoplasmic elements such as filaments and microtubules appear remarkably similar in tissues fixed by either method. Their appear to be few if any endothelial cell structures which are so labile or subject to the effects of chemical fixation that they appear only in cryofixed tissue. However, pronounced qualitative differences have been observed with regard to the membranes of the endothelial vesicular system in several types of blood vessels and in cultured endothelial cells which have been cryofixed by several different procedures (Wagner, 1988). The number of vesicular profiles per unit of endothelial cytoplasm is higher by up to several hundred percent in glutaraldehyde-fixed endothelial cells than in cryofixed cells. This has been shown in capillaries from lung (Mazzone and Kornblau, 19811, diaphragm (Casley-Smith, 1985) and cremaster (Robinson et al., 1984) muscle capillaries and capillaries of the rete mirabile (Wagner and Andrews, 1985). Large blood vessel endothelial cells both in situ (McGuire and Tweitmeyer, 1983)and in culture (Robinson et al., 1984; Wood et al., 1987) exhibit similar effects due to glutaraldehyde fixation. One study (Froekjaer-Jensen and Reese, 1985)

Fig. 15. Low magnification micrograph of a n endothelial cell cultured on a n amniotic membrane and cryofixed by impact freezing. The flat, attenuated profile of the cells in culture favors good freezing throughout the depth of the cells and no evidence of ice crystal reticulation is evident in either the cytoplasm or nucleoplasm. x 18,000. Scale marker = 0.6 pm. Fig. 16. A higher magnification micrograph of upper right region of the cell in Fig. 15 exhibiting well-preserved cisternae of the rough ER which appear filled with material. A cytoplasmic filament bundle appears closely associated with the basal plasma membrane of the cell (arrow). x 30,600. Scale marker = 0.3 pm. Fig. 17. Higher magnification micrograph of the low left region of the cell in Fig. 15 showing well-preserved endothelial caveolae and vesicle profiles and filament bundles in cross section (arrow). x 64,300. Scale marker = 0.15 pm.


detected no changes in vesicular profile number between glutaraldehyde-fixed and cryofixed mesenteric capillaries. This magnitude of change in the amount of membrane comprising the vesicular system implies either a drastic increase in the amount of membrane synthesis or a recruitment of vesicular membrane from a preexisting source brought about by glutaraldehyde. Since the time periods involved in chemical fixation are short, new membrane synthesis would seem to be precluded as an explanation. Vesiculation of plasma membrane or budding of vesicles from intracellular membrane systems could provide sources for additional vesicular profiles. Measurements of the amount of surface membrane present in both glutaraldehyde-fixed and cryofixed rete capillaries (Wagner and Andrews, 1985) and endothelial cells in culture (Wood et al., 1987) indicate no appreciable difference. Glutaraldehyde may therefore induce budding of vesicles from preexisting cisternal membrane systems within the endothelial cell. A rough correlation exists between the thickness of the endothelial cell and the amount of intracellular membrane systems and the degree of difference between the two methods of fixation (Wagner and Andrews, 1985; Froekjaer-Jensen and Reese, 1985). Three dimensional reconstruction of consecutive ultrathin (150 A) serial sections through the capillary wall (Bundgaard et al., 1979; Froekjaer-Jensen, 1980) has revealed that the endothelial vesicular system is comprised of fused clusters of contiguous membrane compartments connected with either lumenal or ablumenal membrane. Such a configuration is not evident in single sections where many apparently free vesicular profiles are observed. It might be supposed that along with an increase in the amount of vesicular membrane, interconnection of vesicles is also an artifactual event brought about by glutaraldehyde fixation. However, such is not the case. In capillaries of the rete mirabile which have been freshly cryofixed, the interconnectedness of the vesicular membranes is nearly total as it is in chemically fixed rete (FroekjaerJensen et al., 1988).Therefore, although a severe quantitative change is brought about by glutaraldehyde fixation, the three dimensional interconnectedness of the vesicles is unaffected. It is almost certain that cryofixation immobilizes macromolecular movement within cells within milliseconds. However, little can be inferred from static electron images about the natural motions and interactions of membrane systems within living cells. The role of the vesicular system in transendothelial and intraendothelial transport thus remains enigmatic (Wagner, 1988). Experiments utilizing intravital electron dense tracers and timed intervals of fixation indicate that the vesicular system does provide a route across endothelial boundaries in living cells. The kinetics and mechanism of this process, however, remain unknown. Future experiments utilizing tracers in endothelial cells should employ rapid freezing methods since vesicles induced by chemical fixation probably bear little relationship to the function of this system in living tissues. Cryofixation of endothelial cells relieves many un-



Meuller, S., and Thorton, S. (1984) Human adult endothelial cell certainties about artifactual effects of chemical fixagrowth in culture. J. Vasc. Surg., 1:757-764. tives but also indicates that quantitative changes can Johnson, T.G.A., and Rash, J.E. (1981) Glutaraldehyde fixation chemoccur if fixation is not rapid enough. Several artifacts istry: Substituted pyridine polymers result from glutaraldehydeare also unique to cryofixation, such as tissue compresamine reactions. J. Cell Biol., 35:213-236. sion, cellular disruption, and ice crystal damage, most Mackenzie, A.P. (1981) Modeling of ultra-rapid freezing of cells and tissues. In Microprobe Analysis of Biological Systems. T.E. of which, however, are detectable as such and can be Hutchinson and A.P. Somlyo, eds. Academic Press, New York, pp. discounted in ultrastructural analysis. Cryofixation is 397-421. a laborious and technically involved method of prepar- Mazzone, R.W., and Kornblau, S.M. (1981) Pinocytic vesicles in the endothelium of rapidly-frozen rabbit lung. Microvascular Res., 21: ing tissues for electron microscopy. However, it in193-211. creases confidence that what is observed represents a McGuire, P.G., and Tweitmeyer, T.A. (1983) Morphology of rapidlycloser semblance of cells in living tissues than chemifrozen aortic endothelial cells: Glutaraldehyde fixation increases cally fixed tissues. the number of caveolae. Circ. Res., 53:424-430.

REFERENCES Bald, W.B. (1984) The relative efficiency of cryogenic fluids used in the rapid quench cooling of biological samples. J. Microsc., 134: 261-270. Bridgman, P.C., and Reese, T.S. (1981) The structure of cytoplasm in directly-frozen cultured cells. I. Filamentous meshworks in the cytoplasmic ground substance. J. Cell Biol., 99:1655-1668. Buckley, I.K. (1973) Studies in fixation for electron microscopy using cultured cells. Lab. Invest., 29398-410. Bundgaard, M., Froekjaer-Jensen, J., and Crone, C. (1979) Endothelial plasmalemmal vesicular profiles as elements in a system of branching invaginations from the cell surface. Proc. Nat. Acad. Sci. U.S.A., 76:6439-6442. Casley-Smith, J.R. (1985) Vesicular form and fusion as revealed by freeze-immobilization and stereoscopy of semithin sections. Prog. Appl. Microcirc. Res., 21:6-20. Costello, M.J., and Corless, J.M. (1978) The direct measurement of temperature changes within freeze-fracture specimens during quenching in liquid coolants. J. Microsc., 11217-37. Dobochet, J., Lepault, J., Freeman, R., Berrimen, J.A., and Homo, J.C. (1982) Electron microscopy of frozen water and aqueous solutions. J . Microsc., 128:219-237. Froekjaer-Jensen, J. (1980) Three dimensional organization of plasmalemmal vesicles in endothelial cells: An analysis be serial sectioning of frog mesenteric capillaries. J. Ultrastructure Res., 73: 9-20. Froekjaer-Jensen, J . , and Reese, T.S. (1985) The plasmalemmal vesicular system in directly-frozen, freeze-substituted frog mesenteric capillaries. J. Physiol. (Lond.), 371384P. Froekjaer-Jensen, J., Wagner, R.C., Andrews, S.B., Hageman, P., and Reese, T.S. (1988) Three-dimensional organization of the plasmalemma1 vesicular system in directly-frozen capillaries of the rete mirabile of the eel. Cell Tissue Res., 25417-24. Gupta, B.L., and Hall, T.A. (1981) The x-ray microanalysis of frozenhydrated sections in scanning electron microscopy: An evaluation. Tissue Cell, 13:623-643. Heuser, J.E., Reese, T.S., Dennis, M.J., J a n , Y., Jan, L., and Evans, L. (1979) Synaptic vesicle exocytosis captured by quick freezing and correlated with quanta1 transmitter release. J. Cell Biol., 81975300. Heuser, J.E., Reese, T.S., and Landis, D.M.D. (1976) Preservation of synaptic structure by rapid freezing. Cold Spring Harbor Symp. Quant. Biol. 15:17-24. Hopwood, D. (1967) Some aspects of fixation with glutaraldehyde. Am. J. Anat., 106:82-92. Humbel, B.M., and Muller, M. (1986) Freeze substitution. In Science of Biological Specimen Preparation. M. Muller, R.P. Becker, and J.J. Wolosewick, eds. Scanning Electron Microsc. Inc., AMF O’Hare, Chicago, pp. 175-183. Jarrel, B.E., Shapiro, S., Williams, S., Carabasi, R.A., Levine, E.,

Ornberg, R.L., and Reese, T.S. (1981) Quick freezing and freeze-substitution for x-ray microanalysis of calcium. In Microprobe Analysis of Biological Systems. T.E. Hutchinson and A.P. Somlyo, eds. Academic Press, New York, pp. 213-223. Plattner, H., and Bachman, L. (1982) Cryofixation: A tool in biological ultrastructure research. Int. Rev. Cytol., 79237-304. Robards, A.W., and Sleyter, U.B. (1985) Low temperature methods in biological electron microscopy. In Practical Methods in Electron Microscopy. A.M. Glauert, ed. Elsevier, Amsterdam, pp. 5-46. Robinson, J.M., Hoover, R.L., and Karnovsky, M.J. (1984) Vesicle (caveolae) number is reduced in cultured endothelial cells prepared for electron microscopy by rapid-freezing. J. Cell Biol., 99:287a. Simionescu, N., and Simionescu, M. (1988) Endothelial Cell Biology in Health and Disease. Plenum Press, New York. Sitte, H., Neumann, K., and Edelman, L. (1986) Cryofixation and cryosubstitution for routine work in transmission electron microscopy. In Science of Biological Specimen Preparation. M. Muller, R.P. Becker, and J.J. Wolosewick, eds. Scanning Electron Microsc. Inc., AMF O’Hare, Chicago, pp. 103-118. Sitte, H., Edelman, L., and Neumann, K. (1987) Cryofixation without pretreatment at ambient pressure. In Cryotechniques in Biological Electron Microscopy. R.A. Steinbrecht and K. Zierold, eds. Springer-Verlag, Berlin, pp. 87-110. Somlyo, A.V., Shuman, H., and Somlyo, A.P. (1977) Elemental distribution in striated muscle fibers and the effects of hypertonicity: Electron probe analysis of cry0 sections. J. Cell Biol., 74:828-857. Steinbrecht, R.A., and Zierold, K. (1987) Cryotechniques in Biological Electron Microscopy. Springer-Verlag, Berlin and New York. Van Harreveld, A,, and Crowell, J . (1964) Electron microscopy after rapid freezing on a metal surface and substitution fixation. Anat. Rec., 149:381-386. Wagner, R.C. (1988) Ultrastructural studies of capillary endothelium: Compartmental tracing, high voltage electron microscopy and cryofixation. In: Endothelial cell Biology in Health and Disease. N. Simionescu and M. Simionescu, eds. Plenum Press, New York, pp. 23-47. Wagner, R.C., and Andrews, S.B. (1985) Ultrastructure of the vesicular system in a rapidly-frozen capillary endothelium of the rete mirabile. J . Ultrastructure Res., 90:172-182. Wagner, R.C., and Casley-Smith, J.R. (1980) Endothelial vesicles. Microvascular Res., 21:267-298. Wagner, R.C., Froehlich, R., Hossler, F.E., and Andrews, S.B. (1987) Ultrastructure of cauillaries in the red body (rete mirabile) of the eel swimbladder. M&rovascular Res., 34349-362. Williams, S.K., Jarrel, B., Friend, L., Radomski, J., Carabasi, R., Koolpe, E., Mueller, S., Thornton, S., Marinucci, T., and Levine, E. (1985) Adult human endothelial cell compatibility with prosthetic graft material. J . Surg. Res., 38518-629. Wood, M.R., Wagner, R.C., Andrews, S.B., Greener, D.A., and Williams, S.K. (1987) Rapidly-frozen endothelial cells in culture: Comparison of vesicles in prefixed vs fresh-frozen samples. Microcirc. Endothelium Lymphatics, 3:323-358.

Cryofixation of vascular endothelium.

Cryofixation refers to the immobilization of tissue components by the rapid removal of heat from the specimen, so that the structure is interred and s...
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