This presentation was supported by a Clinical Innovator Award to JLC from the Flight Attendant Medical Research Institute and by the United States Environmental Protection Agency. Although the research described in this article has been funded wholly or in part by the United States Environmental Protection Agency through Cooperative Agreement CR83346301 with the Center for Environmental Medicine, Asthma, and Lung Biology at The University of North Carolina at Chapel Hill, it has not been subjected to the Agency&#8217;s required peer and policy review, and therefore does not necessarily reflect the views of the Agency and no official endorsement should be inferred. Mention of trade names or commercial products does not constitute endorsement or recommendation for use.

1. Preparation of Biological Specimens for Freeze-fracture/Freeze-etch
<ol>
	<li>Use a conventional EM fixative formulation such as 2% glutaraldehyde + 2% paraformaldehyde in 0.1 M phosphate buffer for 1 hr. to perform primary fixation of biological tissues. NOTE: While it may be desirable to freeze some types of specimens without prior fixation, universal blood borne pathogen precautions mandate appropriate fixation where the specimen consists of human tissue.</li>
	<li>Following primary fixation, rinse the specimen in the same buffer supplemented with 0.2 M sucrose for no more than 1 hr.</li>
	<li>Transfer the specimen to a cryoprotectant solution consisting of 25% glycerol in 0.1 M phosphate buffer for no more than 1 hr.</li>
</ol>
2. Freezing and Storage of Specimens In Advance of Freeze-fracture
<ol>
	<li>Select gold or copper specimen stubs of appropriate size and shape for the specimen being processed&#160;(Figure 4).</li>
	<li>Using fine grade forceps and/or small gauge syringe needles position small fragments of the specimen on the top of a metal specimen stub.
	<ol>
		<li>Reduce the liquid content of the specimen to a sticky, slightly glue like consistency by drawing off liquid from the edge of the stub with filter paper.</li>
		<li>For single replicas, use small gauge syringe needles to create a mound of tissue on the stub. For double replicas, invert another stub and position it exactly over the stub and place lightly onto the specimen surface creating a sandwich (Figure 5A). Do not press the specimen out from between the two stubs.</li>
	</ol>
	</li>
	<li>Fill two insulated dewar vessels with liquid nitrogen. NOTE: The first vessel accommodates a metal post containing a small well which is used to liquefy a small volume of propane gas from a commercially available cylinder (Figure 5B). The second vessel is a partitioned storage/holding vessel for briefly maintaining frozen specimens under liquid nitrogen.
	<ol>
		<li>Using the cylinder nozzle positioned in the propane well, open the cylinder valve and allow gas to flow into the well of the first vessel where it will liquefy. NOTE: CAUTION!! PROPANE IS EXPLOSIVE, MAY CAUSE FREEZING INJURY ON CONTACT, AND IS AN ASPHYXIANT. Use propane only in a chemical hood certified for such use taking care also to avoid extraneous ignition sources, using garments protective against low temperatures, and maintaining adequate ventilation.</li>
		<li>As quickly as possible, pick up the specimen mount with fine grade forceps and plunge it into the liquefied gas in the first vessel for several seconds then quickly transfer to the second holding vessel of liquid nitrogen (Figure 5B).</li>
		<li>Once the specimens are frozen, store under liquid nitrogen until ready to transfer to the freeze-fracture plant. Transfer the stubs to a larger storage liquid nitrogen Dewar container for extended storage if desired, however care should be taken not to allow the stubs to thaw.</li>
	</ol>
	</li>
</ol>
3. Operation of the Freeze-fracture Instrument and Specimen Processing
<ol>
	<li>Loading specimens on the brass holder and into the chamber
	<ol>
		<li>Load the gold specimen stubs into a booklet-type double replica specimen holder or clamp device under liquid nitrogen and maintain there until ready to position in the specimen chamber (Figure 6).</li>
		<li>When the chamber has achieved high vacuum (approximately 2 x 10-6 mbar) and the stage has been cooled to liquid nitrogen temperature, turn off the pump, vent the chamber, and position the mounted specimen stubs onto the specimen table as quickly as possible.</li>
		<li>Turn on the pump, reestablish high vacuum, and use electronic stage and arm temperature controls to adjust the specimen table temperature to -100 &#176;C and direct liquid nitrogen to the microtome arm and bring it to liquid nitrogen temperature.</li>
	</ol>
	</li>
	<li>The fracture process
	<ol>
		<li>Upon achieving high vacuum, a stage temperature of -100 &#186;C and microtome arm at liquid nitrogen temperature, remotely open the double replica specimen holder thereby fracturing the specimens on the specimen mounts (freeze-fracture_movie1.mov).</li>
		<li>In the case of specimens intended for etching following fracture, use a razor blade maintained in a clamp on the microtome arm to shave (i.e., fracture) the surface of the specimens in lieu of step 3.2.1 (freeze-etch_movie2.mov).</li>
		<li>If the added freeze-etching step is to be performed, position the cooled microtome arm over the fractured surfaces for one to several minutes. NOTE: This maneuver will sublime (i.e., etch) water from the fractured surface. The effect of this maneuver is to reveal true cell surfaces, extracellular matrix, and/or molecular assemblies by sublimation of water, in addition to fracture faces passing through the lipid bilayer of membranes.</li>
	</ol>
	</li>
	<li>Metal shadowing and replica support evaporation using electron guns 
	<ol>
		<li>Activate the platinum/carbon electron or resistance gun and allow it to evaporate a thin layer (approximately 2 nm) of platinum/carbon over the fractured surface from an angle of 30&#186; - 45&#186;. This typically requires approximately 15 - 20 sec.</li>
		<li>Activate the carbon electron or resistance gun as in step 3.3.1 and evaporate a thin layer of carbon to the fractured specimen surface from directly overhead (90&#186;) to give support to the replica. This typically requires approximately 15 - 20 sec.</li>
	</ol>
	</li>
	<li>Retrieval of replicas
	<ol>
		<li>Upon completion of replication shadowing and carbon stabilization, turn off the vacuum pump, vent the chamber, and remove the specimen mount from the instrument.</li>
		<li>Using a pair of fine grade forceps remove each gold specimen stub from the double replica booklet/clamp and allow to thaw for several seconds before gently lowering the stub onto a water surface in a spot plate (Figure 7). NOTE: The carbon/platinum replica containing adherent specimen material will float onto the water surface.</li>
		<li>Maneuver the replicas using either a fine wire loop or a conventional copper electron microscope grid held by fine grade forceps and transfer to another spot plate containing 5% sodium dichromate in 50% sulfuric acid for one to several hours. NOTE: This bath digests the actual biological specimen material from the replica. Full strength household bleach may be substituted for the dichromate/sulfuric acid solution.</li>
		<li>When the digestion is complete, transfer the replica back to a clean water surface and retrieve onto a standard copper electron microscopy grid. (NOTE: Replicas may fragment into smaller pieces during digestion but even very small replicas may contain substantial information and should be retrieved and examined.) Store replica supporting grids in commercially available grid boxes where they will remain stable for years with gentle handling.</li>
	</ol>
	</li>
</ol>
4. Ultrastructural Examination of Freeze-fracture/etch Replicas
<ol>
	<li>View replicas in a transmission electron microscope at an accelerating voltage typically from 50 - 80 kV.</li>
	<li>Record relevant images on standard TEM film or with a high resolution digital camera.</li>
</ol>

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The author has nothing to disclose.

In the years following its introduction and commercial availability, freeze-fracture/etch procedures were widely utilized for investigations of biological membrane structure. Indeed, the best perspectives of some of the structural specializations of membranes have been obtained in freeze-fracture/etch preparations. These studies not only contributed to understanding of the structural organization of membranes but also provided insights into how structure and function are related.
The advent of routine biological electron microscopy brought about an awareness that chemical treatments with the aldehyde fixatives, reactive chemicals, and heavy metals required to confer differential electron density to biological specimens themselves contributed to the generation of artifact. The development of freeze-fracture technology was based in part on an effort to reduce processing artifact. Nevertheless, it has been recognized that freeze-fracture/etch introduces its own set of artifacts, a major one of which is ice crystal damage. Tissues plunged directly into liquid nitrogen experience a reduced cooling rate due to the formation of an insulating layer of nitrogen gas surrounding the specimen which results in the formation of ice crystals that obliterate ultrastructural detail. This problem is overcome with the use of cryoprotectants such as glycerol and by freezing the tissue first in liquid nitrogen-cooled propane. Historically, plunge freezing was performed in liquid nitrogen cooled chlorofluorocarbons but these materials have been phased out of use due to their adverse effects on the environment. Even in the best cryoprotective conditions, the nature of the process inevitably results in a coarse, gravel like surface in cytoplasmic fractures; however, fractures through membrane structures reveal well organized features having specific planar organizations relative to their orientation in the cell membrane. These images are possible only with the use of cryoprotectant agents such as glycerol that limit ice crystal damage, and glycerol itself in some cases also may introduce artifactual redistribution of membrane associated particles. Moreover, glycerol cannot be sublimated from the fractured surface thus limiting the opportunity for successful etching. If the subsequent etching procedure is desirable, then the specimen must be either rapidly frozen in a fashion that limits ice crystal damage using either liquid helium or nitrogen slush and/or approached by the use of a cryoprotectant such as acetone which is readily sublimated from fractured surfaces.
Although Steer&#8217;s initial report was focused on imaging viruses, it is in membrane structure that freeze-fracture/etch found more extensive application. Indeed, structures such as tight and gap junctions and ciliary membrane specializations have been elegantly revealed in freeze-fracture preparations. Freeze-fracture has found considerable utility in achieving an understanding of membrane differentiation during development15,17,24 and in membrane pathology25-30. Previous reports from this laboratory, have demonstrated the pattern of differentiation of tight junctional complexes during airway development, disruption of ciliary membrane structures associated with experimental infectious agent exposure, and degradation of tight junctional complexes in airway epithelial cells associated with air pollutant and tobacco smoke exposure in experimental animals and human subjects.
Technical Extensions of the Freeze-Fracture/Freeze-etch Procedure
The quenching method using liquefied propane to freeze specimens described here is perhaps the most common approach to freezing biological specimens for conventional freeze-fracture. However, other approaches to freezing must be appropriated where freeze-etching is to be performed. Since glycerol is a non-etchablecryoprotectant, specimens for freeze-etching must be frozen in ways that limit ice crystal formation. This may be accomplished by using an etchablecryoprotectant such as acetone and/or by freezing the specimen mount against a metal surface cooled with liquid helium or liquid nitrogen slush. Rapid freezing without cryoprotectants also may be accomplished using spray freezing or high pressure freezing in liquefied propane.
Recent efforts to further relate structure and function using freeze-fracture technology have appropriated this technique in concert with immunohistochemical labeling31,32. Here, antibodies to specific antigens can be effectively and accurately localized to membrane sites (Figure 11).
While freeze-fracture with unidirectional shadowing is by far the most widely used protocol, some experimental designs will call for rapid freezing with little or limited cryoprotection, accompanied by etching of the fractured surface to reveal true cell surfaces, extracellular matrix, and/or macromolecular complexes, and rotary shadowing to impart contrast16 (Figure 12 and Figure 13).
There is a growing interest in applications of freeze-fracture in nanotechnology, materials science, and in using this technique as an imaging tool in molecular biology. Indeed, nanomaterials in many ways may be consistent in their organization with viral structures for which freeze-fracture was originally envisioned as a means for imaging. Moreover, manufactured liposomes, structures similar in form to biological membranes, are being widely studied for targeted delivery of pharmaceutical agents and directed gene therapy. Because their organization is essentially membrane like, freeze-fracture processing may have considerable utility in producing revealing images as a component in developing these therapies33.
In summary, freeze-fracture technologies have been of historic importance in characterizing the organization and function of cell membranes. However, freeze-fracture is finding new areas of utility in coordination with molecular biology and nanotechnology and almost certainly will contribute to advances in these rapidly emerging fields.

The concept and practical application of freeze-fracture processing of biological specimens was introduced by Steere1 over half a century ago. The early apparatus appropriated disparate components into a working self-contained unit1. The original apparatus was modified and refined into commercially available instruments in order to accommodate the critical need for remote manipulation, maintenance of high vacuum, and the evaporation of carbon and metals to produce a replica suitable for examination by transmission electron microscopy (Figure 1 and Figure 2).
The typical instrument consists of a high vacuum chamber with specimen table and microtome arm having regulable liquid nitrogen throughputs (Figure 1). The chamber also houses two electron guns, one for stabilizing carbon evaporation positioned at a 90&#186; angle to the specimen stage and the other for platinum/carbon shadowing at an adjustable angle, typically 15&#186; - 45&#186; (Figure 2). Power to the unit is applied to operate the vacuum pump and electronic panels regulate temperature adjustment and electron gun control.
Originally conceived as a means to achieve improved imaging of viruses, freeze-fracture gained even more popularity as a technique for the examination and analysis of cell membranes and their specializations2,3. Indeed, this procedure has been integral to elucidating structure/function relationships in cells and tissues and many of these studies stand as classic contributions to cell and molecular biology4-9. The major goal and rationale for the development of the freeze-fracture technique was to limit artifacts observable at electron microscopic resolution deriving from chemical fixation and processing used in conventional biological electron microscopy. Here the goal is to limit chemical fixation and to freeze the specimen with sufficient speed and frequently in the presence of a cryoprotectant in order to limit ice crystal formation and other freezing artifacts. More recently, this technique has found a resurgence of interest from molecular biologists and materials science investigators for examination of nanoparticles and nanomaterials.
Freeze-fracture and freeze-etch images exhibit a three-dimensional character and sometimes are mistaken for scanning electron micrographs. However, freeze-fracture preparations are examined by transmission electron microscopy and their major contribution to high resolution morphologic studies is their unique representation of structure/function elements of cell membranes. Freeze-fracture processing is initiated by freezing cells and tissues with sufficient speed to limit ice crystallization and/or with the use of cryoprotectant agents such as glycerol. The specimens are then fractured under vacuum and a replica is generated by evaporation of carbon and platinum over the fractured surface. The original specimen is digested from the replica which is retrieved onto a standard EM specimen grid. Another common misinterpretation of freeze-fracture images is that they depict cell surfaces. However the basic premise of freeze-fracture is that biological membranes are split through the lipid bilayer by the fracture process (Figure 3). This process in biological membranes yields two fracture faces, one which reveals the organization of the half of the membrane adjacent to the cytoplasm, the PF-face, and one which reveals the half of the bimolecular leaflet of the membrane that is adjacent to the extracellular milieu, the EF-face. True cell surfaces are not represented in freeze-fracture images but only appear when the subsequent added step of freeze-etching following the fracture procedure is used. In order to effectively etch previously fractured specimens to reveal surface detail, specimens must be frozen at a rapid rate and without unetchablecryoprotectant. Etching of water from the surface of the fractured specimen revealing underlying features is accomplished by positioning the cooling microtome arm over the specimen stage creating a temperature differential between the stage holding the specimen and the cooling microtome arm which causes water to sublime from the surface. When water is sublimed from the surface of the fractured specimen during the freeze-etching maneuver, then aspects of actual cell surfaces, extracellular matrix, cytoskeletal structures, and molecular assemblies may be revealed at high resolution. Thus freeze-fracture and freeze-etch are not interchangeable terms but rather reflect a step-wise process the latter of which may not be necessary or desirable depending on the needs of the particular study.
Following the freeze-fracture/freeze-etch procedures, the fractured surfaces are subjected to directed evaporative coats of carbon and platinum in order to provide support and imaging contrast to the replica. The platinum/carbon imaging evaporation may be unidirectional or rotary and is accomplished by either resistance or electron guns. Unidirectional shadowing from a known angle, typically 30&#186; - 45&#186;, is useful in performing certain morphometric calculations. Specimens that have been subjected to deep-etching typically are rotary shadowed and the resultant images of these specimens are photographically reversed for evaluation.
The historical as well as a present goal of the freeze-fracture/freeze-etch technique is to limit chemical fixation and processing specimen artifacts that are associated with more conventional transmission electron microscopy procedures. However, this technique provides a substantive advantage in its ability to confer three-dimensional detail and thus facilitate acquisition of morphometric data in biological, material science, and nanotechnology specimens. Freeze-fracture and freeze-etch procedures are complex and multi-faceted and some aspects of its application are customized. This presentation offers a survey view of the major features of the process and the reader is referred to comprehensive published protocols10,11 in order to address the details and customize the process for specific research needs.

<table border="0" cellpadding="0" cellspacing="0" width="573">
	<tbody>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">Name of Material/ Equipment</td>
			<td style="width:71px;">Company</td>
			<td style="width:119px;">Catalog Number</td>
			<td style="width:167px;">Comments/Description</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">Balzers Freeze-fracture/freeze-etch plant</td>
			<td style="width:71px;">Balzers</td>
			<td style="width:119px;">BAF400T</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">Standard buffers</td>
			<td style="width:71px;">various suppliers</td>
			<td style="width:119px;"></td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">standard aldehyde fixatives</td>
			<td style="width:71px;">various suppliers</td>
			<td style="width:119px;"></td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">sodium dichromate</td>
			<td style="width:71px;">various suppliers</td>
			<td style="width:119px;"></td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">sulfuric acid</td>
			<td style="width:71px;">various suppliers</td>
			<td style="width:119px;"></td>
			<td></td>
		</tr>
		<tr height="84">
			<td height="84" style="height:84px;width:216px;">Disposable supplies for Platinum/Carbon Evaporation</td>
			<td style="width:71px;">Technotrade International</td>
			<td style="width:119px;"></td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">Liquid nitrogen</td>
			<td style="width:71px;">various suppliers</td>
			<td style="width:119px;"></td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">Freon</td>
			<td style="width:71px;">various suppliers</td>
			<td style="width:119px;"></td>
			<td></td>
		</tr>
		<tr height="84">
			<td height="84" style="height:84px;width:216px;">Disposable supplies for electron microscopy</td>
			<td style="width:71px;">Electron Microscopy Sciences</td>
			<td style="width:119px;"></td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">Transmission electron microscope</td>
			<td style="width:71px;">Carl Zeiss Inc.</td>
			<td style="width:119px;"></td>
			<td></td>
		</tr>
	</tbody>
</table>

fundamental technical elements of freeze-fracture/freeze-etch in biological electron microscopy

Freeze-fracture/freeze-etch describes a process whereby specimens, typically biological or nanomaterial in nature, are frozen, fractured, and replicated to generate a carbon/platinum &#8220;cast&#8221; intended for examination by transmission electron microscopy. Specimens are subjected to ultrarapid freezing rates, often in the presence of cryoprotective agents to limit ice crystal formation, with subsequent fracturing of the specimen at liquid nitrogen cooled temperatures under high vacuum. The resultant fractured surface is replicated and stabilized by evaporation of carbon and platinum from an angle that confers surface three-dimensional detail to the cast. This technique has proved particularly enlightening for the investigation of cell membranes and their specializations and has contributed considerably to the understanding of cellular form to related cell function. In this report, we survey the instrument requirements and technical protocol for performing freeze-fracture, the associated nomenclature and characteristics of fracture planes, variations on the conventional procedure, and criteria for interpretation of freeze-fracture images. This technique has been widely used for ultrastructural investigation in many areas of cell biology and holds promise as an emerging imaging technique for molecular, nanotechnology, and materials science studies.

The key premise of freeze-fracture image interpretation is that fracture planes pass through the lipid bilayer of membranes conferring two fracture faces, called by convention the PF-face (plasma fracture-face) and EF-face (extracellular fracture-face) (Figure 3). The PF-face is the half of the membrane lipid bilayer adjacent to the cytoplasm of the cell and the EF-face is the half of the membrane lipid bilayer adjacent to the extracellular milieu. The freeze-fracture technique is particularly useful for the investigation of membrane structure and the two faces are typically distinctively different with PF-faces being populated with many membrane associated particles in contrast to EF-faces which contain fewer. The cytoplasmic contents of freeze-fractured cells are typically coarse in appearance and are not particularly revealing although membrane bound organelles such as nuclei and Golgi can be readily identified. Of particular interest to investigators using this technique are specializations of membrane structure that can be interpreted for their function. These include specific distributions of membrane associated particles, ciliary membrane specializations, and intercellular junctions.
Cilia-associated Structures
Residing in the cell membrane at the base of each eukaryotic cilium and flagellum is an array of membrane associated particles organized into strands encircling the shaft of the cilium and known as the ciliary necklace (Figure 8)12-16. This structure exhibits some degree of morphologic heterogeneity in various species and it has been speculated to be multifunctional inasmuch as it appears prior to the emergence of the ciliary shaft during ciliogenesis and remains in place in the mature cilium. The nascent ciliary necklace derives from the aggregation and organization of membrane particle arrays prior to the time of the emergence of the ciliary shaft during ciliogenesis suggesting its possible participation in early organization of the developing axoneme14.
Tight Junctions and Epithelial Permeability
In various epithelia, freeze-fracture studies reveal a sophisticated organization of anastomosing strand and groove structures encircling the apical aspect of the basolateral borders. This belt-like structure is known as the tight junction or zonulaoccludens (Figure 9) and is believed to act as a regulator of epithelial permeability to the paracellular flux3, 17-20. Tight junctions with few strands have been described as &#8220;leaky&#8221; while those with more complex organization are described as &#8220;tight&#8221;. This organization appears to fit the function of the epithelia to which they are associated. The PF-faces of tight junctions appear as strands while the EF-faces exhibit complementary grooves.
Gap Junctions and Intercellular Communication
Gap junctions (Figure 10) appear in freeze-fracture preparations of a variety of tissues and are represented by dense arrays of particles on PF-faces and complementary pits on EF-faces. These structures are thought to represent membrane sites that facilitate intercellular communication. In tissues where they are present, demonstrations of the passage of low molecular weight fluorescent dyes and calcium fluxes have been demonstrated21-24 suggesting that they represent a physical route for the passage of signaling molecules.
<img alt="Figure 1" fo:content-width="5in" src="/files/ftp_upload/51694/51694fig1highres.jpg" width="500" /> 
Figure 1.&#160;Layout of a commercial freeze-fracture/etch plant. At far left is a pressurized Dewar feeding cooling liquid nitrogen to the specimen stage and cooling microtome arm under controlled conditions. At center is the unit with specimen chamber and electronic control panels to the right. At right is an additional liquid nitrogen tank on which the vent gas is used to bring the specimen chamber to atmosphere.
<img alt="Figure 2" fo:content-width="5in" src="/files/ftp_upload/51694/51694fig2highres.jpg" width="500" /> 
Figure 2.&#160;Freeze-fracture/etch plant specimen chamber with door opened to reveal positions of directionally aimed electron guns, cooling shroud, and specimen post where specimens in brass holders are positioned. Specimens are introduced to the chamber through a port to the left of the chamber.
<img alt="Figure 3" fo:content-width="5in" src="/files/ftp_upload/51694/51694fig3highres.jpg" width="500" /> 
Figure 3.&#160;Diagrammatic representation of the principle of freeze-fracture. The fracture faces derive from the splitting of the membrane through the lipid bilayer producing an extracellular face (EF-face) and a face that is proximal to the cytoplasmic aspect, the PF-face. True unfractured membrane surfaces are seen only by etching subsequent to the freeze-fracture maneuver.
<img alt="Figure 4" fo:content-width="5in" src="/files/ftp_upload/51694/51694fig4highres.jpg" width="500" /> 
Figure 4.&#160;Two types of specimen mounts for freeze-fracture/etch procedures. (Left) A double replica specimen holder booklet is used for conventional freeze-fracture procedures performed without etching. The specimen is sandwiched between two gold specimen stubs and positioned in the brass booklet. The specimen is fractured by opening the booklet under vacuum in the specimen chamber (See freeze-fracture_movie1.mov). (Right) A mount used primarily for freeze-etching. The specimen is positioned on the gold stubs and locked into the brass holder. The specimen is fractured by gentle shaving with a razor blade fixed in the microtome arm. This configuration is preferred for etching procedures (See freeze-etch_movie2.mov).
<img alt="Figure 5" fo:content-width="5in" src="/files/ftp_upload/51694/51694fig5highres.jpg" width="500" /> 
Figure 5.&#160;Preparation of a freeze-fracture specimen mount. The specimen is sandwiched between two gold mounts (A) and frozen in a well containing liquid nitrogen cooled propane (foreground) with subsequent transfer to a holding Dewar containing liquid nitrogen (background) (B).
<img alt="Figure 6" fo:content-width="5in" src="/files/ftp_upload/51694/51694fig6highres.jpg" width="500" /> 
Figure 6.&#160;Demonstration of positioning of a frozen freeze-fracture specimen into the double replica specimen holder booklet in preparation for introduction to the specimen chamber of the freeze-fracture/etch plant.
<img alt="Figure 7" fo:content-width="5in" src="/files/ftp_upload/51694/51694fig7highres.jpg" width="500" /> 
Figure 7.&#160;Retrieval of replicas following fracture and shadowing in the freeze-fracture/etch plant. &#8220;A&#8221; is a clean water surface in a spot plate. The replica floated from the specimen stub is transferred to &#8220;B&#8221; a well containing an acidified solution of sodium dichromate or full strength household bleach for digestion of the tissue from the replica. &#8220;C&#8221; represents subsequent clean water surfaces to which the cleaned replica is transferred prior to being retrieved onto a copper electron microscopy grid.
<img alt="Figure 8" fo:content-width="5in" src="/files/ftp_upload/51694/51694fig8highres.jpg" width="500" /> 
Figure 8.&#160;Cilia related structures revealed by freeze-fracture. White arrows point to specialized membrane particle arrays on the luminal border of epithelial cells undergoing ciliogenesis. These particle arrays represent nascent ciliary necklaces which reside at the bases of emergent and mature cilia (black arrow). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/51694/51694fig8highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 9" fo:content-width="5in" src="/files/ftp_upload/51694/51694fig9highres.jpg" width="500" /> 
Figure 9.&#160;Fracture through epithelial cell membranes revealing examples of PF- and EF-fracture faces. The strand and groove organization of tight junctional complexes is evident and the intercellular space is marked by arrows. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/51694/51694fig9highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 10" fo:content-width="5in" src="/files/ftp_upload/51694/51694fig10highres.jpg" width="500" /> 
Figure 10.&#160;A freeze-fracture image of a gap junction in rat liver. Particles are evident on the PF-face and complementary pits are evident on the EF-face. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/51694/51694fig10highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 11" fo:content-width="5in" src="/files/ftp_upload/51694/51694fig11highres.jpg" width="500" /> 
Figure 11.&#160;Immunohistochemical localization of a connexin using a colloidal gold labeled antibody on a freeze-fracture preparation illustrating specific localization on the gap junction. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/51694/51694fig11highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 12" fo:content-width="5in" src="/files/ftp_upload/51694/51694fig12highres.jpg" width="500" /> 
Figure 12.&#160;An example of rapid freezing with deep etching and rotary shadowing. The epithelial cell has been fractured through the cytoplasm and a PF-face is evident (arrowheads). Behind the arrowheads can be seen a true cell surface revealed by subsequent etching. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/51694/51694fig12highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 13" fo:content-width="5in" src="/files/ftp_upload/51694/51694fig13highres.jpg" width="500" /> 
Figure 13.&#160;An example of rapid freezing with deep etching and rotary shadowing to reveal molecular complexes comprising a bovine tracheal ciliaryaxoneme. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/51694/51694fig13highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Fundamental Technical Elements of Freeze-fracture/Freeze-etch in Biological Electron Microscopy at JoVE.com

Fundamental Technical Elements of Freeze-fracture/Freeze-etch in Biological Electron Microscopy

Basic techniques and refinements of freeze-fracture processing of biological specimens and nanomaterials for examination by transmission electron microscopy are described. This technique is a preferred method for revealing ultrastructural features and specializations of biological membranes and for obtaining ultrastructural level dimensional and spatial data in materials sciences and nanotechnology products.

Freeze-fracture/freeze-etch describes a process whereby specimens, typically biological or nanomaterial in nature, are frozen, fractured, and replicated ...

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33.1K Views.  The University of North Carolina at Chapel Hill. Basic techniques and refinements of freeze-fracture processing of biological specimens and nanomaterials for examination by transmission electron microscopy are described. This technique is a preferred method for revealing ultrastructural features and specializations of biological membranes and for obtaining ultrastructural level dimensional and spatial data in materials sciences and nanotechnology products.

Video: Fundamental Technical Elements of Freeze-fracture/Freeze-etch in Biological Electron Microscopy

Preparation of embryos for Electron Microscopy of the Drosophila embryonic heart tube

The morphogenesis of the Drosophila embryonic heart tube has emerged as a valuable model system for studying cell migration, cell-cell adhesion and cell shape changes during embryonic development. One of the challenges faced in studying this structure is that the lumen of the heart tube, as well as the membrane features that are crucial to heart tube formation, are difficult to visualize in whole mount embryos, due to the small size of the heart tube and intra-lumenal space relative to the embryo. The use of transmission electron microscopy allows for higher magnification of these structures and gives the advantage of examining the embryos in cross section, which easily reveals the size and shape of the lumen. In this video, we detail the process for reliable fixation, embedding, and sectioning of late stage Drosophila embryos in order to visualize the heart tube lumen as well as important cellular structures including cell-cell junctions and the basement membrane.

Preparation of embryos for Electron Microscopy of the Drosophila embryonic heart tube

We describe a process for fixation, embedding, sectioning, and imaging of late stage Drosophila embryos for Trasmission Electron Microscopy of the embryonic heart tube. This technique allows for the visualization of the heart tube lumen as well as the basement membrane, which lines the lumen of the heart.

The morphogenesis of the Drosophila  embryonic heart tube has emerged as a valuable model system for studying cell migration, cell-cell adhesion and cell ...

Time-lapse Microscopy of Early Embryogenesis in Caenorhabditis elegans

Caenorhabditis elegans has often been used as a model system in studies of early developmental processes. The transparency of the 
embryos, the genetic resources, and the relative ease of transformation are qualities that make C. elegans an excellent model for early embryogenesis. Laser-based 
confocal microscopy and fluorescently labeled tags allow researchers to follow specific cellular structures and proteins in the developing embryo. For example, 
one can follow specific organelles, such as lysosomes or mitochondria, using fluorescently labeled dyes. These dyes can be delivered to the early embryo by means 
of microinjection into the adult gonad. Also, the localization of specific proteins can be followed using fluorescent protein tags. Examples are presented here 
demonstrating the use of a fluorescent lysosomal dye as well as fluorescently tagged histone and ubiquitin proteins. The labeled histone is used to visualize 
the DNA and thus identify the stage of the cell cycle. GFP-tagged ubiquitin reveals the dynamics of ubiquitinated vesicles in the early embryo. Observations 
of labeled lysosomes and GFP:: ubiquitin can be used to determine if there is colocalization between ubiquitinated vesicles and lysosomes. A technique for 
the microinjection of the lysosomal dye is presented. Techniques for generating transgenenic strains are presented elsewhere (1, 2). For imaging, embryos 
are cut out of adult hermaphrodite nematodes and mounted onto 2% agarose pads followed by time-lapse microscopy on a standard laser scanning confocal 
microscope or a spinning disk confocal microscope. This methodology provides for the high resolution visualization of early embryogenesis.

Time-lapse Microscopy of Early Embryogenesis in Caenorhabditis elegans

This article describes a technique for the visualization of the early events of embryogenesis in the nematode Caenorhabditis elegans.

Caenorhabditis elegans has often been used as a model system in studies of early developmental processes.  The transparency of the 
embryos, the genetic ...

Microscopy of Fission Yeast Sexual Lifecycle

The fission yeast Schizosaccharomyces pombe has been an invaluable model system in studying the regulation of the mitotic cell cycle progression, the mechanics of cell division and cell polarity. Furthermore, classical experiments on its sexual reproduction have yielded results pivotal to current understanding of DNA recombination and meiosis. More recent analysis of fission yeast mating has raised interesting questions on extrinsic stimuli response mechanisms, polarized cell growth and cell-cell fusion. To study these topics in detail we have developed a simple protocol for microscopy of the entire sexual lifecycle. The method described here is easily adjusted to study specific mating stages. Briefly, after being grown to exponential phase in a nitrogen-rich medium, cell cultures are shifted to a nitrogen-deprived medium for periods of time suited to the stage of the sexual lifecycle that will be explored. Cells are then mounted on custom, easily built agarose pad chambers for imaging. This approach allows cells to be monitored from the onset of mating to the final formation of spores.

We provide a reproducible basic method for the long-term microscopy of the fission yeast sexual lifecycle. With minor adjustments described, the presented protocol allows research focus on different steps of the reproductive process.

The fission yeast Schizosaccharomyces pombe has been an invaluable model system in studying the regulation of the mitotic cell cycle progression, the ...

Characterization of Calcification Events Using Live Optical and Electron Microscopy Techniques in a Marine Tubeworm

Characterizing the first event of biological production of calcium carbonate requires a combination of microscopy approaches. First, intracellular pH distribution and calcium ions can be observed using live microscopy over time. This allows identification of the life stage and the tissue with the feature of interest for further electron microscopy studies. Life stage and tissues of interest are typically higher in pH and Ca signals. 
Here, using H. elegans, we present a protocol to characterize the presence of calcium carbonate structures in a biological specimen on the scanning electron microscope (SEM), using energy-dispersive X-ray spectroscopy (EDS) to visualize elemental composition, using electron backscatter diffraction (EBSD) to determine the presence of crystalline structures, and using transmission electron microscopy (TEM) to analyze the composition and structure of the material. In this protocol, a focused ion beam (FIB) is used to isolate samples with dimension suitable for TEM analysis. As FIB is a site specific technique, we demonstrate how information from the previous techniques can be used to identify the region of interest, where Ca signals are highest.

We demonstrate the use of various microscopy methods that are useful in observing the calcification of a tubeworm, Hydroides elegans, as well as locating and characterizing the first calcified material. Live microscopy and electron microscopy are used together to provide functional and material information that are important in studying biomineralization.

Characterizing the first event of biological production of calcium carbonate requires a combination of microscopy approaches. First, intracellular pH ...

Preparation and Observation of Thick Biological Samples by Scanning Transmission Electron Tomography

This report describes a protocol for preparing thick biological specimens for further observation using a scanning transmission electron microscope. It also describes an imaging method for studying the 3D structure of thick biological specimens by scanning transmission electron tomography. The sample preparation protocol is based on conventional methods in which the sample is fixed using chemical agents, treated with a heavy atom salt contrasting agent, dehydrated in a series of ethanol baths, and embedded in resin. The specific imaging conditions for observing thick samples by scanning transmission electron microscopy are then described. Sections of the sample are observed using a through-focus method involving the collection of several images at various focal planes. This enables the recovery of in-focus information at various heights throughout the sample. This particular collection pattern is performed at each tilt angle during tomography data collection. A single image is then generated, merging the in-focus information from all the different focal planes. A classic tilt-series dataset is then generated. The advantage of the method is that the tilt-series alignment and reconstruction can be performed using standard tools. The collection of through-focal images allows the reconstruction of a 3D volume that contains all of the structural details of the sample in focus.

This report describes a sample preparation protocol and specific imaging conditions for performing scanning transmission electron tomography of thick biological specimens.

This report describes a protocol for preparing thick biological specimens for further observation using a scanning transmission electron microscope. It ...

Biological Sample Preparation by High-pressure Freezing, Microwave-assisted Contrast Enhancement, and Minimal Resin Embedding for Volume Imaging

The described sample preparation technique is designed to combine the best quality of ultrastructural preservation with the most suitable contrast for the imaging modality in a focused ion beam scanning electron microscope (FIB-SEM), which is used to obtain stacks of sequential images for 3D reconstruction and modelling. High-pressure freezing (HPF) allows close to native structural preservation, but the subsequent freeze substitution often does not provide sufficient contrast, especially for a bigger specimen, which is needed for high-quality imaging in the SEM required for 3D reconstruction. Therefore, in this protocol, after the freeze substitution, additional contrasting steps are carried out at room temperature. Although these steps are performed in a microwave, it is also possible to follow traditional bench processing, which requires longer incubation times.The subsequent embedding in minimal amounts of resin allows for faster and more precise targeting and preparation inside the FIB-SEM. This protocol is especially useful for samples that require preparation by high-pressure freezing for a reliable ultrastructural preservation but do not gain enough contrast during the freeze substitution for volume imaging using FIB-SEM. In combination with the minimal resin embedding, this protocol provides an efficient workflow for the acquisition of high-quality volume data.

Here we present a protocol for combining two sample processing techniques, high-pressure freezing and microwave-assisted sample processing, followed by minimal resin embedding for acquiring data with a focused ion beam scanning electron microscope (FIB-SEM). This is demonstrated using a mouse tibial nerve sample and Caenorhabditis elegans.

The described sample preparation technique is designed to combine the best quality of ultrastructural preservation with the most suitable contrast for the ...

The CryoAPEX Method for Electron Microscopy Analysis of Membrane Protein Localization Within Ultrastructurally-Preserved Cells

Key cellular events like signal transduction and membrane trafficking rely on proper protein location within cellular compartments. Understanding precise subcellular localization of proteins is thus important for answering many biological questions. The quest for a robust label to identify protein localization combined with adequate cellular preservation and staining has been historically challenging. Recent advances in electron microscopy (EM) imaging have led to the development of many methods and strategies to increase cellular preservation and label target proteins. A relatively new peroxidase-based genetic tag, APEX2, is a promising leader in cloneable EM-active tags. Sample preparation for transmission electron microscopy (TEM) has also advanced in recent years with the advent of cryofixation by high pressure freezing (HPF) and low-temperature dehydration and staining via freeze substitution (FS). HPF and FS provide excellent preservation of cellular ultrastructure for TEM imaging, second only to direct cryo-imaging of vitreous samples. Here we present a protocol for the cryoAPEX method, which combines the use of the APEX2 tag with HPF and FS. In this protocol, a protein of interest is tagged with APEX2, followed by chemical fixation and the peroxidase reaction. In place of traditional staining and alcohol dehydration at room temperature, the sample is cryofixed and undergoes dehydration and staining at low temperature via FS. Using cryoAPEX, not only can a protein of interest be identified within subcellular compartments, but also additional information can be resolved with respect to its topology within a structurally preserved membrane. We show that this method can provide high enough resolution to decipher protein distribution patterns within an organelle lumen, and to distinguish the compartmentalization of a protein within one organelle in close proximity to other unlabeled organelles. Further, cryoAPEX is procedurally straightforward and amenable to cells grown in tissue culture. It is no more technically challenging than typical cryofixation and freeze substitution methods. CryoAPEX is widely applicable for TEM analysis of any membrane protein that can be genetically tagged.

This protocol describes the cryoAPEX method, in which an APEX2-tagged membrane protein can be localized by transmission electron microscopy within optimally-preserved cell ultrastructure.

Key cellular events like signal transduction and membrane trafficking rely on proper protein location within cellular compartments. Understanding precise ...

Serial Block-Face Scanning Electron Microscopy (SBF-SEM) of Biological Tissue Samples

Serial block-face scanning electron microscopy (SBF-SEM) allows for the collection of hundreds to thousands of serially-registered ultrastructural images, offering an unprecedented three-dimensional view of tissue microanatomy. While SBF-SEM has seen an exponential increase in use in recent years, technical aspects such as proper tissue preparation and imaging parameters are paramount for the success of this imaging modality. This imaging system benefits from the automated nature of the device, allowing one to leave the microscope unattended during the imaging process, with the automated collection of hundreds of images possible in a single day. However, without appropriate tissue preparation cellular ultrastructure can be altered in such a way that incorrect or misleading conclusions might be drawn. Additionally, images are generated by scanning the block-face of a resin-embedded biological sample and this often presents challenges and considerations that must be addressed. The accumulation of electrons within the block during imaging, known as "tissue charging," can lead to a loss of contrast and an inability to appreciate cellular structure. Moreover, while increasing electron beam intensity/voltage or decreasing beam-scanning speed can increase image resolution, this can also have the unfortunate side effect of damaging the resin block and distorting subsequent images in the imaging series. Here we present a routine protocol for the preparation of biological tissue samples that preserves cellular ultrastructure and diminishes tissue charging. We also provide imaging considerations for the rapid acquisition of high-quality serial-images with minimal damage to the tissue block.

Serial Block-Face Scanning Electron Microscopy SBF-SEM of Biological Tissue Samples

This protocol outlines a routine method for using serial block-face scanning electron microscopy (SBF-SEM), a powerful 3D imaging technique. Successful application of SBF-SEM hinges on proper fixation and tissue staining techniques, as well as careful consideration of imaging settings. This protocol contains practical considerations for the entirety of this process.

Serial block-face scanning electron microscopy (SBF-SEM) allows for the collection of hundreds to thousands of serially-registered ultrastructural images, ...

Cryo-Electron Microscopic Grid Preparation for Time-Resolved Studies using a Novel Robotic System, Spotiton

The capture of short-lived molecular states triggered by the early encounter of two or more interacting particles continues to be an experimental challenge of great interest to the field of cryo-electron microscopy (cryo-EM). A few methodological strategies have been developed that support these &#34;time-resolved&#34; studies, one of which, Spotiton-a novel robotic system-combines the dispensing of picoliter-sized sample droplets with precise temporal and spatial control. The time-resolved Spotiton workflow offers a uniquely efficient approach to interrogate early structural rearrangements from minimal sample volume. Fired from independently controlled piezoelectric dispensers, two samples land and rapidly mix on a nanowire EM grid as it plunges toward the cryogen. Potentially hundreds of grids can be prepared in rapid succession from only a few microliters of a sample. Here, a detailed step-by-step protocol of the operation of the Spotiton system is presented with a focus on troubleshooting specific problems that arise during grid preparation.

The protocol presented here describes the use of Spotiton, a novel robotic system, to deliver two samples of interest onto a self-wicking, nanowire grid that mix for a minimum of 90 ms prior to vitrification in liquid cryogen.

The capture of short-lived molecular states triggered by the early encounter of two or more interacting particles continues to be an experimental ...

Strategies for Optimization of Cryogenic Electron Tomography Data Acquisition

Cryogenic electron tomography (cryoET) is a powerful method to study the 3D structure of biological samples in a close-to-native state. Current state-of-the-art cryoET combined with subtomogram averaging analysis enables the high-resolution structural determination of macromolecular complexes that are present in multiple copies within tomographic reconstructions. Tomographic experiments usually require a vast amount of tilt series to be acquired by means of high-end transmission electron microscopes with important operational running-costs. Although the throughput and reliability of automated data acquisition routines have constantly improved over the recent years, the process of selecting regions of interest at which a tilt series will be acquired cannot be easily automated and it still relies on the user's manual input. Therefore, the set-up of a large-scale data collection session is a time-consuming procedure that can considerably reduce the remaining microscope time available for tilt series acquisition. Here, the protocol describes the recently developed implementations based on the SerialEM package and the PyEM software that significantly improve the time-efficiency of grid screening and large-scale tilt series data collection. The presented protocol illustrates how to use SerialEM scripting functionalities to fully automate grid mapping, grid square mapping, and tilt series acquisition. Furthermore, the protocol describes how to use PyEM to select additional acquisition targets in off-line mode after automated data collection is initiated. To illustrate this protocol, its application in the context of high-end data collection of Sars-Cov-2 tilt series is described. The presented pipeline is particularly suited to maximizing the time-efficiency of tomography experiments that require a careful selection of acquisition targets and at the same time a large amount of tilt series to be collected.

The increasing demand for large-scale data collection in cryogenic electron tomography requires high-throughput image acquisition routines. Described here is a protocol that implements the recent developments of advanced acquisition strategies aimed at maximizing the time-efficiency and throughput of tomographic data collection.