[Methods in Cell Biology] Cellular Electron Microscopy Volume 79 || Cryopreparation Methods for...

CHAPTER 2

METHODS IN CELL BIOLCopyright 2007, Elsevier Inc.

Cryopreparation Methods for ElectronMicroscopy of Selected Model Systems

Kent McDonaldElectron Microscope LaboratoryUniversity of CaliforniaBerkeley, California 94720

I. I

OGY,All rig

ntroduction

VOL. 79 0091hts reserved. 23 DOI: 10.1016/S0091

-679X-679X

II. E
quipment and Materials A. H igh-Pressure Freezers B. S pecimen Carriers C. A ccessories D. C ryoprotectants/Fillers E. T ools Useful for Most HPF Specimen Loading Operations
III. G
eneral Rules for Loading Samples for HPF A. W ork Only with Healthy, Unstressed Cells B. W ork Quickly C. D o Not Let Your Cells/Tissues Dry Out D. A void Surrounding Your Cells/Organisms with Aqueous Media E. A void Mechanical Damage F. F ill the Carrier Correctly G. U se the Smallest Volume of Sample You Can
IV. M
ethods for Specific Organisms A. Y easts (Saccharomyces cerevisiae and Schizosaccharomyces pombe) B. N ematodes (C. elegans) C. D rosophila Embryos
V. P
ostfreezing Processing A. F reeze-Substitution B. E mbedding C. F S and Embedding for Tomography D. F S and Embedding for Immunocytochemistry E. A Case Study of Adjusting FS Variables to Improve Visualization F. S ectioning
/07 $35.00(06)79002-1

24 Kent McDonald

G.
M icroscopy and Evaluation of Results H. A rtifacts of HPF
VI. S
ummary R eferences
Specimen preparation for electron microscopy (EM) is a critically important process

because distortions or artifacts introduced at this stage will be represented in the final

images taken on the microscope. Nowhere is this more important than in electron

tomographic three-dimensional (3D) modeling and analysis of cells because the 3D

reconstruction eVorts can takeweeks or evenmonths to complete.Molecular and cell

biologists working on model systems are increasingly turning to EM to analyze

mutant phenotypes or to immunolocalize molecules at high resolution. In this

chapter, we describe in detail EM specimen preparation methods for three of the

most powerful and popular model systems in today’s biology: Saccharomyces cere-

visiae,Caenorhabditis elegans, andDrosophila melanogaster.Each of these organisms

is diYcult to fix for EM by conventional methods that use buVered fixatives at room

temperature because they each have formidable diVusion barriers that hinder the freeexchange of reagents. However, when low-temperature fixation methods such as

high-pressure freezing and freeze-substitution are used, the preservation of cellular

ultrastructure can be excellent. We discuss the basic equipment for high-pressure

freezing and how to use it, what to do with frozen material to prepare it for routine

EM analysis, cellular tomography, or EM immunolabeling. We suggest some strate-

gies for improving the visualization of particular cytoplasmic components such as

membranes, and comment briefly on the problem of artifacts of cryopreparation

methods.

I. Introduction

Structural biology today is increasingly turning to 3D methods at the highest

possible resolution to visualize biological ultrastructure from molecules to whole

cells. Small objects such as individual molecules or molecular assemblies are

regularly analyzed in vitreous ice by crystallographic or single particle averaging

methods (Nogales and GrigorieV, 2001). Even cell organelles and extremely small

whole cells can be studied in ice using cryo-electron tomography (cryo-ET) (Lucic

et al., 2005). With larger cells, however, it is diYcult to use these methods because

the electron microscope beam will not readily penetrate them, and high-quality

image formation is impossible (McIntosh et al., 2005). As a result, some kind of

sectioning is required. An excellent, but technically diYcult choice is to freeze

whole cells or tissues and then cut them into sections at very low temperatures;

the resulting sections can be imaged in cryo-transmission electron microscopy

(cryo- TEM) ( Al-Amo udi et al ., 2004; Zhang et al., 2004 ; Chapter 15 by Dubo chet

2. Cryopreparation Methods for Electron Microscopy 25

et al., this volume). This method, however, is fraught with problems, particularly

for serial reconstruction of large volumes, so a more productive approach is to

cryoimmobilize cells, freeze-substitute them into resins, and cut resin sections at

room temperature for viewing in conventional TEMs. The resulting resolution

may be slightly less than with vitreous cryosections, but it is suYcient for the study

of a far greater number of questions in cell biology. Perhaps the best available

method for analyzing resin sections at high resolution is ET (McIntosh et al.,

2005). In this chapter, we will emphasize methods that result in resin-embedded

cells that can be viewed by conventional TEM or ET.

Preserving ultrastructure with the greatest fidelity and at the highest possible

resolution is essential for structural work that will improve our understanding of

the interactions among molecules in a cell. It has been known for years that

cryofixation methods for electron microscopy (EM) specimen preparation give

the closest approximation to the cell’s native state (see historical accounts in:

Echlin, 1992; Gilkey and Staehelin, 1986; Robards and Sleytr, 1985; see Murk

et al., 2003 for a comparison of conventionally fixed and high-pressure frozen

cells). However, only the smallest cells can be fixed without ice crystal damage by

plunging them into a cryogen, and even powerful methods like impact freezing can

preserve only a few micrometers of cell depth. High-pressure freezing (HPF) is

therefore the best available method for cryofixation of most cells and tissues. HPF

has been around for about 20 years (Moor, 1987), but it has not yet been widely

adopted by the EM community. True, the necessary machines are relatively

expensive, but so are many other pieces of EM equipment. EM laboratories

typically spend from one-half to several million dollars for microscopes and

related instrumentation. It is ironic, then, that the distortions of cellular fine

structure introduced by conventional methods of specimen processing can be

orders of magnitude greater than the resolving power of the microscopes. We

look forward to the day when more laboratories have adopted cryomethods for

routine EM analysis because the quality of EM data, and thus our understanding

of cell fine structure, will benefit. In this chapter, we will give detailed methods for

improving preservation of ultrastructure in some of the most popular and power-

ful cell systems now being studied. Coincidentally, these are cell types that aremore

than usually diYcult to fix by conventional EM specimen preparation methods.

How does HPF work and why is it a better cryofixation choice than other types

of freezing? HPF was developed by Moor and Riehle and first reported at the

European Congress of Electron Microscopy in Rome in 1968 (Moor, 1987; Moor

and Riehle, 1968). The impetus to develop such a technique was probably related

to the freeze-fracture work that was very popular at the time. Because freeze-

fracture requires the freezing of a large volume of sample (so it can more easily be

fractured), it was then common to get freezing artifacts (ice damage) unless some

cryoprotectant was used. HPF is a method that can, in theory, freeze samples that

are as much as 600-mm thick without ice damage, even when no cryoprotectant

is used. This greatly expands the types of cells and tissues that can be studied

by fast-freezing methods, and this remains the reason why HPF is currently so

26 Kent McDonald

important, although freeze-fracture itself is rarely used these days. In practice, we

now know that the 600-mm figure is overly optimistic and that 100–300 mm is a

more realistic working figure for most types of sample, especially if some non-

penetrating cryoprotectant is added to the sample before freezing (Dubochet,

1995; Sartori et al., 1993). However, as long as the size of any ice crystals that

form is smaller than the resolution of the EM method being used, the procedure

will usually be considered acceptable.

HPF works by lowering the freezing point and suppressing the rate of ice crystal

nucleation and growth (Moor, 1987). This follows from the fact that the volume of

water increases as it crystallizes. The rapid application of a high hydrostatic

pressure will inhibit this expansion, slowing the crystallization that occurs with

cooling and letting the water become immobilized in a vitreous state before

crystals can form. The optimal pressure is around 2045 bar and HPF machines

are designed to operate in this range. HPF is not as rapid as some other freezing

methods. Impact cooling may reach rates of 106 K/sec at the surface, where HPF

is about 5 � 104 K/sec. At distances away from the surface the rate slows rapidly,

so at the center of a 600-mm sample the rate may only be a few hundred Kelvin per

second (Moor, 1987). In practice, this means that HPF is not a good method for

arresting fast kinetic events such as synaptic vesicle transmission (Heuser and

Reese, 1981). There is also the currently unexplored possibility that fast molecular

events, like protein polymerization, are aVected by HPF in ways that may lead to

misleading images.

II. Equipment and Materials

A. High-Pressure Freezers

There are currently three types of commercially available high-pressure freezers:

the BAL-TEC HPM 010, the Leica EM PACT2, and the Wohlwend HPF

Compact 01. Space does not permit a detailed description of each instrument,

but for the BAL-TEC HPM 010 see Moor (1987) and for the EM PACT see

Studer et al. (2001). There is no published account of the Wohlwend HPF

Compact 01, but it is probably similar in design to the BAL-TEC HPM 010

because Wohlwend was an engineer for this machine. In the subsequent discus-

sions, these two machines will be considered together because the loading and

specimen carriers are interchangeable.

One significant diVerence between the EM PACT machine and the others is its

portability. It sits on a cart with wheels, is significantly smaller and lighter than the

others, and can be moved from place to place, for example, to a sophisticated light

microscope where it can be used for correlative light and electron microscope

studies. All of these machines work well with a variety of materials, and the choice

of which to use is mostly related to the freezing needs of the users and how these

match the capabilities of the machines. In the end, it is not the machine that

matters most for achieving good freezing. They are like computers in that what


comes out is directly related to what goes in. Specimen loading, on the other hand,

is far more critical to the successful freezing of samples. In the sections that follow,

we will show the variety of specimen loading containers that are available and how

best to fill these for freezing.

B. Specimen Carriers

The most significant aspects of specimen carrier design are the heat transfer

properties of its materials and its geometry. Among common metals, copper and

aluminum have the best heat transfer properties and most carriers are made of one

or the other. Gold is another good choice because it is inert and will not react with

cells or their growth media solutions, but it is more expensive. Sapphire is a

specimen loading accessory that is used for growing cells in culture, and it is an

excellent choice because its heat transfer properties are almost twice as good as

copper or aluminum and it is transparent.

1. BAL-TEC HPF 010 and Wohlwend HPF Compact 01

a. Cup-Shaped Specimen CarriersThe most common geometry of sample holder for HPF is a simple, round cup.

In the BAL-TEC and Wohlwend machines, the cups are 2 mm in diameter, and

they range in depth from 100 to 300 mm. Because two cups are fitted together to

make a chamber, one can have well depths from 100 to 500 mm in 50-mmincrements and a 600-mm-deep well.

b. Jet Freezing Device CarriersIn theWohlwendHPFCompact 01 system, there is a variant on the simple cup that

is a rectangular-shaped depression 200-mm deep in a thin layer of copper (Walther,

2003). These are holders for the BAL-TEC JFD 030 jet freezer device that can be

used for both the Wohlwend and BAL-TEC HPF machines by using a special

specimen holder tip available from Engineering OYce Wohlwend, Sennwald,

Switzerland. To cover the well, there is a flat-sided copper piece that fits on the top.

A possible advantage of this type of specimen holder is that the walls are thinner than

the conventional sample holder for the Wohlwend/BAL-TEC machines. In theory,

this should lead to faster cooler rates at the surface of the sample, although if the

sample is thicker than 100 mm, it may have no eVect on the cooling rates at the center

of the sample (Shimoni and Muller, 1998; Studer et al., 1995).

c. Specialty CarriersIn the BAL-TEC Consumables catalog (http://www.bal-tec.com/products/

CONSUMABLES.htm), there are a variety of specialized holders available for

specialized applications such as freeze-fracture or freezing in gold tubes between

two clamp rings (Shimoni and Muller, 1998).

http://www.bal-tec.com/products/CONSUMABLES.htm

http://www.bal-tec.com/products/CONSUMABLES.htm

28 Kent McDonald

2. Leica EM PACT2

a. Cup-Shaped CarriersThe specimen cup diameters in the EM PACT machines are smaller than those

of the other HPF machines. They are 1.2–1.5 mm in diameter and have depths of

100, 200, and 400 mm. These cups are not used in pairs, so the depth combinations

are more restricted than with the BAL-TEC styles. In the original cup design, the

pressurizing fluid hit the sample directly and caused deformation of the material

in line with the hole. Subsequently, the Leica engineers have devised a new type

of cup that has a thin, flexible membrane across the cup and this transmits the

pressure from the pressurizing fluid to the material in the cup. These new cups

come in depths of 100 and 200 mm, are 1.5-mm wide, and are referred to as

membrane carriers. They have a further advantage over the previous versions of

the Leica cup specimen carriers in that they are much thinner, which reduces the

total thermal mass and probably improves the freezing rates.

b. Tube-Shaped CarriersIn the EM PACT, there is a copper tube 16-mm long with an inner diameter of

300 mm that can be used as a capillary tube to draw up cell suspensions for

freezing. It will also hold 200-mm diameter cellulose capillary tubing (Hohenberg

et al., 1994) filled with cells. This arrangement can give excellent freezing, as long

as the air space between the cellulose tubing and copper tube is filled with

hexadecene or some other fluid that will eVectively transfer heat. This two-step

process has the advantage that the distribution of biological material pulled into

the cellulose tubing can be checked before freezing. With the copper tubing alone,

it is impossible to see if there are trapped air bubbles or whether the material is

tightly packed in the tube. The major application of these copper tubes is for

vitreous cryosectioning. Once frozen, the tubes can be transferred to a cryoultra-

microtome where the copper is trimmed away with a diamond-trimming tool. The

resulting block face can be sectioned at�160 �C and the sections viewed directly in

a cryo-TEM (Al-Amoudi et al., 2004).

c. Live-Cell CarriersWith the addition of an optional rapid transfer system (RTS) to an EM PACT2

machine, users have the ability to do correlative light microscopy (LM) and EM

fixation by HPF with a time resolution of about 5 sec. To accomplish this, it is

necessary to modify the stage of an inverted light microscope to accept a special

stage that will hold the RTS loader. The live-cell carrier is loaded into the tip of the

RTS loader and inserted into the stage for observation by LM. At the moment

when the operator wants to fix the cells, the RTS loader is removed from the light

microscope and inserted into the RTS module, where it is automatically secured

into the freezer specimen carrier holder, inserted into the HPF and frozen. Further

detai ls of co rrelative LM/E M by this machi ne can be found in Chapt er 4 by

Muller-Reichert et al., this volume.


d. Specia lty Car riersThe Leica system also ha s specia lty carri ers for sampling by micr obiopsy needle

(Hohe nberg et al ., 1996 ) or for freeze-f racture or cryo-s canning EM.

C. Accesso ries

Thr ee impor tant ac cessories to use with HPF specim en carri ers are sapphir e

disks, cell ulose micro dialysis tubing, and a micro biopsy gun. Thes e are used with

certain types of cells and tissues as explained be low.

1. Sapphir e disk s. As mention ed earli er, sapphir e has an extre mely goo d

coe Y cient of heat transfer, almos t twice as good as the meta ls us ed for specim en

carriers. Sap phire can be mach ined into smal l disks that fit into the specim en

carriers of both types of HPF machi nes. Thes e disks hav e excell ent optical proper-

ties and can thu s be used as substr ates for grow ing tissue cultur e cells ( Hess et al .,

2000; Reipert et al ., 2004a ). The quality of cell gro wth can be easily checked in a

light microscop e; in the EM PACT2 wi th RTS , they can be observed up until 5 sec

before freez ing.

If the cell s on sap phire disks are process ed by freez e-subs titutio n (FS) and

embedded in resin, the sapph ire disk is present in the resi n after polyme rization.

Methods have been worke d out for removi ng the sapphir e disk from the cells in

the resi n and these can be found in McDon ald et al. (2006) .

2. Micro dialysis tubi ng . The idea of using kidney dialy sis tubing as a carrier

for smal l cell s an d even small multicel lular organ isms was first proposed by

Hohenberg et al. (1994). The tubing has an inner diameter of about 200 mm and

a pore exclusion size of about 10-kDa MW. The trick to successfully using the

tubing is learning how to crimp the ends so that the cells do not get lost during

process ing. This is exp lained in more detai l in Chapt er 4 by M u ller-R eichert et al. ,

this volume.

3. Microbiopsy device. Hohenberg et al. (1996) were also the first to publish a

method using a very small biopsy needle to facilitate rapid sampling of soft tissues.

This works much better than cutting up delicate organ tissues with razor blades

or scalpels or large punches because it is faster and causes less mechanical damage.

The EM PACT2 HPF has a special biopsy loading station that also speeds up the

loading.

D. Cryoprotectants/Fillers

Many of the samples loaded into specimen carriers do not completely fill the

cavity; something must be used to fill these spaces or else the samples will not

freeze well. Fillers should have the following properties:

1. They should transfer heat eVectively during cooling. Aqueous solutions

are to be avoided because they turn into ice. As this part of the sample

30 Kent McDonald

crystallizes, the latent heat of fusion must be removed, greatly slowing the

cooling of adjacent regions of the sample. Moreover, once ice forms it

becomes a poor conductor of heat.

2. They should be excluded from the cells, so the organization of the biological

sample is not altered or compromised by their presence.

3. They should be physiologically compatible with the cells or tissues.

4. They should have cryoprotective properties, that is, bind water molecules

such that they will not rearrange into ice.

5. They should separate from the cells/tissues after FS, either by dissolving in

the FS solvent or physically separating from the samples.

Cryoprotectants/fillers are classified as either extracellular, because they do

not penetrate into cells, or intracellular, because they are penetrating (Gilkey and

Staehelin, 1986). Some common extracellular cryoprotectants include: 1-hexadecene,

yeast paste, Escherichia coli paste, cold water fish gelatin, sucrose, serum albumin,

and high molecular weight carbohydrate compounds such as dextran and ficoll.

Some common intracellular cryoprotectants include: glycerol (5–15%), methanol,

ethanol, dimethyl sulfoxide, and ethylene glycol.

The issue of cryoprotectants for HPF is one that needs to be considered very

carefully. The nonpenetrating cryoprotectants are to be preferred because they

should disrupt the internal cell structure less than penetrating cryoprotectants. But

these compounds are not always easy to use, or they may have consequences for

downstream processing. For example, dextran works well as a cryoprotectant, but

after FS, it forms a hard shell around whatever tissue it surrounds, sometimes

making sectioning diYcult. Unless physically removed or fractured under liquid

nitrogen, hexadecene can form a barrier to free exchange of solvents during FS

because it does not dissolve at FS temperatures (Hohenberg et al., 1994; Thijssen

et al., 1998). For flies, we prefer to make a paste of dry baker’s yeast with 10%

methanol or buVer. Mix the liquid and dry components in roughly equal volumes,

then adjust the final consistency so that the paste is as thick as possible but can still

be used to pack around the fly embryos. Some workers like to use the yeast paste

that is used on fly egg-laying plates or trays. For worms, we often use the E. coli

‘‘lawn’’ that is used as food on worm plates. Worm laboratories will have lots of

dishes with E. coli, and if you can find plates that are a little older, that is, when the

E. coli has a thicker consistency, those will work well for filling the spaces around

worms. Thus for both flies and worms, we are using materials that normally

surround these organisms in their laboratory growth habitat.

Another choice for flies and worms is 20% bovine serum albumin (BSA) made

up in phophate-buVered saline (PBS) or M-9 buVer (Sulston and Hodgkin, 1988)

for worms. This is used for correlative LM/EM work on single worm embryos

( Chapter 4 by Mu ller -Reichert et al ., this volume ) and routi nely gives goo d freez ing.

It has also been shown to be a good cryoprotectant for HPF of tissue culture cells


(Reipert et al., 2004a), murine skin cells (Reipert et al., 2004b), and a variety of

other organisms (McDonald et al., 2006 and unpublished results reported to

the author).

Intracellular cryoprotectants are less desirable because they have the potential

to interfere with cell physiology, and they may change ultrastructure. Space does

not permit an extensive discussion of this issue here, but see Gilkey and Staehelin

(1986) for more details. Fortunately, for the organisms that are the subject of this

chapter, the choice of cryoprotectant is not usually a problem. Yeast, for example,

are simply concentrated to a thick pastelike consistency in their normal growth

medium. Drosophila embryos and Caenorhabditis elegans worms are surrounded

by impermeable barriers, such as vitelline envelopes or cuticle, so the material

surrounding them for the 30 sec or so before freezing appears to have little or no

eVect on internal ultrastructure.

E. Tools Useful for Most HPF Specimen Loading Operations

1. Fine forceps for picking up specimen cups and other small items.

I particularly like the long fine forceps such as Cat. No. 72919-SS from

EMS. For the BAL-TEC machine, an additional pair of bent tip forceps

such as Cat. No. 72703-D from EMS or other EM vendors are useful for

picking up the specimen cups.

2. Paper points (Ted Pella, Redding, CA, Cat. No. 115–18) or some other

wicking material.

3. Fine needles.

4. Fine paint brushes. These are useful for transferring delicate samples, or

for loading minute amounts of cryoprotectants into the specimen carriers.

III. General Rules for Loading Samples for HPF

Sample loading is the most critical part of the HPF process!!! This point cannot

be overemphasized. Hans Moor, the inventor of the technique, said as much in his

comprehensive article on the theory and practice of HPF (Moor, 1987). If you

want good results from HPF you must be aware of this fact and act accordingly.

The machines are robustly designed and work the same way every time, freezing

event after freezing event. So if you are not getting good results it is unlikely to be

the fault of the machine. To remedy the situation, you have to check what you are

doing before the freezing (specimen loading) or after (e.g., FS). In most cases, it is

mistakes with specimen loading that are the problem. We will deal with FS issues

in a later section. Here, we will give some general rules to consider and to use when

you are freezing. If you use them carefully, you will be rewarded with better yields

of well-frozen samples.

32 Kent McDonald

A. Work Only with Healthy, Unstressed Cells

Cells/tissues must be in optimal physiological condition. Yeast cells in liquid

culture, for example, should be in early to mid log phase growth, if at all possible.

They should be at the right temperature, in the most physiologically appropriate

medium, and in most cases should be on a shaker. Cells should not be concen-

trated and allowed to sit on ice. Worms are best picked directly from food

plates and not washed oV with M-9 buVer or otherwise be surrounded by liquid

(Section III.D). They should be well fed and be at optimal temperature and

humidity for good growth and development. The same goes for flies on egg-

layi ng plate s. As discus sed in detail by Hess (Chapt er 3, this volume ), there are

other stresses on organisms, including genetic manipulations, that can adversely

aVect the preservation of ultrastructure.

B. Work Quickly

For all types of preparations, one should go from living to frozen cells in

less than a minute. While this is not always possible with some preparations,

it is easy for others. If concentration procedures are necessary, they should

be done as quickly as possible and be followed by freezing with a minimum lag.

One must always consider the kinetics of the process under study and realize

that the interval between the time of departure from optimum conditions and

the time of freezing is available for the creation of artifacts that can surpass the

damage done by chemical fixation. A well-frozen dead or sick cell is not worthy of

study.

C. Do Not Let Your Cells/Tissues Dry Out

The volume of the specimen carriers of nearly all shapes and sizes is less than a

microliter. This volume will dry out very fast, especially if the humidity around the

sample is low. Try working in a moist chamber or otherwise keeping your material

moist over the time you are taking samples for freezing. When loading yeast into a

cup-shaped specimen carrier, put a top on the cup (BAL-TEC and Wohlwend

machines), or insert the sample into the machine (EM PACT2 with RTS) as soon

as the cup is filled. As an experiment, load some yeast into a cup then watch under

a dissecting microscope to see how long it takes for the surface to change from a

shiny (wet) to a matte finish (dry). If your yeast are highly concentrated to begin

with, this will take less than a minute.

When working with worms or flies in E. coli or yeast paste, have a vial of M-9

buVer, appropriate fly buVer, PBS, or 20% BSA close at hand and add a little with

a fine-tipped paint brush if the specimens in the cup begin to dry out. The worms

should be moving in the cup (unless they are paralyzed mutants) when you load

them into the HPF machine. For fly embryos in yeast paste, use a little 8%

methanol solution in water if you need to moisten the paste.


D. Avoid Surrounding Your Cells/Organisms with Aqueous Media

This is a common mistake that can lead to poor results, even when you do

everything else correctly. The importance of this issue has to do with the path

of heat transfer from the deep interior of your cells or tissues, through the extra-

cellular medium, and into the metal of the specimen holder. The deeper the

specimen cup, the more this is a problem. If there is a significant amount of free

water around the cells, this will form ice during the freezing process and block or

retard the transfer of heat from your cells. This is the rationale for using external

cryoprotectants that will bind or replace the free water outside the cells.

Good practical advice regarding thesemodel systems is to: (1)make sure the yeast

are more pastelike than liquid when freezing, (2) avoid concentrating worms from a

plate by rinsing them into a tube with M-9. If that has to be done for some reason,

try to transfer some worms back to an agar plate and let the agar absorb the excess

liquid before loading the worms into a specimen cup, and (3) if fly embryos are

dechorionated in liquid solutions, transfer them to a Nitex screen or some other

filter to let the excess liquid drain oV before loading them into the specimen cup.

E. Avoid Mechanical Damage

Sloppy cutting of tissues or the overfilling of specimen cups can damage your cells.

Use biopsy needles where feasible, always use the sharpest scalpels and razor blades

for bulk cutting. Disposable dermatology biopsy punches in 1.0-, 1.5-, and 2.0-mm

diameters are useful for leaves and other hard materials. It is better to have too little

material and fill in the spaceswith a suitable cryoprotectant thanhave sample sticking

out the top of the carrier that will get crushed when the top piece is put on.

F. Fill the Carrier Correctly

Use a filler that is compatible with the cells being frozen, that has good heat

transfer properties, and do not overfill or underfill the carrier. The ideal fill is a

very slight overfill that will contact the top piece over the cup and ensure that no

air is trapped in the cup as the top comes down. Air is not only a barrier to heat

transfer but will collapse under pressure and perhaps cause shear that will damage

your tissue. In either case the result will be bad freezing. You can tell with the

BAL-TEC and Wohlwend systems if there has been air in the cups because they

will collapse inward and leave a concave depression on the outer sides. The best

way to learn to recognize this phenomenon is to deliberately underfill a cup and

observe the subsequent shape of the cup walls.

G. Use the Smallest Volume of Sample You Can

This may be the single most important factor to getting high yields of well-frozen

cells. All the HPF machines give you a choice of specimen carriers with diVerent

34 Kent McDonald

depths or volumes. The important dimension is the depth, for example, of the

cup-shaped carriers. If you are freezing a cell suspension such as yeast, always use

the shallowest depth available, usually 100 mm.With the BAL-TEC andWohlwend

machines you can even use slot grids to create variable-depth spacers between two

flat-sided specimen carriers (McDonald et al., 2006). For worms, the 100-mm-deep

carriers also work well because they are deeper than the thickness of adult worms.

Fly embryos, on the other hand, are closer to 200 mm in width, so 200-mm-deep

carriers must be used. For cells in suspension, dipping a mesh grid into the suspen-

sion and sandwiching it between two flat-sided specimen cups or very shallow

custom-made cups (Muller et al., 1980; Murk et al., 2003) is another option.

IV. Methods for Specific Organisms

A. Yeasts (Saccharomyces cerevisiae and Schizosaccharomyces pombe)

1. Background

Yeast has long been associated with developing EM cryomethods, going back to

the early days of freeze-fracture studies (Moor, 1967; Moor and Muhlethaler,

1963). Using plunge freezing methods, both S. pombe (Tanaka and Kanbe, 1986)

and S. cerevisiae (Baba and Osumi, 1987) were shown to have vastly improved

ultrastructural preservation after FS and thin sectioning. Despite these early

observations, studies of yeast ultrastructure have continued to use conventional

room temperature methods of specimen preparation, and even in a relatively

recent publication (Heiman and Walter, 2000), a modified potassium permanga-

nate method was employed. The idea that freezing followed by FS would improve

visualization of fungal cell ultrastructure in thin sections goes back to the

pioneering work of Howard and Aist (1979). Mendgen and coworkers began

using HPF as the freezing method for a number of diVerent fungi (Knauf and

Mendgen, 1988; Knauf et al., 1989; Welter et al., 1988). The work by Ding et al.

(1993) on S. pombe and Winey et al. (1995) on S. cerevisiae were early eVorts usingHPF to study yeasts where the focus was on the biology rather than the technique.

By now, there are dozens of papers using HPF on yeasts to address a wide variety

of biological questions.

2. Materials

a. Vacuum source (pump or house line)

b. Vacuum filtration apparatus, typically a 15-ml Millipore setup

c. 0.45-mm pore size polycarbonate filters, 25-mm diameter

d. Toothpicks or other implement to scrape cells from filter

e. Syringe needle bent at 45 � near the tip


f. Yeast cells in liquid culture, growing in early tomid log phase (OD¼ 0.2–0.4),

you will need 5–10 ml for every freezing event, so adjust the total volume to

your needs

g. 2% agar made up with YPD (see Difco Manual or search for ‘‘Yeast Plates’’

on a web browser) and poured into 5- to 8-cm-plastic Petri dishes

3. Methods

a. Set up filtration apparatus.

b. Pour 5–15 ml cells into filtration column.

c. Apply suction to pull down cells onto filter, using care not to let them get too

dry.

d. Remove column from apparatus, place filter with cells on agar plate.

e. Scrape cells from filter with toothpick tool.

f. Fill well of specimen carrier with cells.

g. Remove excess cells from carrier with syringe needle tool by scraping over

the surface. Instead of moving from side to side, put needle across the center

of the cup and move to one side. Repeat moving to the opposite side but

leave a small ridge of yeast across the middle to ensure that the cup is very

slightly overfilled.

h. Freeze.

4. Further Reading

For more detailed discussions of methods for preserving yeast for EM see

Giddings et al. (2001) and McDonald and Muller-Reichert (2002). In the latter

reference, there are also methods for users who do not have access to HPF

technology.

B. Nematodes (C. elegans)

1. Background

There is a long history of EM associated with C. elegans because the organism

was developed as a model system with EM in mind (Brenner, 1973). Early studies

used serial sections to map out the structure of the anterior sensory anatomy

(Ward et al., 1975), of the ventral nerve cord (White et al., 1976), and the pharynx

(Albertson and Thomson, 1976). These, and many other C. elegans EM references

can be found on the C. elegans WWW Server (http://elegans.swmed.edu/) using

‘‘electron microscopy’’ as a search term in the Literature Search link. While the

quality of specimen preservation was suYcient for tracing cell lineages as well as

other descriptive work, it would not be good enough to meet today’s standards for

preservation at the molecular level.

http://elegans.swmed.edu/

36 Kent McDonald

2. Materials

a. A tool for picking worms oV a plate of E. coli

b. E. coli for use with the picking tool, older plates with thicker, ‘‘stickier’’

bacteria work best

c. Alcohol lamp for flame sterilizing the worm-picking tool

d. Specimen carriers and accessories for the appropriate HPF machine:

1. EM PACT machines. Typically we use the100-mm-deep membrane

carriers

2. BAL-TEC machines. For this instrument you will need:

–Type B specimen carriers (flat on one side)

–TEM slot grids (EMS Maxtaform Cu-Rh 1 � 2 mm2, Cat. No. M2010-

CR)

–1-Hexadecene

–60-mm plastic Petri dish fitted with filter paper. Saturate the filter paper

with the hexadecene

–A number 0 red sable paint brush

–A number 11 scalpel blade

–Fine needles

e. M-9 worm buVer solution

f. 20% BSA in M-9 buVer

3. Methods

a. EM PACT2 HPF machine:

1. Place a 100-mm membrane carrier in the loading device.

2. Slightly overfill the cup of the membrane carrier with 20% BSA solution

in M-9.

3. Using the worm-picking tool loaded with E. coli, pick 15–20 worms oV the

worm plate. Touch these to the BSA in the cup and allow the worms to

swim oV into the specimen carrier or push them oV the pick with a fine

needle.

4. Use the paper points or other wicking material to lower the level of liquid

so that there is only a slight excess of material in the cup.

5. Load and freeze in the EM PACT machine.

b. BAL-TEC HPF machine:

1. Place a number of Type B specimen cups (flat side down) and slot grids on

the 1-hexadecene saturated filter paper.

2. Blot oV excess 1-hexadecene on the specimen cup on dry filter paper and

put into the tip of the specimen cup holder flat side up.

3. Put the slot grid (1-hexadecene side down) on top of the specimen cup. Do

not blot excess oV the slot grid.


4. Using the paint brush, put a small amount of M-9 buVer in the slot. It will

bead up and should occupy only about half the slot area.

5. Pick 20–25 worms with the worm pick or E. coli paste.

6. Use the tip of the scalpel blade to scrape the worms and E. coli into the

M-9 droplet in the slot grid.

7. Use fine needles to mix the worms and E. coli. Allow the solution to

thicken and the volume to reduce. Do not let the paste dry out completely.

The worms should continue to move throughout this process. If the paste

becomes too dry, you can reconstitute it with M-9 from the paint brush.

This procedure takes a little practice to get right, and practicing ahead of

time will reduce frustration and promote the necessary skills.

8. When the area of the slot grid is only slightly overfilled with worms or

E. coli solution, take another specimen carrier from the 1-hexadecene pad

and place it flat side down on top of the worms.

9. Close the tip of the specimen loader and freeze in the BAL-TEC or

Wohlwend HPF machine.

4. Further Reading

For more detailed information about cryomethods for preserving nematodes

for EM see Muller-Reichert et al. (2003a).

C. Drosophila Embryos

1. Background

EM studies of Drosophila go back to the early studies of Lowman (1958),

Mahowald (1962, 1963), and King (1960; King et al., 1966; Koch and King,

1966) among others. More than yeast or worms, EM studies of Drosophila have

been published regularly until the present day, although few researchers have

chosen to use rapid freezing and FS methods. Drosophila was among the first

organisms to be studied by HPF (Muller and Moor, 1984) and has been the object

of HPF study in some more recent publications (Bardsley et al., 1993; McDonald

and Morphew, 1993; Schulte et al., 2003).

2. Materials

1. Embryos on food plates or harvested and dechorionated. Do not store

embryos in liquid solutions. Keeping them moist on agar or food plates is

the best solution.

2. Yeast paste. Many fly embryo plates are seeded with yeast paste as food for

the flies. You can use this as a filler or make up a separate paste. Making up a

separate paste using 8% methanol as a wetting agent for dry yeast will give a

better yield of well-frozen embryos and will not aVect them adversely.

3. Number 0 red sable paint brush.

4. 8% methanol solution in an Eppendorf tube.

38 Kent McDonald

3. Methods

1. Working on an agar or food plate, use the paint brush to bring together 20–

25 embryos in a small clump. If the embryos are covered with a liquid layer,

wick that oV, or let them sit a while until the surface appears dry.

2. Take a small amount (practice will tell you how much) of yeast paste on the

tip of the paint brush and mix it with the clump of embryos until all are

thoroughly coated with yeast.

3. Place a 200-mm-deep cup-shaped specimen carrier (either BAL-TEC or

Leica) on the surface next to the coated embryos and transfer the yeast–

embryo mixture into the cup so that it is full but not overfull. The ideal fill is

just to the level of the top edge of the cup. If it is too full, the cells will be

compressed when the top is put on. If it is not full enough there will be an air

space and heat transfer will be seriously impeded.

4. Transfer the specimen carrier to the HPF specimen loader and freeze.

4. Non-embryo Drosophila Tissues

Processing adult or larval fly tissues is diYcult because of the cuticle or other

hard materials that cover the cells. These materials probably do not interfere

much with the freezing process itself, but rather the subsequent FS steps where

organic solvents and fixatives still have to cross the diVusion barrier and replace

the cell water inside the tissues. Another problem is that the adult and larval

tissues are generally too large to fit into even the largest HPF specimen carriers.

Therefore, it is necessary to dissect the desired parts away from the intact organ-

ism before freezing and this may also have negative consequences for cell fine

structure.

5. Further Reading

For more information and details about HPF and subsequent EM processing of

Drosphila tissues see McDonald (1994) and McDonald et al. (2000).

V. Postfreezing Processing

Freezing specimens is only the first of several steps between living tissue and

the electron microscope. Care must be taken at each step to ensure that high-

resolution structural information is not lost. Common options after HPF

include: freeze-fracturing and metal coating, vitreous cryosectioning, and FS and

embedding in resin. In this chapter, we will only concern ourselves with FS

and embedding because this is by far the most common way to prepare samples

after HPF.


A. Freeze-Substitution

A literature search for ‘‘freeze-substitution’’ will turn up a large number of

examples, some going back 40 years or more. The idea of using low-temperature

dehydration and fixation methods for EM is usually credited to Fernandez-Moran

(1960), although the idea of FS was well known in light microscope cytology

(reviewed in Feder and Sidman, 1958). Freeze-drying for EM is a much older

technique and was used in some very early EM observations of biological material

(Richards et al., 1943). Van Harreveld and Crowell (1964) are often cited for using

osmium-acetone as an EM FS fixative. For the history-minded, the book by

Echlin (1992) has references to the earlier literature of cryomethods. Other papers

to consult for general principles and applications of FS are by Steinbrecht and

Muller (1987), Hippe-Sanwald (1993), and Nicolas and Bassot (1993).

FS means to substitute an organic solvent, usually acetone, for the cell water at a

low temperature, typically �78 to �90 �C. Other solvents such as methanol or

ethanol are sometimes used, but the convention for most people today is to use

acetone. Unless one is doing low-temperature embedding with Lowicryls, it is

usually necessary to add a fixative to the FS solvent. Osmium tetroxide is the most

common fixative, but paraformaldehyde and/or glutaraldehyde can also be used.

Unlike conventional processing, the fixation takes place after or perhaps during the

dehydration steps. At �90 �C, acetone will dissolve the cell water over a period of

hours to days, but the fixatives are not thought to be very reactive at this temperature

(Humbel and Muller, 1986). However, the fixative becomes distributed throughout

the entire cell or tissue, and when the temperature is permissive for fixation, the

fixative is ‘‘in place’’ and does not need to diVuse through relatively great distances,

as it does for conventional processing based on diVusion of reagents. The tempera-

ture where fixation begins is not well known, although the studies of Humbel and

Muller (1986) suggest that glutaraldehyde is active at about �50 �C and osmium

tetroxide at �30�C or so. Another important reference that deals specifically with

fixation of chromatin at low temperatures is Horowitz et al. (1990). In this study,

cross-linking of chromatin with 3% glutaraldehyde in acetone begins at �45�C.

1. Which FS Method to Choose for Your Cells?

FS is a process that is poorly understood, and there has been little systematic

research to probe basic mechanisms. Consequently, there are many diVerent meth-

ods published in the literature. The majority of these methods appear to work, so it

seems to be a process with a largemargin of error. The practical problem for the new

user, however, is which method to choose. One suggestion is to search the literature

to see if there are good results on the cell and tissue type that you are working on.

This seems like a good idea, but be aware that diVerences in the reagent chemistry

from one laboratory to another or subtle procedures not published in the original

paper can mean that the method will not give the same good result. Another

approach is to select a general protocol that seems to work for a wide variety of

40 Kent McDonald

tissues and use that as your star ting point. This is the approa ch we have taken in our

laborat ory. By always star ting wi th the same method, we see the variations that are

due to the chemi stry of the cells themselves . If we do not like the resul ts, we can

begin to change some of the varia bles in a systemat ic way to see how we can improve

them. Getting a good FS resul t may requ ire a lot of problem solvi ng and experi -

menta tion wi th the steps in the process . The steps include not only those of tim e,

tempe ratur e, an d chemi stry of the FS process itself, but there are a numb er of

varia bles dur ing HPF that must also be taken into accoun t.

2. Variables During Freezing

Most of these points such as workin g with optimall y healt hy cell s, choosing the

right filler , and so on ha ve alrea dy been discussed in Section III . In addition, there

are some issues that we can menti on here that might help illustrate the subtle

nature of the facto rs that c an aV ect freez ing.

a. The Genotype of the Cells Being FrozenIt is our experience that some mutants do not look as good as wild type even

when they are processed in exactly the same way. As discussed in detail by Hess

( Chapter 3, this vo lume), this is only one of severa l ways that organis ms become

altered in ways that will adversely aVect the preservation of ultrastructure.

b. The Way the Cells Are Grown Can Make a Big DiVerenceFor example, we have heard anecdotally from a yeast researcher that diVerences

in the carbon source in a yeast medium (e.g., acetate vs glucose) can make a

big diVerence in the visualization of mitochondrial membranes. Another plant

researcher claims that using mannitol instead of sorbitol in the buVer will make

the diVerence in the final appearance of plant protoplast cells in the microscope

(Lonsdale et al., 1999).

3. Variables During the FS Run

a. How the FS Cocktail Is Made and StoredWe make batches of fixative in cryotubes and freeze them for use later. They are

stored in liquid nitrogen. This has the advantage that when you are done with

HPF, you can put your sample directly into the fixative and start an FS or you can

store the sample in the cryovial/fixative until you are ready to do the FS. We have

published a detailed recipe for preparing FS fixatives in several earlier papers

(McDonald, 1999; McDonald and Muller-Reichert, 2002; McDonald et al., 2000);

this method works well for us. The important thing is not to leave the fixative

mixture at room temperature any longer than necessary. Also, when freezing the

fixative in cryovials, make sure it is upright when frozen so the liquid does not get

into the threads of the cryovial cap. If this does happen, it makes it diYcult to

unscrew the cap when loading samples under liquid nitrogen.

We always make up our FS solutions from small bottles of just-opened acetone,

as opposed to acetone over molecular sieves. We believe, as has been mentioned in


the literature (Steinbrecht and Muller, 1987), that acetone extracts something

from molecular sieves that can react with osmium tetroxide. Because uranyl

acetate is not readily soluble in acetone, we make up a 5% stock solution in

methanol and dilute that with acetone to get a final concentration of 0.1%.

b. The Chemistry of the FS MediumThis includes choice of solvent(s), choice of fixative and concentration, and

whether or not additives are added to the fixative–solvent cocktail. For example,

we have chosen 1% osmium tetroxide plus 0.1% uranyl acetate in acetone as our

standard FS cocktail for starting morphological studies. Acetone substitutes the

water at a much slower rate than methanol, and this turns out to be a desirable

property. If the substitution goes too rapidly, as it can do with methanol, the

ultrastructure may suVer (McDonald, 1994; Steinbrecht, 1993). It is hard to

generalize about this because every tissue is diVerent and studies using methanol

have produced good preservation of cell fine structure (Muller et al., 1980).

Increasing the concentration of osmium tetroxide may add contrast to some

samples, but we find that 1% works well for the model organisms covered in this

chapter. Additives to the solvent–fixative cocktail can make a diVerence to the

final appearance of cells. We always include about 0.1–0.2% uranyl acetate with

both our osmium and glutaraldehyde FS cocktails. We find that it adds contrast to

membranes and seems to prevent destruction of ultrastructure that can occur if the

cells are in osmium/acetone alone at room temperature for more than an hour.

With the uranyl acetate added, one can leave the cells at room temperature for

longer periods and the solution will not turn black as quickly, another bit of

evidence suggesting that uranium could be aVecting the osmium interactions with

biological material. There are other ways to add membrane contrast, and one of

the more intriguing is to add water to the FS cocktail (Walther and Ziegler, 2002).

This approach is covered in more detai l in Chapter 3 by Hess, this volume .

c. Time and TemperatureMost FS protocols start with the samples held at a temperature of �78 to�90�C

for some amount of time, then warming of the sample over another prescribed time

period. The initial period varies from a few hours to a week, and the warm-up time

can be similarly varied. There are few systematic studies of these variables, but

Steinbrecht did some experimentation on FS times for moth antennae some years

ago (Steinbrecht, 1982; Steinbrecht and Muller, 1987). These showed that if the

time at �80�C were long enough (7 days), the sample could be warmed up in 1 min

without significant ice damage. Likewise, time at �80�C could be reduced to 5 min

if the warm-up time were extended over 6 h. The problem with this experiment, and

all similar experiments, is that they may only be true for the specimens under study.

Given the extreme heterogeneity of biological tissues, generalizations are not really

possible. The approach taken by most investigators is to be conservative, leaving

tissues at �80 to �90�C for 8–72 h, and using warm-up times from 6 to 24 h.

In our laboratory, we tend to use a protocol that varies the period at �90 �C but

leaves the warm-up period constant. For example, if we are doing an FS at the

42 Kent McDonald

beginning of the week, we hold at �90 �C for as little as 5 h. If we are freeze-

substituting over a weekend, it might hold for as long as 36 h. We chose a warm-up

rate of 5 �C/h, andwe halt the warm-up at�25 �C for 12 h, because we found that we

getmoremembrane contrast that way (SectionV.D). Then we continue warm-up to

0 �C where we hold the samples until we are ready for further processing. At that

point, samples are warmed rapidly to room temperature, rinsed three times over

15 min in pure acetone and infiltrated with resin for embedding.

B. Embedding

The choice of embedding resin is another variable that can influence the final

appearance of your high-pressure frozen, freeze-substituted sample.We generally use

an Epon-Araldite formulation of medium hardness (Mollenhauer, 1964) because of

its excellent sectioning and beam stability properties. For yeast, we find that

a mixture of Epon and Spurr’s resins (McDonald and Muller-Reichert, 2002)

works very well for revealing cytoplasmic details (Fig. 1). Generally, we do not use

Spurr’s resin for anything because it will extract membranes and other cytoplasmic

components (Hess, 2003 and Chapter 3, this volume). However, because the

cytoplasm of yeast is so extremely dense compared to most cells, perhaps a little

extraction is useful in this particular case.

C. FS and Embedding for Tomography

For samples of resin-embedded material analyzed by ET, the contrast comes

mostly from electron-dense molecules used in FS (osmium and uranyl acetate) and/

or poststaining (uranyl acetate and lead citrate). In some cells like yeast and in some

structures, like the spindle pole body, the electron density can become so high after

FS and poststaining that it is diYcult to visualize organelle substructure. A strategy

for making medium-density images of well-fixed samples is to freeze-substitute

them in relatively high concentrations of glutaraldehyde instead of osmium tetrox-

ide, and use only the poststaining solutions to generate contrast (McDonald and

Muller-Reichert, 2002; O’Toole et al., 2002). Embedding in a methacrylate resin

such as Lowicryl HM20 also seems to help. For worm tissues, we find that our

standard FS protocols work well for tomographic studies (O’Toole et al., 2003).

So far, we have not done any tomographic studies of Drosophila, but based on

our experience with routine sectioning studies, we expect that standard FSmethods

will work well for most tomographic analyses.

D. FS and Embedding for Immunocytochemistry

For immunocytochemical studies our standard fixative is 0.2% glutaraldehyde

plus 0.1% uranyl acetate in acetone. Again, we find that the addition of uranyl

acetate has a beneficial eVect on membrane contrast, but does not seem to

interfere with most antigen–antibody reactions, at least not at this concentration.

For yeast, we sometimes use another fixative combination made up of 0.1%

Fig. 1 S. cerevisiae visualized in a 60-nm section after HPF-FS and embedment in Epon-Spurr’s resin.

Endoplasmic reticulum (er) extends from the nuclear envelope toward the cell cortex. Three cisternae of

a Golgi apparatus (g) are visible as are the fine projections called fimbriae (f) at the surface of the cell

wall. The cell wall is broken (arrow, upper left), as sometimes happens with high-pressure frozen yeast

cells. Scale bar ¼ 1.0 mm.


glutaraldehyde, 0.5% uranyl acetate, and 0.01% osmium tetroxide in acetone.

This small amount of osmium is suYcient to add membrane contrast, but it still

will permit immunolabeling, at least with some antibodies (McDonald and

Muller-Reichert, 2002; Muller-Reichert et al., 2003b).

The embedding medium that we prefer for immunolabeling work is LR White

resin, the hard formulation.We order our LRWhite from Ted Pella, Inc. (Redding,

CA) because it is shipped with the accelerator separate from the resin. Accelerator

is only added when the resin is made up. Some other vendors have the accelerator in

the resin at shipping, and you do not know how long it has been on their shelves

before that time. We find that LR White is adequate for worms (Fig. 2; Kosinski

et al., 2005) as well as flies (Bardsley et al., 1993; McDonald et al., 2000) and

yeast (Cid et al., 2001). Lowicryl is probably a superior resin for preserving cell

morphology, but when you are working with small, colorless samples in a �50 �Cenvironment, it is sometimes diYcult to see what you are doing. The exception to

that problem is yeast, which tend to form a firm disk after FS that usually releases

from the HPF specimen carrier. These disks can be transferred to gelatin capsules

in the FS device and polymerized at low temperatures. We recommend using

Lowicryl HM20 resin and polymerizing with UV irradiation at �50 �C.

Fig. 2 Part of an amoeboid C. elegans sperm following HPF, FS in 0.2% glutaraldehye plus 0.1%

uranyl acetate in acetone, and embedding in LR White. 10-nm gold particles (arrows) show the

distribution of major sperm protein (MSP). Antibody against MSP courtesy of David Greenstein,

Vanderbilt University Medical School. Scale bar ¼ 200 nm.

44 Kent McDonald

For worms and fly embryos we prefer to embed in thin layers of resin, so we can

screen the organisms in the light microscope prior to sectioning. For this ap-

proach, it is necessary to use LR White resin and work at room temperature.

Although this method was originally developed in conjunction with microwave

polymerization (Lonsdale et al., 2001), we now use oven polymerization more

routinely. We construct the thin layer wells as shown in Lonsdale et al. (2001) but

put them in a sealed Rubbermaid storage container that has been flooded with

nitrogen gas to exclude oxygen. This is then put into the oven overnight, and the

next day the chamber is flooded with N2 gas again and put back into the oven for

one more day to ensure complete polymerization. After screening with the light

microscope, individual worms/fly embryos are cut out, remounted, and precisely

oriented for sectioning. For yeast, we almost always embed small pieces of yeast

pellets in LRWhite in flat-bottomed capsules (Ted Pella Cat. No. 133P) polymerized

in the microwave (Cid et al., 2001).


E. A Case Study of Adjusting FS Variables to Improve Visualization

When you look at a sample in the microscope and find that you do not like the

way certain features look, you should consider how to change the FS methods to

improve the situation. As an example, we present here the changes that evolved in

our work to get improved visualization of the yeast nuclear envelope (Fig. 3).

In our earliest attempts at FS with yeast, we used 2% osmium tetroxide in acetone

Fig. 3 Changes in the appearance of the nuclear envelope of S. cerevisiae cells as a function of changes

in the FS protocol. Beginning with 2% osmium tetroxide in acetone (A) there is no clear density in

the nuclear envelope; it is detectable mostly because of the absence of staining in the lumen between

the double membranes. (B) The addition of 0.1% uranyl acetate to the osmium acetone mixture creates

faint dark lines in the areas where we expect to see membrane. By using the same FS cocktail as in

(B), but holding the FS temperature at �25�C for 12 h during the warm-up period, the membrane

contrast is increased even further (D). If the cells are fixed ‘‘hard’’ with 2% glutaraldehyde during

FS, then some membrane contrast can be generated from uranium and lead poststains alone (C). Scale

bars ¼ 200 nm.

46 Kent McDonald

as the FS fixative cocktail, because this was considered standard at the time. The

results were okay, but a number of people were concerned about the lack of

contrast in the membranes (Fig. 3A). Sometime later, Mary Morphew of the

Boulder Laboratory added uranyl acetate to the FS mix, and this seemed to

present a more familiar membrane image (Fig. 3B). Then, as a result of experi-

ments on other organisms with low membrane contrast, we came up with the idea

of holding the FS warm up at �25 �C overnight to add even more membrane

contrast (Fig. 3D). This is now our current standard starting FS protocol for all

new organisms. In general, if you want more contrast, you can incubate samples in

this FS cocktail for variable periods, and the longer you hold it the more contrast

you will get, up to the point when the reaction saturates. However, it is also

possible to visualize membranes without using osmium at all. In Fig. 3C, the cells

have been prepared for tomographic analysis using 2% glutaraldehyde plus 0.1%

uranyl acetate as the FS fixative. The contrast here comes mostly from the

poststaining of sections in uranyl acetate and lead citrate. These cells were embed-

ded in Lowicryl HM20, which also seems to help the contrast.

We must mention that the desire to see membranes rendered as dark lines is an

interesting paradox. This view does not match what one would predict from the

dist ribution of lipid compo nents in membr anes, but it is one that has evolved as a

consistent artifact of conventional EM preparations; most people are more com-

fort able when they see membr anes this way (see also Chapter 3 by Hess, this

volume ). We should also say that membrane contrast in freeze-substituted samples

is highly variable from organism to organism and even within cells. It probably

reflects the diVerences in the membrane chemistry as well as the FS protocol.

In general, membranes in well-frozen samples will look less contrasty than the

same membranes prepared by conventional methods. For the organisms that are

the focus of this chapter, however, the yeast are the most problematic. Worms

(Figs. 4 and 5) and fly tissues generally give reasonable to good membrane

contrast with the methods we recommend here. For an alternative approach to

the FS of yeast cell membr anes, see Walther and Ziegle r (2002) . We have tried this

method in our laboratory, and the images of yeast membranes are excellent.

F. Sectioning

Material prepared by rapid freezing and FS is more dense than that prepared by

conventional methods because there is less extraction of the cytoplasm. For yeast

cells, especially, it is a good idea to cut sections thinner than you might normally

do. We typically section at 40–60 nm when studying yeast cells, and these sections

look sharper than those cut thicker (see Fig. 4 in McDonald and Muller-Reichert,

2002). For worms and flies, we vary the thickness between 60 and 100 nm,

depending on the cytoplasmic features we are interested in.

The time and type of poststaining on the sections can also be important to how

the cells look in the microscope. With the procedures for FS and embedding that

we normally use, we find that poststaining in 2% uranyl acetate made in 70%

Fig. 4 A cross section through microvilli on the gut epithelial cells of C. elegans. Individual actin

filaments (a) are visible in the cores of the microvilli, and molecules coating both the inner (ic) and outer

(oc) sides of the microvillar membrane are evident. Scale bar ¼ 200 nm.


methanol for 5 min, followed by 3 min in Reynolds (1963) lead citrate gives good

results. Longer staining times can actually reduce the contrast on yeast sections

because while the overall staining density is greater, the contrast goes down

because the range of tones is less. For sections of fly or worm material, we may

go as long as 10 min in methanolic uranyl acetate and 5 min in lead citrate. Staining

sections with 1% aqueous tannic acid for 3 min (then rinse well) prior to staining

them in uranyl acetate and lead citrate can also be very useful if you are interested

in cell walls, or certain classes of membranes and endocytic vesicles (Fig. 6).

G. Microscopy and Evaluation of Results

Looking at sections or micrographs of material prepared by rapid freezing and

freeze-substitution fixation (RF-FSF) techniques will present some unfamiliar

images to even experienced EM users if they are only familiar with conventional

fixation (CF) methods. The cytoplasm of RF-FSF material will be denser and

have less contrast than CF material (Figs. 7 and 8). This is because the cytoplasm

is less extracted and there is less collapse and concentration of proteins and

other structures (Kellenberger, 1987). One of the features of a cell that is easiest

Fig. 5 Details in a cross section of the head region of C. elegans. An E. coli cell (e) can be seen on the

outside of the cuticle (c). A cross section through the body muscle (m) is just inside the cuticle, and

concentric membranes (me) and amphid cilia (a) are seen further in. Scale bar ¼ 0.5 mm.

48 Kent McDonald

to assess regarding fixation quality is the shape of the nucleus. We know from LM

of living cells that most nuclei are round or ovoid; therefore, a section through

the nucleus should look like a circle or an ellipse (Fig. 7A). If only part of the

nucleus is visible, the nuclear envelope should form a smooth curve. If the nucleus

has an irregular outline (Figs. 7B and 8B), then it has most likely suVered from

distortions during specimen preparation. Of course, some nuclei, such as those

occurring in certain classes of leucocytes, are lobed naturally but these are usually

characterized by deep invaginations in the nuclear envelope, as opposed to the

kinds of patterns shown in Figs. 7B and 8B. Other organelle membranes can

show similar distortions due to collapse during specimen preparation. Space

Fig. 6 Use of special poststain procedures on sections to enhance certain cytoplasmic details. A method

developed by Susan Hamamoto at the University of California, Berkeley uses a 3-min incubation in 1%

aqueous tannic acid, followed by extensive rinsing in dH2O before conventional poststaining with uranyl

acetate and lead citrate.Golgimembrane cisternae (g inA) aswell as certain vesicles located in the emerging

bud (arrows in B) react with the tannic acid plus the other poststains and become darkly stained.

Reproduced from McDonald and Muller-Reichert (2002), courtesy of Academic Press, San Diego. Scale

bar in (A) ¼ 0.5 mm; Scale bar in (B) ¼ 0.25 mm.


does not permit showing all the diVerences that one encounters between cells

prepared by cryomethods and CF, but a study of the literature should help make

the diVerences clear. References to hundreds of papers using HPF and FS methods

can be found on theUniversity of California ElectronMicroscope LaboratoryWeb

site (http://em-lab.berkeley.edu/EML/protocols/hpflit.php), or one can search bib-

liographic databases for similar items. It should be pointed out, however, that not

all papers claiming to get improved results by cryomethods actually do so. Freezing

brings its own set of specimen preparation artifacts, mostly those that result from

ice crystal damage.

H. Artifacts of HPF

Alterations of cell ultrastructure due to poor freezing are well known and not

restricted to HPF work. In fact, for large pieces of tissue, they are less common in

HPF than with other cryoimmobilization methods. These artifacts are usually

referred to as ‘‘ice damage’’ and occur when the rate of freezing is too slow to

prevent rearrangement of cellular water molecules into ice crystals. Because the

rate of heat transfer is highest at the surface that is in contact with the cooling

http://em-lab.berkeley.edu/EML/protocols/hpflit.php

Fig. 7 Comparison of Drosophila embryo cells processed by high-pressure freezing and freeze-

substitution (HPF-FS) as seen in (A), with the same tissue prepared by CF (Fig. 7B). Note the overall

lower contrast in (A) compared to (B), and how the membranes in the HPF-FS material show smooth

contours compared to those prepared conventionally. Scale bars ¼ 1.0 mm.

50 Kent McDonald

source and slows down rapidly away from the surface, it is not uncommon to find

a gradient of ice damage from the surface toward the interior of cells and tissues

(Moor, 1987; Shimoni and Muller, 1998). The diVerence between HPF and other

methods is the depth of good freezing before ice crystals appear (Robards and

Sleytr, 1985). Nevertheless, poor freezing technique with any instrument will result

in ice damage and HPF is no exception. Students of HPF need to recognize the

various forms of ice damage before they unwittingly publish poor results while

claiming superior preservation. Unfortunately, the literature of freezing does not

include many papers that specifically address the issue of ice damage and what it

looks like. Steinbrecht (1982, 1985, 1993) has been one of the few to systematically

study this problem. The beginner with freezing techniques would benefit from

reading these papers. The problem is not with severe ice damage, because that is

easy to recognize; the subtle distortions are more problematic. Another valuable

source of information for beginners about what ice crystal damage looks like is the

careful examination of their own material. It is unlikely in the extreme that all

freezing eVorts will be successful, even for the most experienced HPF researcher.

When one looks at sections of freeze-substituted and embedded samples, it is not

uncommon to find a gradient from excellent preservation to subtle or even gross

ice damage within a single section (Fig. 1 in Keene and McDonald, 1993). Those

new to cryotechniques can benefit greatly from taking lots of images of poorly

Fig. 8 Regions taken from Fig. 7 displayed at higher magnification, comparing the details of mem-

brane morphology (arrows) in cells prepared by HPF-FS (A) and by conventional methods (B). Note

how the membranes in the conventionally prepared samples are distorted compared to the smooth

membrane profiles in the HPF-FS sample. Scale bars ¼ 200 nm.


frozen material to help them learn how to recognize even subtle manifestations of

ice damage.

DiVerent cell components also vary in their susceptibility to ice damage. Those

that are highly charged, and thus bind a lot of water, are most likely to show

damage first. Chromatin is a good example; it is one of the first places to check

when evaluating the quality of freezing. Mitochondria are another essentially

universal cell component that show ice damage by having a clear ‘‘halo’’ around

their outer membrane. Ice damage to both chromatin and mitochondria in a worm

cell are illustrated in Fig. 9.

Besides ice damage, there are some structural aberrations that may be due to the

high pressure. One of these is rupture of the yeast cell wall as shown in Fig. 1.

These ‘‘cracks’’ often appear where two cells are touching, although they are not

restricted to that arrangement. Another artifact that is probably pressure-related

is the ‘‘eruption’’ of dense protein storage vacuoles that are often found in

embryonic tissues (see Fig. 6 in McDonald, 1999). As more data come in from

high-resolution tomographic studies, we may also find new categories of pressure

artifacts that have not previously been noticed.

Fig. 9 Ice-damagedC. elegans cells. Even at lowmagnification it is possible to recognize the pattern of

ice crystal damage in the chromatin (c) and the clear halos that surround some mitochondria (arrow).

Scale bar ¼ 1.0 mm.

52 Kent McDonald

VI. Summary

Following an era when many people believed LM would make EM mostly

unnecessary, EM is returning as a premier investigative tool in molecular cell

biology. In fact, LM used in conjunction with EM is emerging as one of the most

powerful approaches to learning the structural basis of cell function. Both LM and

EM are evolving technically to provide 3D visualizations of cells with increased

resolution. On the EM side, one of the most exciting developments is the emer-

gence of cellular electron tomography (cET) as an imaging method. Because cET

can image molecules at 5- to 6-nm resolution or better in the context of the intact

cell, specimen preservation techniques are of critical importance. In this chapter,

we have given detailed methods for preserving ultrastructure in several model

systems that are among the most widely used in cell biological research. When cells

of S. cerevisiae, C. elegans, andD. melanogaster are processed by HPF and fixed at

low temperature for subsequent resin embedding, the quality of the ultrastructure

and preservation of antigenicity are superior to more conventional EM methods.

We anticipate that EM and cET of these systems will play an increasingly

important role in understanding the molecular basis of cell function.

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