Has negative staining still a place in biomacromolecular electron microscopy?

Ultramicroscopy 46 (1992) 85-111 North-Holland

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Has negative staining still a place in biomacromolecular electron

microscopy? A n d r e a s B r e m e r a, Christian H e n n a, A n d r e a s E n g e l ~, W o l f g a n g B a u m e i s t e r b and Ueli A e b i ~,c " M.E. Miiller Institute for High-Resolution Electron Microscopy at the Biocenter, Unicersity of Basel, Klingelbergstrasse 70, CH-4056 Basel, Switzerland b Max Planck Institute for Biochemistry, Department for Structural Biology, W-8033 Martinsried/Munich, Germany ' Department of Cell Biology and Anatomy, The Johns Hopkins Unicersity School of Medicine, Baltimore, MD 21205, USA Received at Editorial Office 3 July 1992

Transmission electron microscopy of proteins has provided molecular- and in a few cases near-atomic-resolution structural information. In this review, we critically evaluate the potential and the limitations in obtaining molecular resolution, particularly with negatively stained specimens, and put these into perspective with cryomicroscopy of unstained frozen-hydrated and sugar-embedded preparations.

I. Introduction

Solving the first 3D structures of proteins by X-ray crystallography [1,2] revealed that proteins adopted distinct conformations. The 3D structure of a protein provides the molecular basis for its function. For instance, in spite of little sequence similarity, the atomic structures of actin, hexokinase, and the heat-shock cognate protein HSC 70 are largely superimposable since they share a conserved 3D protein framework defining the spatial arrangement of the amino acid residues involved in the specific binding and hydrolysis of their ligand, ATP [3]. Today, 3D structural analysis of proteins with resolutions ranging from molecular (i.e., beyond 3.0 nm) to near-atomic (i.e., beyond 0.4 nm) can be performed by X-ray crystallography, nuclear magnetic resonance (NMR) and electron microscopy (EM). In addition, both scanning tunneling microscopy (STM) and atomic force microscopy (AFM) can provide near-atomic-resolution images of conducting as well as insulating surfaces (for a review, see ref. [4]).

X-ray crystallography was the first and still is the predominant method for analyzing the atomic structure of proteins in spite of two severe limitations: (i) the large 3D protein crystals required for diffraction analysis are sometimes difficult to grow; and (ii) to compute electron density maps both the amplitudes and phases of the diffracted X-rays are required, but only the amplitudes can directly be measured. To provide the missing phase information, experimental (i.e., multiple isomorphic replacement) as well as knowledgebased (i.e., molecular replacement) phasing are most frequently used [5]. Hence, these methods require the availability of heavy-atom derivatives, or of previously solved structures of similar proteins. Multi-dimensional NMR has emerged as a promising method to analyze the 3D structure of proteins in solution. However, a molecular mass of about 30 kDa may define the upper size limit of molecules amenable to structural analysis by this method (for a recent review, see ref. [6]). EM is a well established, versatile and fast method to investigate protein structures at molecular or even near-atomic resolution. The major

0304-3991/92/$05.00 © 1992 - Elsevier Science Publishers B.V. All rights reserved

86

A. Bremer et al. / Negative staining

advantages of EM over both X-ray crystallography and NMR are that (i) only small quantities (i.e., /zg rather than mg amounts) of the protein are required, (ii) the protein does not necessarily have to be highly pure provided it can unambiguously be recognized in crude preparations [7], (iii) in the EM direct images containing both amplitude and phase information about the specimen can be recorded, (iv) specimens can often be analyzed in a relatively natural environment, for instance, with 2D crystals of membrane proteins reconstituted in a lipid bilayer rather than in a 3D crystal, and (v) even very large proteins or protein assemblies lacking distinct molecular symmetry can be analyzed by EM. On the other hand, structural analysis by EM has to cope with three major disadvantages: (i) the specimen has to be exposed to a high vacuum, (ii) the inherent contrast of biological matter is low, and (iii) biological matter is extremely radiation-sensitive and thus deteriorates rapidly in the electron beam. As a consequence of these limitations, the resolution of reproducible structural detail obtained with proteins is typically only about 1.5-4.0 nm, more than one order of magnitude worse than the nominal resolving power of an EM. Higher resolution, i.e., beyond 2.0-1.5 nm, usually requires major efforts. It was a "milestone" in the electron crystallography of proteins when Henderson

et al. [8] solved the 3D structure of bacteriorhodopsin in glucose-embedded purple membranes of Halobacterium halobium to 0.35 nm resolution since this structure allowed the proposition of the first atomic model of a protein based on EM rather than NMR or X-ray diffraction data.

2. Why negative staining? For EM, specimens have to be exposed to a high vacuum, hence the simplest way to prepare specimens for EM is air-drying. However, the surface tension acting on protein molecules or their supramolecular assemblies at the air-liquid interface often causes collapse and distortion of protein molecules. Removal of a protein's hydration shell leads to partial denaturation and thus renders most proteins very "sticky" [9]. Imaging proteins in a more native state in the EM necessitates (i) protection against dehydration and (ii) "sustaining", i.e., stabilization against collapse. The preparation techniques most frequently used in high-resolution EM are compared in table 1. Theoretically, keeping the specimen in a frozenhydrated state is the prime choice: dehydration and surface tension are avoided, and adsorption artifacts are eliminated since no specimen sup-

Table 1 Comparison of the most commonly used preparation methods in high-resolution EM of proteins and their supramolecular assemblies Preparation

Structure preservation

Contrast

Resolution

Support film

Radiation sensitivity

Structural information

Frozenhydrated

Very good, no dehydration

Low, PH~o = 1.0 versus

Theoretically very good, practically usually _>1 nm

Not required

High

Projected density

Required

High

Projected density

Pprotein

Very good, resolution down to 0.15 nrn has been achieved

High, due to heavy metal atoms

Moderate, typically 1.5-3 nm

Required

Low

Projected molecular envelope

Pprotein =

1.35 g/ml Sugarembedding

Negative staining

Good

Very low Psugar =

Good

A. Bremer et al. / Negatit,e staining

port film is required. Nevertheless, since the vitreous layer of ice surrounding the specimen should be very thin (ideally not much thicker than the specimen itself), some interaction of the protein with the air-liquid interface is likely to take place prior to freezing the specimen. However, the major disadvantage of frozen-hydrated preparations is their relatively low contrast and their radiation sensitivity. An alternative means to prevent or minimize spread-flattening and denatura-

87

tion of proteins and their supramolecular assemblies is air-drying in the presence of a non-volatile "sustaining" medium such as heavy-metal salts (for a review, see ref. [10]) or sugars (glucose [11]; trehalose [12]). Sugar-embedding, probably generating a non-volatile water-like environment for protein molecules, offers good sustaining of the specimen and, at the same time, allows for very high resolution. Unfortunately, the contrast of sugar-embedded specimens is very low, and they

Fig. 1. Hexagonally packed intermediate (HPI) layer. Stained with (a) aurothioglucose and (b) Cd-thioglycerol. The upper panels depict micrographs and the lower panels the corresponding correlation averages. Scale bar: 10 nm (upper panels).

88

A. Bremer et al. / Negative staining

are extremely radiation sensitive. Increasing the contrast of such preparations using heavy-atom derivatives of glucose or glycerol has been examined [13-15]. This is illustrated in fig. 1: (a) depicts the hexagonally packed intermediate (HPI) layer stained with aurothioglucose, while (b) displays the same specimen stained with Cdthioglycerol. Figs. lc and ld depict correlation averages of the above images. Cryomicroscopy (i.e., imaging at liquid-nitrogen or liquid-helium temperatures) of specimens stained with such compounds has also been explored [14,16]. Negative staining is a very rapid and simple technique, providing good sustaining and radiation protection of the specimen and, particularly with uranyl salts, high contrast (see table 2). Whereas frozen-hydrated and sugar-embedded preparations yield information on internal protein density, negative staining only reveals the structure of the stain-accessible molecular envelope in the resolution range of 1-3 nm. The fidelity of mapping structural details of a protein molecule depends primarily on the penetration properties of the heavy-metal salt used as the negative stain, i.e., its size, hydrophilicity a n d / o r hydrophobicity and other parameters. As summarized in table 2, negative stains can be used from relatively acidic pH (pH 3-4: uranyl salts, phosphotungstatic acid (PTA)) to moderately basic pH (pH 7-10: N a / K phosphotungstate, methyl phosphotungstate, methylamine tungstate, aurothioglucose), and they may be neutral (e.g., aurothioglucose, Cd-thioglycerol), anionic (e.g., tungstates) or cationic (e.g., uranyl salts).

3. Resolution, protein flexibility and ordered protein arrays

The high contrast of negatively stained preparations permits the visualization of single protein molecules. Nevertheless, revealing the reproducible structural features of a protein molecule requires one to compare many molecular images and to generate averages since (i) biological specimens are very radiation sensitive and the heavymetal replica surrounding the specimen therefore has to be imaged with relatively low electron

doses (typically 100-1000 electrons/nm 2) when trying to preserve high-resolution structural detail, thus resulting in noisy electron micrographs; and since (ii) unlike some small organic molecules, proteins are highly flexible structures. Averaging can readilybe performed with ordered arrays of protein molecules (e.g., 2D crystals, helical or icosahedral structures) that provide guidelines for the alignment of the constituting repeating units. Various naturally occurring, well ordered protein arrays have been found (e.g., the purple membrane of Halobacterium halobium [8,11], bacterial surface layers [17], and viral shells [18]. Frequently, ordered array formation can be induced artificially [19-22]. Yet, using elaborate image classification and alignment procedures, 3D reconstructions with molecular resolution have also been obtained by combining the information from projections of hundreds of single particles or molecules [23]. Multiple interactions with neighboring molecules in three dimensions can minimize the deviations from ideal lattice points, rendering 3D crystals frequently highly ordered, and thus diffracting in X-ray crystallographic experiments to near-atomic resolution. 2D crystals and helical arrays as they are frequently used in EM are more prone to departures from perfect crystallinity since the constituent protein molecules establish fewer interactions with neighboring molecules. Molecular disorder (i.e., protein flexibility), lattice disorder, and lattice defects (e.g., ref. [24]) can limit the resolution achievable by EM. In other words, to increase the resolution, the order of a protein array has to be improved, or the lattice distortions have to be corrected by digital image processing. Increasing the crystalline order of a protein array by specific treatments (e.g., lipase digestion [25]) or by adding specific effector molecules has been demonstrated. The stabilizing effect of one such effector molecule, the mushroom toxin phalloidin, on the structure of the F-actin filament is illustrated by fig. 2: (a) reveals control F-actin filaments, and (b) depicts phalloidin-stabilized Factin filaments. Phalloidin stabilization decreases the intrinsic disorder of F-actin filaments: the crossover spacing histograms shown as insets in

Table 2 Properties of various negative stains a) Stain

Density (g/ml)

Useful pH range

Radiation sensitivity

Contrast

References

Comments

Uranyl acetate

2.89

3- 4

Moderate

High

Van Bruggen et al. [101]

Fixative effect

Uranyl oxalate

2.50-3.07 b)

3- 7

Moderate

High

Haschemeyer and Meyers [10]; Mellema et al. [102]

Very light sensitive, store frozen

Uranyl nitrate

2.81

3- 4

Low

High

Uranyl formate

3.70

3- 4

Moderate

High

Haschemeyer and Meyers [10]; Leberman et al. [103]

Fixative effect, smallest grain size

Uranyl sulfate

3.28

3- 4

Low

High

Estis et al. [104]

Reported not to recrystallize upon irradiation with electrons

Na/Kphosphotungstate

1.69 c~

4- 9

Low

High

Brenner and Horne [105]

Positive staining, increases with lowering the pH; destructive effect on phospholipid membranes

Na silicotungstate

2.84 d)

4- 8

High

High

Sherman et al. [106], Terry [107] Haschemeyer [108]

4 - 9.5

Low

Medium

Oliver [109]

Methylphosphotungstate Methylamine tungstate

3.88

3-10

low

High

Faberge and Oliver [110], Shaw and Hills [111]

Supposed not to be a positive stain at any pH. With glycoproteins, add tannic acid

Ammonium molybdate

2.28

5- 8

Moderate

Medium

Bohonek [112], Manella and Frank and [113]

Good for membranes, some fibrous proteins

Aurothioglucose

2.92

4-10

High

Low

Kiihlbrandt [114], Kiihlbrandt and Unwin [13]

Yields Aucrystallites upon electron irradiation

Cadmiumthioglycerol

2.0

4-10

Moderate

low

Jakubowski et al. [15]

No crystallite formation upon electron irradiation, possibly useful with undecagold

Vanadate

2.85 e)

low

low

Very light stain, can be used with undeca-gold labelling

a) Most of the density values were obtained using the Gmelin on-line database for inorganic and metallo-organic compounds. Many of the other data were originally compiled by C.L. Woodcock. b) Depending on the amount of bound water. c) Density of a saturated solution of phosphotungstic acid at 22°C. a) Density of silicotungstic acid. e) Density of NaVO 3.

90

A. Bremer et al. / Negatit,e staining

Fig. 2. Effects of the mushroom toxin phalloidin on the F-actin filament structure. (a) F-actin filaments polymerized with 2mM MgCl 2/50raM KCI for 2 h at room temperature; (b) F-actin filaments polymerized similarly but in the presence of phalloidin in a 2 : 1 molar ratio relative to actin. (Insets) Side-on and end-on views of volume-rendered 3D reconstructions of such filaments. With phalloidin-stabilized F-actin filaments, a pronounced "bridge" (indicated by arrows) putatively comprising the phalloidin binding site that connects adjacent subunits across the two long-pitch helical strands of the F-actin filament. Compared to the control (a), the phalloidin-stabilized filament s (b) appear more compact with more regular crossover spacings. This is quantitated in the respective crossover spacing histograms included in (a) and (b). The mean crossover spacings as well as the standard deviations are indicated. Scale bars: 100 nm.

Table 3 Compilation of structure determinations with a resolution of 1.0 nm or better from electron micrographs a n d / o r electron diffraction patterns recorded by cryomicroscopy Specimen

Type of Array

Subunit size and stoichiometry

Reconstruction

Preparation

References

Purple membrane (i.e., bacteriorhodopsin)

2D

Membrane protein, 27 kDa

3D reconstruction, 0.35 nm resolution parallel to the membrane

Glucoseembedding

Henderson et al. [8]

2D reconstruction, 0.28 nm resolution

Glucoseembedding

Baldwin et al. [78]

2D reconstruction, 0.39 nm resolution

Glucoseembedding

Bullough and Henderson [79]

Three different 2D reconstructions, 0.34 nm resolution

Tannic acidpreserved, glucoseembedded, and frozen-hydrated specimens

Wang and Kiihlbrandt [80]

3D reconstruction, 0.6 nm resolution

Tannic acidpreservation

Kiihlbrandt and Wang [81]

Lightharvesting complex

PhoE porin

2D

2D

Membrane protein, three different subunits with 25 kDa each

Membrane protein, 37 kDa

3D reconstruction, about Trehalose0.6 nm resolution parallel embedding to the membrane and about 0.8 nm resolution normal to it

Jap et al. [82]

2D reconstruction, 0.35 nm resolution

Trehaloseembedding

Jap et al. [12]

OmpF porin

2D

Membrane protein, 37 kDa

2D reconstruction, 0.35 nm resolution

Glucoseembedding

Sass et al. [83]

Crotoxin

3D

2 subunits, 10 and 14 kDa

2D reconstruction, 0.35 nm resolution

Glucoseembedding

Jeng et al. [84]

2D reconstruction, 0.4 nm resolution

Glucoseembedding

Bullough and Tulloch [32]

t~-helical coiledcoil protein from praying mantis CaATPase

3D

Membrane protein, 109 kDa

2D reconstruction, 0.6 nm resolution

Frozen-hydrated

Stokes and Green [85]

Cytochrome oxidase

2D

Membrane protein, 7 subunits with M r ranging between 4.5 and 45 kDa; 125 kDa total

2D reconstruction, 0.7 nm resolution

Negatively stained

Valpuesta et al. [86]

Bacterial S-layer

2D

Extracellular proteins, 2D reconstruction, monomer > 140 kDa 1.0 nm resolution

Negatively stained

Lembcke and Zemlin [16]

monomer 91 kDa

2D reconstruction, 0.8 nm resolution

Negatively stained

Rachel et al. [14]

T4 DNA helix destabilizing protein gp 32 * I

Multilayered sheets

Cytoplasmic protein, 27 kDa

2D reconstruction, 0.85 nm resolution

Frozen-hydrated

Grant et al. [87]

Tobacco mosaic virus

Helical

C~toplasmic protein, 17.5 kDa

3D reconstruction, 1.0 nm resolution

Frozen-hydrated

Jeng et al. [88]

The type of reconstruction (i.e., 2D or 3D), the resolution achieved and the specimen preparation (i.e., sugar-embedding, frozen-hydrated, or negative staining) are indicated. The order of the examples listed is according to the resolution.

92

A. Bremer et al. / Negatit,e staining

fig. 2 reveal the same mean spacing of 36.1 nm for phalloidin-stabilized and control F-actin filaments but a statistically significant decrease in the standard deviation with phalloidin-stabilized filament preparations (_+ 2.7 nm versus ___3.4 rim; n = 200). The structural basis for this increased filament order is an increased strength of the intersubunit contacts between the two long-pitch helical strands of actin subunits in the F-actin filament: the phalloidin-stabilized filaments appear more compact (compare figs. 2a and 2b), an impression that is confirmed by 3D reconstructions (see insets in fig. 2) showing additional mass in the form of "bridges" (marked by arrows in fig. 2b) that probably contain the phalloidin binding site and connect adjacent subunits between the two long-pitch helical strands of the F-actin filament (see also ref. [26]).

Digital processing of electron micrographs can be used to at least partially correct for lattice imperfections that can be parametrized (e.g., correction of lattice distortions [27-29], or unbending of slightly curved filaments [26,30,31]). Realspace averaging techniques in conjunction with pattern recognition [24] or distortion analysis [29] have become powerful means to retrieve a maxim u m of structural information from imperfect 2D protein arrays.

4. Resolution and preservation of structural detail High-performance EMs can readily attain atomic resolution with radiation-resistant inor-

Table 4 Survey of specimens investigated with both negatively stained and frozen-hydrated preparations Specimen

Negative staining

Frozen-hydrated

Ca 2 + release c h a n n e l / ryanodine receptor

2D, 2.5 nm resolution; reconstruction of Saito et al. [89], resolution stated in Radermacher et al. [90]

2D, 3.0 nm; R a d e r m a c h e r et al. [90]

3D, 3.7 nm: Wagenknecht et al. [91] Actin filaments

3D, 2.8 nm; Bremer et al. [26]

sheets

3D, 1.5 nm; Smith et al. [38]

T4 D N A helix destabilizing protein gp 32"1

3D, 1.5-1.8 nm (anisotropic); Grant et al. [87]

3D, ~ 3.0 nm; Milligan et al. [57]

2D, 0.85 nm; Grant et al. [87]

3D, 2.0-3.0 n m (anisotropic); Cohen and Chiu [92]

CaATPase

2D, 1.5 nm; Stokes and Green [93]

2D, 0.6 nm; Stokes and Green [85]

Cytochrome oxidasc

2D, 0.7-1.5 nm, depending on the stain and the temperature; Valpuesta et al. [86]

2D, 1.5 nm; Valpuesta et al. [86]

Fragments of the N A D H : ubiquinone oxidoreductase

2D, 1.5-3.8 nm, depending on the stain used; Brink et al. [52]

2D, 1.4-1.9 nm, depending on the amount of defocus (i.e. 400 n m - l . 3 ~m); Brink et al. [94]

Tobacco mosaic virus

3D, 1.2 nm; Unwin and Klug [95]

3D, 1.0 nm; Jeng et al. [88]

Bacterial T-layer

3D, 2.5 nm; Lepault et al. [96]

2D, better than 3.0 nm; Lepault and Pitt [97]

Gap junctions

2D, 1.8 nm; Zampighi and Unwin [98]

2D, 2,0 nm; Unwin and Ennis [99]

3D, 1.8 nm; Unwin and Zampighi [100] Indicated are the type of reconstruction (i.e., 2D or 3D), the resolution achieved and the reference. The examples are listed inversely chronologically (i.e., starting with the most recent reference).

A. Bremer et al. / Negatit'e staining

ganic materials, e.g., gold crystals. Electron diffraction patterns recorded from glucose-embedded 2D crystals of biological matter have also

93

demonstrated a resolution of up to 0.15 nm [32]. As is documented in table 3, structure determination to resolutions significantly better than 1.0 nm

b (-5,3)

( 4 , 0 ) '.

Fig. 3. Negatively stained crystalline actin sheets. (a) Electron micrograph of a double-layered rectangular actin sheet recorded under low-dose conditions. Staining was with 0.75% uranyl formate, pH 4.25. (b) Representative optical diffraction pattern of rectangular actin sheets. The highest diffraction orders are indicated: the (4, 0) spot corresponds to a 1.4 nm spacing while the ( - 5 , 3) spot corresponds to a spacing of 1.0 nm. (c) Single-sided Fourier filtration reveals a unit cell containing two actin molecules that are related by a two-fold axis of symmetry. The actin molecules appear to be composed of two domains of similar size. Negative stain is printed black, protein bright. Scale bars: 100 nm (a), (2.5 nm) i (b), 5 nm (c).

A. Bremer et al. / Negatit~e staining

94

b

- "'= ( - 1 , 6 )

(4,0)

T

Fig. 4. Frozen-hydrated actin sheets. (a) Electron mlcrograph of a double-layered rectangular actin sheet recorded under low-dose conditions (prepared and recorded by Dr. J. Lepault, at the E M B O Lab, Heidelberg). (b) Optical diffraction pattern (composed of two different halves) recorded from sheets similar to the one in (a). The highest diffraction orders are indicated: the (4, 0) spot corresponds to a 1.4 nm spacing while the ( - 1, 6) spot corresponds to a spacing of 1.1 nm. (c) Single-sided Fourier filtration reveals a unit cell containing two actin molecules that are related by a two-fold axis of symmetry. Compared to fig. 3c, the actin molecule is less obviously composed of two domains. Ice is printed black, protein bright. Scale bars: 100 n m (a), (2.5 nm) I (b), 5 nm (c).

A. Bremer et al. / Negative staining

has been possible with a few biological specimens. Theoretically, frozen-hydrated preparations should yield optimal preservation (see above and table 1). EM of frozen-hydrated specimens is commonly thought of as being synonymous with higher resolution compared to negative staining, albeit without firm experimental support. Table 3 summarizes 18 different 2D and 3D reconstructions extending to a resolution of 1.0 nm or better (for a review, see ref. [33]). Out of these, the vast majority (nine) were determined by cryomicroscopy of sugar-embedded specimens, whereas the number of _< 1.0 nm-resolution reconstructions determined from frozen-hydrated specimens (four) and from negatively stained specimens (three) was almost identical. Table 4 depicts specimens where reconstructions have been computed from both negatively stained a s well as frozen-hydrated preparations. Accordingly, negatively stained preparations mostly yield very similar (e.g., TMV, N A D H : ubiquinone oxidoreductase, gap junctions, actin) or even higher (Ca 2+ release channel, cytochrome oxidase), rather than lower (see T4 DNA helix destabilizing protein, Ca2+-ATPase) resolutions to frozen-hydrated preparations. This apparent discrepancy has been ascribed to beaminduced specimen/film movements, charging effects and background scatter [2l]. Moreover, sugar-embedding and negative staining may help to decrease bending or other types of long-range disorder resulting from the lack of a specimen support. Most proteins in table 3 are relatively small (less than 37 kDa) and thus may be relatively compact and rigid compared to large multi-domain proteins. In addition, 12 of the 18 specimens listed in table 3 are membrane proteins that are possibly protected against preparation artifacts at least in their membrane-spanning parts. The structures of the larger proteins listed in table 3 have either been determined from 3D crystals (Ca2+-ATPase) that may shield their internal subunits, or from sustained preparations (negatively stained: cytochrome oxidase, S-layer proteins; tannic acid-embedded: light-harvesting complex). With the exception of the light-harvesting complex, the specimens analyzed by EM of

95

frozen-hydrated specimens are 3D, or multilayered 2D crystals (Ca2+-ATPase, T4 DNA helix destabilizing protein gp 32" I) or otherwise very sturdy structures (TMV). Again, with the exception of the light-harvesting complex, the resolution achieved with frozen-hydrated specimens is considerably lower than with sugar-embedded specimens. Hence, specimen support also emerges as a possible resolution-increasing factor. Both glucose-embedded and negatively stained specimens have been adsorbed to a flat supporting film while non-supported frozen-hydrated specimens may exhibit long-range disorder such as bending. Adsorption does not at all decrease the resolution: with double-layered structures, the face in contact with the support film yields higher-resolution structural detail and is better ordered [34,35]. Thus, 2D arrays of small membrane proteins and 3D arrays of larger proteins, preferentially sugar-embedded, seem to be the specimens of choice for high resolution electron crystallography. As can be gathered from table 4, negative staining has the potential to preserve the structure and crystalline order of proteins to a resolution of 1.0 to 2.0 nm. This is illustrated in figs. 3 and 4: negatively stained (fig. 3a; see ref. [36]) as well as frozen-hydrated (fig. 4a; see ref. [37]) crystalline actin sheets diffract to a resolution of about 1.0 nm (see figs. 3b and 4b, diffraction patterns). Fourier-filtered images appear similar though there are subtle differences which are difficult to interpret in projection images. A rigorous comparison would require 3D reconstructions which, as yet, have only been obtained for negatively stained actin sheets to a resolution of 1.5 nm [38]. Apparently, the averaged unit cell of frozen-hydrated actin sheets reveals less structural detail than that of negatively stained sheets (compare fig. 3c with 4c).

5. Resolution and radiation damage

The radiation-induced limitation of resolution can be minimized by low-dose EM but also by radiation protection by negative staining, i.e., by shielding the biological material with a heavy-

96

A. Bremer et al. / Negatit.e staining

metal salt replica. Cryomicroscopy, i.e., k e e p i n g the s p e c i m e n at l i q u i d - n i t r o g e n or l i q u i d - h e l i u m t e m p e r a t u r e , can also increase the r a d i a t i o n resistance of biological material a n d h e n c e increase the a t t a i n a b l e resolution (for a recent review, see ref. [39]). However, at least for a few negatively stained specimens, there is n o obvious correlation b e t w e e n the a c c u m u l a t e d electron dose a n d the

achievable resolution. This is d e m o n s t r a t e d in fig. 5 where images of the T-layer of B a c i l l u s bret'is have b e e n recorded at increasing a c c u m u l a t e d electron doses (i.e., from 210 (fig. 5a) to 2900 e l e c t r o n s / n m 2 (fig. 5d)), a n d the c o r r e s p o n d i n g diffraction patterns, correlation averages, a n d radial a m p l i t u d e correlation functions [28] are compared. In this case, the resolution d e t e r m i n e d

Fig. 5. Dose series of a T-layer of Bacillus bre~,is. The same region of a negatively stained T-layer (left column) was imaged in a scanning transmission EM with an electron dose of (a) 210 e /nm 2, (b) 380 (accumulated dose 590) e /rim 2, (c) 760 (accumulated 1350) e /nm z, and (d) 1550 (accumulated 2900) e /rim 2. For each image shown, the diffraction pattern, a correlation average including ~ 400 unit cells, and the radial amplitude correlation function (from left to right) are shown. Scale bars: 100 nm (image), and 10 nm (correlation average).

A. Bremer et al. / Negatice staining

1.0

1.0

0.5

0 5

97

0.0" l

0.0

0.2

I

0.4

I

0.6 nm" 1

I

0.8

1.0

0.0

0.2

0.4

0.6 nm-1

0.8

1.0

Fig. 6. Effect of the electron dose on the resolution attainable. A negatively stained (0.75% uranyl formate, pH 4.25) E. coli O m p F porin sheet has been imaged with an accumulated electron dose of (a) 2000 e l e c t r o n s / n m 2 and (b) 10000 e l e c t r o n s / n m 2. The corresponding radial amplitude correlation functions are displayed in (c) and (d), revealing a resolution of about 1.1 nm for the lower-dose and about 1.8 nm for the higher-dose image. The correlation averages computed for both electron dose images are displayed in (el and (g). Inspection of the difference image between (el and (g) that is depicted in (f) reveals that the " b o o m e r a n g - s h a p e d " structures lining the pores ((e-g), black arrowheads) are "'blurred" due to a redistribution of the negative stain primarily by migration into the pores (black arrows), and by shrinkage a n d / o r redistribution into the lipid-bearing regions (white arrowheads). Scale bars: 50 nm (a, b), 5 nm (e-g).

A. Bremer et al. / Negatit,e staining

98

b

I

°

C 1.0

m

90°~ 0.5 m 45°

I

]

I

~

0~

1

0.5

1.0

1.5 [nm-l]

'

0

~

t

i

i

[

0.5

I

t

J

i

J

1.0

I

I

1

I

1.5 [nm-ll

Fig. 7. Revealing distinct resolution-dependent structural features with negatively stained bacteriophage T4 polyheads. (a) Negatively stained polyhead (40 min 1% sodium silicotungstate, 1 min 1% uranyl acetate) imaged in a scanning transmission EM. (b) The diffraction pattern of such a polyhead exhibits distinct six-fold symmetry (left half). The right half of the diffraction pattern was obtained from an average of seven polyheads (amounting to a total of about 700 unit cells) after six-fold symmetrization. (c) Radial amplitude (left) and phase (right) correlation functions of this polyhead average. The Fourier-filtered polyhead average was limited to a nominal resolution of (d) 3.0 and (f) 1.25 nm. As is evidenced by the difference image (e), including the resolution range between 1.25 and 3.0 nm into the correlation average reveals additional structural information: the protomers (see arrows) appear more or less globular in (d) while they are "leaf-shaped" in (f). Moreover, the interprotomer connectivity (arrowheads) present in (f) is absent in (d). Scale bars: 20 nm (a), (1.9 n m ) - 1 (b), 10 nm (d-f).

A. Bremer et al. / Negative staining

from the radial amplitude correlation function increases with increasing electron dose from 2.78 nm in fig. 5a to 1.72 nm in fig. 5d. The effect of the accumulated electron dose on the nominal resolution and the correlation averages of a negatively stained crystalline sheet reconstituted from E. coli OmpF porin is demonstrated in fig. 6: (a) depicts an image recorded with an electron dose of 2000 electrons/nm 2 while (b) reveals the same sheet after an accumulated electron dose of 104 electrons/rim 2. The respective radial amplitude correlation functions [28] are shown in figs. 6c and 6d and yield a resolution of about 1.l nm for the lower-dose and of about 1.8 nm for the higher-dose image. Figs. 6e and 6g depict the correlation averages computed for the images in figs. 6a and 6b. The "boomerang-shaped" structure (see black arrowheads) lining the pores in figs. 6e and 6g (protein appears bright) is separated into two distinct domains in fig. 6e which are no longer resolved in fig. 6g. Comparison of the difference image of figs. 6e and 6g that is depicted in fig. 6f reveals significant redistribution of the negative stain by migration into the pores (marked by black arrows

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in the central part of figs. 6e and 6g), and into the lipid-bearing regions (see white arrowheads). Possible factors and mechanisms governing such dose-dependent stain redistribution and shrinkage have been explored in detail before [40]. 6. The merits of increased resolution

The amount of biologically interpretable - or useful - information is not simply a question of nominal resolution, e.g., as judged by "counting diffraction orders". For instance, in the 2 nm resolution 3D reconstruction of negatively stained actin-DNase I tubes [41], it was relatively straightforward to separate the actin from the DNase I moiety after fitting the actin dimer [38] into the map. By contrast, the X-ray crystallographic 0.6 nm resolution electron density map of the actin-DNase I complex alone did not unambiguously allow separation of the electron density contributed by the actin moiety from that of the DNase I moiety [42]. At least in part, the similarity of the electron density in intersubunit and inter domain interfaces may account for this difficulty.

Fig. 8. Native neurofilaments isolated from bovine spinal cord. (a) Glycerol-sprayed/rotary metal-shadowed preparations of neurofilaments appear distinctly millepede-like due to their "side-arms". (b) Upon negative staining with uranyl formate (0.75%, pH 4.25), these side-arms probably collapse into "knobs". Scale bar: 100 nm.

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A. Bremer et al. / Negatice staining

The additional information provided by an extra diffraction order that amounts to 0.1% or even 1% of the total power of a diffraction pattern may sometimes only be marginal, particularly with negatively stained specimens. Therefore, "differential filtering" [43] should be applied to decide whether particular diffraction spots really increase the resolution in the sense that additional structural features are revealed upon adding them to the Fourier-filtered image. This is illustrated in fig. 7 where an average of 7 bacteriophage T4 polyheads (corresponding to about 700 unit cells) is differentially filtered [43]. The difference image (fig. 7e) includes the resolution range between 1.25 and 3.0 nm. U p o n adding this resolution to the image limited to 3.0 nm resolution (fig. 7d), additional structural information is revealed: the protomers are transformed from a more or less globular shape in fig. 7d into a " l e a f - s h a p e " in fig. 7f (see arrows) and establish connectivity in the region of the c a p s o m e r center (arrowheads). Nevertheless, often the high-resolution diffraction spots of negatively stained specimens only cause edge-sharpening of the filtered

image rather than adding distinct structural detail. Therefore, filtered images rather than only diffraction patterns or radial correlation functions [28] should be compared.

7. Different preparation methods and different stains may reveal different aspects of the same specimen Proteins and their supramolecular assemblies may respond sensitively to changes in their aqueous environment (e.g., pH, ionic strength, concentrations of multivalent cations) with sometimes dramatic structural changes. Often, the structure of protein assemblies seen by EM depends on the preparation technique used. As shown in fig. 8a, " s i d e - a r m s " r e n d e r neurofilaments distinctly millepede-like in g l y c e r o l - s p r a y e d / r o t a r y metalshadowed preparations [44,45]. In contrast, as illustrated in fig. 8b, negatively stained preparations of the same sample reveal " k n o b s " rather than side-arms, possibly as a result of collapse or aggregation of the side-arms onto the central rod

Fig. 9. Negatively stained cultured human epidermal cell (CHEC) keratin filaments. (a) CHEC keratin filaments in 10mM Tris, pH 7.5, and negatively stained with 0.75C¢ uranyl formate, pH 4.25, appear compact with more or less uniform width. (c) Negative staining with 25~ sodium phosphotungstate, ptl 7.0, reveals local unraveling of the same filaments. (b) Even more dramatic unraveling is observed when the filaments are briefly washed with 10mM sodium phosphate (NAP,, pH 7.0) prior to staining them with 0.75(/~ uranyl formate, pH 4.25. Scale bar: 100 nm.

A. Bremer et al. / Negatice staining

of the filament. The heavy-metal salt used as the negative stain for contrasting the specimen may also affect its structure. As illustrated in fig. 9a, cultured human epidermal cell (CHEC) keratin filaments in 10raM Tris, pH 7.5 appear rather compact and featureless if stained with uranyl formate (pH 4.75). By contrast, if stained with

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sodium phosphotungstate (NaPT, pH 7.0), they become locally unraveled (see fig. 9c). However, this result does not just represent a preparation artifact: if the filaments are briefly washed with 10raM sodium phosphate (NaP i, pH 7.0) and negatively stained with uranyl formate, they appear even more unraveled (see fig, 9b). These data

Fig. 10. Effect of the state of hydrolysis of the specifically bound nucleotide on the structure of the F-actin filament. (a) ADP-F-actin filaments resuspended in 5mM ATP, and (b) filaments polymerized from monomers that have the nonhydrolyzable ATP analog AMP-PNP bound appear compact with more or less evenly spaced crossovers, with a clean background (i.e., no significant amounts of monomers or small oligomers), and little if any indication of local unraveling (arrowhead in (a)). (c) Compared to these putative structural analogs of the ATP-F-actin state, ADP-F-actin filaments unveil more internal structural (i.e., they are less compact), yield a significantly increased background, and exhibit frequent local unraveling of their two long-pitch helical strands (see arrowheads). (d) These features become even more obvious in the presence of 50ram sodium phosphate (NAP,, pH 7.4). With such preparations, "splayed ends" (i.e., local unraveling of filament ends) are sometimes observed. Negative staining in ( a - d ) was with 0.75% uranyl formate, pH 4.25. (e) Similar to uranyl-formate-stained ADP-P,-F-actin filaments (d), ADP F-actin filaments negatively stained with 1% sodium phospbotungstate, pH 7.0, reveal even more dramatic local unraveling and splayed ends (see arrowheads). Scale bar: 100 nm.

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A. Bremer et al. / Negatit'e staining

suggest that P~ may act as a modulator of the lateral interaction of protofilaments or protofibrils in the filament [46]. In fact, a similar response has been observed with F-actin filaments where the state of hydrolysis of the specifically bound nucleotide appears to modulate the relative strength of the bonds along and between the two long-pitch helical strands defining the filament [47-49]. This phenomenon is demonstrated in figs. 10a-10d: F-actin filaments appear compact in the presence of ATP (fig. 10a) or the nonhydrolyzable ATP analog, AMP-PNP (fig. 10b). F-actin filaments with bound ADP (fig. 10c) appear less compact and show indications of local unravelling into their two long-pitch helical strands (arrowheads). This is even more dramatic after adding mM amounts of Pi (fig. 10d). As with CHEC keratin filaments (see above, fig. 9), the specific effect of P~ is mimicked by staining with NaPT (see fig. 10e). Similar structural changes in response to the state of hydrolysis of the specifi-

cally bound nucleotide have also been observed with frozen-hydrated F-actin filaments [50]. This latter observation eliminates the possibility that local unraveling of F-actin filaments [26] is solely a specimen preparation artifact caused by negative staining a n d / o r air-drying. The fidelity of mapping structural details of a protein molecule by negative staining depends primarily on the penetration properties of the heavy-metal salt used, i.e., its size, hydrophilicity a n d / o r hydrophobicity as well as other parameters. To explore possible differences in structural details depending on the stain used, Woodcock and Baumeister [51] have compared the structure of the tetragonal surface layer of Clostridium aceticum using various negative stains. As documented in figs. l l a - l l d , in 3D reconstructions of specimens stained with sodium phosphotungstic acid (PTA)(figs. l l a and 11c) and uranyl acetatc (UA) (figs. l l b and l i d ) representing two extreme situations, the primary structural differ-

~ ~c>....~

i

h

Fig. I 1. The effect of different negative stains on the 3D structure of the surface layer of Clostridium aceticum. 3D reconstructions of sodium phosphotungstic-acid-stained preparations are shown in the upper row, while the lower row depicts reconstructions of urany]-acetate-stained specimens. Volume-rendered representations of (a,b) the outside and (c,d) the inside of 3D reconstructions. (e,f) 2D projections of the 3D reconstructions depicted in (a-d) perpendicular to the plane of the surface layer. (g,h) Schematic representations of the projected asymmetric unit cells shown in (eX). The major structural differences between the two reconstructions are observed in their connectivity patterns (compare a and c with b and d): accordingly, an outward movement of the center of mass of the protomer (highlighted by a circle in the schematic representation in (g,h)) in the reconstruction of uranyl-acetate-stained versus phosphotungstic-acid-stained preparations prevents establishing contact between the side-lobes. Scale bar: 5 nm (a f).

A. Bremer et al. / Negatice staining

ences are observed in the connectivity between the protomers. In 2D projections of these 3D reconstructions (figs. l l e and llf), the differences appear less pronounced, resulting in a small movement of the center of mass of the protomer (schematically shown in figs. l l g and llh). This

103

movement enables the side-lobes of the protomer to make contact in the reconstruction of PTAstained but not of the UA-stained specimens. However, it is not clear whether these differences are due to different stain distributions or to alterations of the structure during staining. Likewise, Brink et al. [52] compared 2D reconstructions of 2D crystals of fragments of N A D H : u b i q u i n o n e oxidoreductase stained with ammonium molybdate, phosphotungstic acid, uranyl acetate, uranyl nitrate, uranyl sulfate, sodium tungstate and sodium silicotungstate. Multivariate statistical analysis unveiled subtle systematic differences between reconstructions obtained using the different stains. However, the lattice vectors of the 2D crystals were not affected by the different stains. Comparison of the staining properties of different heavy-metal salts can also provide information other than purely structural. For instance, Baker et al. [53] stained gap junctions with one molybdate, three tungstate, and three uranyl stains. The cationic uranyl stains penetrated the axial connexon channel whereas the anionic stains were largely excluded. This result was interpreted to indicate that negatively charged residues may line the channel.

Fig. 12. Comparison of a 3D reconstruction from negatively stained F-actin filaments with electron density maps computed from an atomic model of the actin filament. A volumerendered representation of a 3D reconstruction from negatively stained F-actin filaments (average over about 200 subunits from three different filament stretches) is shown to the left at two different contouring levels to include 100% (top) and 50% (bottom) mass assuming a hydrated mass density of 810 D a / n m -~. Electron density maps for an atomic model of the actin filament [56] have been computed to nominal resolutions of 2.5, 3.0, and 3.5 nm, respectively, and volume-rendered as described for the EM-based 3D reconstruction shown to the left. The shape of the actin subunit that is composed of two domains demarcated by a cleft (indicated by a line in the top row) as well as the intersubunit contact between the two long-pitch helical strands (see arrows in top row) in the 3D reconstruction are very similar to the 3.0 nm resolution electron density map. The 50% mass representation of the 3D reconstruction displays both intersubunit contacts along and between the two strands; the intersubunit connectivity pattern therefore more closely resembles the 2.5 nm resolution electron density map. Scale bar: 5 nm.

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A. Bremer et al. / Negatil'e staining

Fig. 13. Comparison of the actin molecule as revcaled by 3D reconstruction of negatively stained F-actin filaments with differenl representations of the atomic structure of the actin molecule. I'art of one volume-rendered long-pitch helical strand of the reconstruction shown in fig. 12 is overlaycd onto (left) a ribbon representation and (right) a space-filling van der Waals sphere representation of thc atomic structure of the actin moleculc [55]. The major structural differences are observed in the rcgion of the actin molecule corresponding to the so-called DNasc 1 binding loop (see arrows, for discussion see ref. [49]). Scale bar: 0.5 nm.

8. Negatively stained F-actin filaments: comparison with X-ray diffraction data A b u n d a n t i n f o r m a t i o n on t h e s t r u c t u r e o f the actin m o l e c u l e a n d t h e actin f i l a m e n t is a v a i l a b l e , t h u s a l l o w i n g for a m e a n i n g f u l critical c o m p a r i son o f e l e c t r o n m i c r o s c o p y and X - r a y d i f f r a c t i o n d a t a (fl~r a review, see refs. [49,54]): (i) t h e a t o m i c s t r u c t u r e s o f t h e actin m o l e c u l e in c o m p l e x with D N a s e 1 [55] and t h e g e l s o l i n f r a g m e n t 1 [P. McLaughlin, personal communication] have been d e t e r m i n e d by X - r a y d i f f r a c t i o n analysis; (ii) an a t o m i c m o d e l of t h e F - a c t i n f i l a m e n t has b e e n built [56] by t r a n s f o r m i n g the c o o r d i n a t e s of an

a t o m i c s t r u c t u r c of t h e actin m o l e c u l e to fit 0.8 n m r e s o l u t i o n X - r a y f i b e r d i f f r a c t i o n d a t a obt a i n e d with o r i e n t e d gels o f p h a l l o i d i n - s t a b i l i z e d F - a c t i n f i l a m e n t s : (rid a 1.5 n m r e s o l u t i o n 3 D m o d e l of t h e actin m o l e c u l e was d e t e r m i n e d f r o m e l e c t r o n m i c r o g r a p h s o f 2 D c r y s t a l l i n e actin s h e e t s [38]; a n d (iv) h i g h - r e s o l u t i o n 3 D r e c o n s t r u c t i o n s of e l e c t r o n m i c r o g r a p h s o f f r o z e n - h y d r a t e d [57] as well as n e g a t i v e l y s t a i n e d F - a c t i n f i l a m e n t s [26] have been presented and are remarkably similar to e a c h o t h e r ( r e v i e w e d in ref. [49]). T h e r e f o r e , actin f i l a m e n t s can s e r v e as a test s p e c i m e n to critically c o m p a r e E M - b a s e d 3 D r e c o n s t r u c t i o n s with results o b t a i n e d by X - r a y d i f f r a c t i o n .

Fig. 14. Identification of the ~- and ,6-subunits of proteasomcs from Thermoplasma acidophilum. Protcasomcs were reacted with antibodics directed against (a) the ~-subunit and (b) the ~-subunit and negatively stained with 2% uranyl acetate. Formation of distinct comph:xes between antibody molecules and protcasomcs is observed: (a) anfibody-proteasome complexes fl~rmcd with anti-~-subunit antibodies mostly join thc proteasomes in an end-to-end fashion, whereas (b) side-by-side aggregates predominate with anti-~-subunit antibodies [67]. (insets) Low-pass-filtered images of selected antibody proteasome comple×es illustrate the distinct complex formation more clearly. (c) Single-particle average of the Thermol?lasma proteasome determined from negatively stained preparations. (d) 3D model of the Thermoplasma proteasome consisting of two outer ~-subunit disks (dark) and two inner ~-subunh disks (bright: see also refs. [67,68]). Scale bars: 100 nm (a,bk 5 nm (c,d).

A. Bremer et al. / Negatit'e staining

d

105

106

,4. Bremer et a L / Negatit,e staining

In fig. 12, a 3D reconstruction of negatively stained F-actin filaments (left panel) is compared to electron density maps (right panels) computed

from an atomic model of the F-actin filament (according to ref. [56]). The electron density maps were limited to a resolution of 2.5, 3.0, and 3.5

Fig. 15. Specific labeling of distinct bacteriophage T4 polyhead capsomer epitopes using monovalent Fab fragments. (a-c) Electron micrographs and ( d - f ) their respective Fourier-filtered unit cells (for nomenclature, see ref. [64]) of negatively stained polyheads, a polymorphic variant of the E. coli bacteriophage T4 capsid, are shown. (a,d) Native polyhead, (b,e) polyhead stoichiometrically decorated with label 1, and (c,f) polyhead stoichiometrically decorated with label 2. Labels 1 and 2 are monovalent Fab fragments prepared from rabbit antisera directed against different antigenic determinants on the major capsid protein gp23* (for details, see refs. [63,64]). In (d-f), the upper panel shows roughly one "superunit" cell, the middle panel depicts the same data with a one-contour level map of the native polyhead superimposed to facilitate detecting displacements/rotations of the six "petals" relative to the position in the native polyhead. In the lower panel, a difference map obtained by subtracting the superunit cell of native polyheads ((e), top) from the respective top panel. As in the middle panel, a one-contour level map of the native polyhcad is superimposed. Scale bars: 100 nm (a-c), 10 nm (d-f).

A. Bremer et al. / Negatit~e staining

nm (left to right), respectively, while the 3D reconstruction of negatively stained F-actin filaments included data extending to a nominal resolution of 2.8 nm. The upper panels of fig. 12 are contoured to include 100% of the total mass. The lower panels are contoured to include only 50% of the total mass and reveal a resolution-dependent transition in the high-density intersubunit connectivity pattern from contact along and between the two long-pitch helical strands to exclusive contact between the two strands. Thus, possibly as a result of comparatively low resolution, some previous EM-based 3D reconstructions and models of the F-actin filament using a two-sphere approximation, or a low-resolution structure of the actin subunit, were erroneously interpreted to indicate strong contact between the two long-pitch helical strands and little, if any, contact along them (see ref. [58], and references therein). More recent higher-resolution reconstructions [26,57,59] displayed strong contact along the two strands. According to the intersubunit connectivity pattern at the 50% mass level, the 3D reconstruction of negatively stained F-actin filaments appears more similar to the 2.5 nm resolution electron density map in displaying prominent long-pitch helix contact (fig. 12, lower panels). The overall size and shape of the 3D reconstruction of negatively stained F-actin filaments appears remarkably similar to the electron density maps at 3.0 and 3.5 nm resolution. In the 100% mass representation (fig. 12, upper panels), the cleft demarcating the two domains of the actin molecule (indicated for one subunit by a line), and the intersubunit contact area between the two strands (see arrows) appears to be more similar to the electron density map at 3.0 nm resolution. Likewise, the intersubunit connectivity pattern of our reconstruction revealed in the 50% mass representation is most similar to the electron density map at 2.5 nm resolution. As shown in fig. 13, subunits of the 3D reconstruction cut-out from negatively stained F-actin filaments closely envelope the actin molecule (see fig. 13a), most evident with a space-filling representation of the actin molecule as shown in fig. 13b. However, there are slight differences in the region of the DNase I binding loop in subdomain

107

2 of the molecule (marked by an arrow). This difference has been interpreted to indicate either significant flexibility in this region of the molecule or a different conformation of the DNase I binding loop in the F-actin filament compared to the a c t i n - D N a s e I complex [49]. Recent refinement of the atomic model of the actin filament has now borne out the latter assumption [K.C. Holmes, personal communication].

9. Some applications of negative stain The integration of biochemical and structural data from sources other than EM often requires the localization of subunits in ordered arrays and the determination of their orientation, the localization of binding sites for other proteins or ligands, or the mapping of specific amino acid residues. The most common way of localizing subunits in ordered arrays consisting of more than one type of subunit is by computing difference maps in the presence and absence of stoichiometric amounts of one or more of the subunits (e.g. refs. [57,60,61]). Substoichiometric binding of ligands and binding proteins is often more difficult to analyze and may require the use of multivariate statistical analysis [62]. A versatile approach to mapping subunits, surface-exposed cofactors and specific amino acid sequences is the use of monospecific antibodies or antibody fragments to label the sites of interest [60,63-65]. This is exemplified in figs. 14 and 15. Proteasomes isolated from Thermoplasma acidophilum have previously been shown to consist of a- and /3-subunits in a 1 : 1 stoichiometry [66]. In fig. 14, such proteasomes have been reacted with antibodies directed against the a-subunit (fig. 14a) and the /3-subunit (fig. 14b). The complexes formed with antibodies directed against the a-subunit mostly join the proteasomes in an end-to-end fashion while with anti-/3-subunit antibodies primarily side-by-side aggregates were observed [67]. This is particularly evident in lowpass-filtered images that are displayed in the insets of figs. 14a and 14b. As shown in fig. 14c, the structure of the Thermoplasma proteasome is barrel-like composed of four discs [68]. The la-

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A. Bremer et al. / Negatice staining

belling experiments described above permitted proposition of a model for the T h e r r n o p l a s m a proteasome that consists of two outer a-subunit and two inner /3-subunit discs (fig. 14d, see also ref. [67]). In fig. 15, (a) depicts a negatively stained unlabelled control of a polyhead, a polymorphic variant of the bacteriophage T4 capsid. Figs. 15b and 15c show polyheads labelled with two different monovalent antibody Fab fragments directed against distinct epitopes of the major phage T4 capsid protein gp23* [64]. The top panels in figs. 15d-15f represent Fourier-filtered images of the micrographs shown in figs. 15a-15c. The middle panels reveal the same filtrations but with a single contour of the filtration in fig. 15d superimposed. The lower panels show difference maps relative to fig. 15d with a contour superimposed as in the middle panel. As can be seen from the difference map in fig. 15e, the Fab fragments used in fig. 15b bind to two distinct epitopcs, the first one located on the six petals of the hexameric capsomer and a second one in the stainpenetrated center of the capsomer. In contrast, as shown in fig. 15f, the label used in fig. 15c only binds to the central part of the capsomer. Sequence-specific antibodies directed against surface-exposed residues can be generated by epitope scanning. This strategy has recently been used to map specific sequences in proteasomes [Grziwa et al., submitted] and, in conjunction with EM, may prove to be a powerful tool to locate specific surface-exposed amino acid sequences on a protein molecule. Alternatively, specific amino acids, particularly cysteins [57], or cofactors, can be labelled with small, electrondense metal clusters, e.g., undecagold (ref. [69], and references therein). Undecagold clusters were designed to be imaged with unstained specimens in a STEM or in frozen-hydrated preparations. However, metal clusters can also be m a p p e d with lightly stained specimens [J. Hainfield, personal communication]. 10. Conclusions

For proteins with M r between 50 and 200-300 kDa, X-ray crystallography will certainly continue

to be the method of choice for the determination of their 3D structure at near-atomic resolution. N M R may soon take over the leading role in structural analysis of smaller proteins (M~ _< 1050 kDa) in solution. Cryomicroscopy and electron crystallography of sugar-embedded or frozen-hydrated specimens will probably become increasingly important in structural biology if high resolution is required, possibly in combination with energy filtering (see ref. [70]). Progress in instrumentation (cryo-holders, automatic data acquisition procedures, recording devices, image restoration; see ref. [71]) may, however, be required to exploit the full potential of cryomicroscopy of both sugar-embedded and frozen-hydrated samples. EM can provide structural information about large protein complexes such as ribosomes (reviewed in rcf. {72]), proteasomes (e.g., ref. [68]) or the nuclear pore complex {73]. Large protein assemblies such as whole viruses or oligomeric enzymes may require combining EM with X-ray crystallography a n d / o r NMR. A recent example for a fruitful complementation of all three methods has been the structural analysis of the dihydrolipoyl transacetylase [74]. By X-ray crystallography, a rigid core was solved to atomic resolution [75]. N M R has provided structural information about the flexible parts of the molecule [76,77]. Electron microscopy could finally provide the constraints required to integrate this information into a detailed model of the molecule [74]. Negative staining has proven its potential to preserve protein structures to resolutions of up to about 1.0 nm. It is an elegant, simple and very rapid technique. In the few cases where comparable data from X-ray crystallography or frozen-hydrated specimens are available, the preservation of structural detail at the level of 1.5-3.5 nm resolution has been remarkable. Therefore, negative staining will certainly continue to provide 2D and 3D structures in this range of resolution that is sufficient to reveal important information about (i) the overall size and shape of proteins, (ii) their orientation in macromolecular assemblies, and (iii) their interaction with other proteins or molecules.

A. Bremer et al. / Negatil,e staining

Acknowledgements Drs. Wolfgang Kabsch and Ken Holmes kindly provided the coordinates of the atomic structure of the actin molecule as well as the atomic coordinates of their F-actin filament model. The approved team of Ms. H. Frefel, Ms. M. Steiner and Ms. M. Zoller provided high-quality photographic work very rapidly. This work was supported by the M.E. Mtiller Foundation of Switzerland, grants of the Swiss National Science Foundation (31-30129.90 to U. Aebi, 31-25684.88 to A. Engel), and a fellowship of the Studienstiftung des Deutschen Volkes to A. Bremer.

References [1] M.F. Perutz. Science 140 (1963) 863. [2] J.C. Kendrew, Science 139 11963) 1259. [3] K.M. Flaherty, D.B. McKay, W. Kabsch and K.C. Holmes, Proc. Natl. Acad. Sci. USA 88 11991) 5041. [4] A. Engel, Annual Rev. Biophys. Biophys. Chem. 20 (1991) 79. [5] A.T. Brfinger, Curr. Opin. Struct. Biol. 1 (1991) 1016. [6] N.E. MacKenzie, P.R. Gooley and L.A. Hardway, Annual Rev. Physiol. 54 (1992) 749. [7] B.M. Phipps, A. Hoffmann, K.O. Stetter and W. Baumeister, EMBO J. 10 (1991) 1711. [8] R. Henderson, J.M. Baldwin, T.A. Ceska, F. Zemlin, E. Beckmann and K. Downing, J. Mol. Biol. 213 (1990) 899. [9] J. Kistler and E. Kellenberger, J. Ultrastruct. Res. 59 11977) 76. [10] R.H. Haschemeyer and R.J. Myers, Negative Staining, in: Principles and Techniques of Electron Microscopy, Vol. 2, Ed. M.A. Hayat (Van Nostrand, New York, 1972) pp. 101-147. [11] R. Henderson and P.N.T. Unwin, Nature (London) 257 (1975) 28. [12] B.K. Jap, K.H. Downing and P.J. Walian, J. Struct. Biol. 103 (1990) 57. [13] W.U. Kiihlbrandt and P.N.T. Unwin, J. Mol. Biol. 156 (1982) 431. [14] R. Rachel, U. Jakubowski, R. Tietz, R. Hegerl and W. Baumeister, Ultramicroscopy 10 (1986) 305. [15] U. Jakubowski, R. Hegerl, H. Formanek, S. Volker, U. Santarius and W. Baumeister, Inst. Phys. Conf. Ser. 93, Vol. 3 (1988) 381. [16] G. Lembcke and F. Zemlin, Proc. 12th Int. Congr. Electr. Microsc. (San Francisco Press, San Francisco, 1991) pp. 102-103.

109

[17] W. Baumeister and H. Engelhardt, in: Electron Microscopy of Proteins, Vol. 6, Membraneous Structures, Eds. J.R. Harris and R.W. Horne (Academic Press, London, 1987)pp. 109-154. [18] S.C. Hfirrison, Curr. Opin. Struct. Biol. 2 11992) 293. [19] U. Aebi, W.E. Fowler, E.L. Buhle, Jr. and P.R. Smith, J. Ultrastruct. Res. 88 (1984) 143. [20] R.D. Kornberg and S.A. Darst, Curr. Opin. Struct. Biol. 1 (1991) 642. [21] W. Kiihlbrandt, Q. Rev. Biophys. 25 (1992) 1. [22] B.K. Jap, M. Zulauf, T. Scheybani, A. Hefti, W. Baumeister, U. Aebi and A. Engel, Ultramicroscopy 46 11992) 45. [23] J. Frank, Q. Rev. Biophys. 23 (19911) 281. [24] G. Lembcke, R. Diirr, R. Hegerl and W. Baumeister, J. Microscopy 161 (1991) 263. [25] C.A. Manella, Science 224 (1984) 165. [26] A. Bremer, R.C. Millonig, R. Sfinerlin, A. Engel, T.D. Pollard and U. Aebi, J. Cell Biol. 115 (1991) 689. [27] R.S. Crowther and U.B. Sleytr, J. Ultrastruct. Res. 58 11977) 41. [28] W.O. Saxton and W. Baumeister, J. Microscopy 127 (1982) 127. [29] R. Diirr, Ultramicroscopy 38 (1991) 135. [30] A.C. Steven, R. Stall, P.M. Steinert and L.TR. Benes, in: Proc. 43rd Annual EMSA Meeting, Ed. G.W. Bailey (San Francisco Press, San Francisco, 1985) p. 738. [31] E.H. Egelman, UItramicroscopy 19 (1986) 367. [32] P.A. Bullough and P.A. Tulloch, J. Mol. Biol. 215 (1990) 161. [33] F. Zemlin, Curr. Opin. Struct. Biol. 1 11991) 1023. [34] R.K.L. Bijlenga, U. Aebi and E. Kellenberger, J. Mol. Biol. 103 (1976) 469. [35] A. Brisson and P.N.T. Unwin, J. Cell Biol. 99 (1984) 12O2. [36] U. Aehi, W.E. Fowler, G. Isenberg, T.D. Pollard and P.R. Smith, J. Cell Biol. 91 (1981) 340. [37] U. Aebi, W.E. Fowler. E.L. Buhle, Jr. and P.R. Smith, E U R E M 84, Budapest, 1984, Vol. 2, pp. 1467-1480. [38] P.R. Smith, W.E. Fowler, T.D. Pollard and U. Aebi, J. Mol. Biol. 167 (1983) 641. [39] E. Zeitler, Radiation Damage in Biological EM, in: Biophysical EM, Eds. P. Hawkes and U. Valdr?~ (Academic Press, London, 1990) pp. 288-308. [40] P.N.T. Unwin, J. Mol. Biol. 87 (1974) 657. [41] W.E. Fowler, E.L. Buhle and U. Aebi, Proc. Natl. Acad. Sci. USA 81 (1984) 1669. [42] D. Suck, W. Kabsch and H.G. Mannherz, Proc. Natl. Acad. Sci. USA 78 11981) 4319. [43] U. Aebi, P.R. Smith, J. Dubochet, C. Henry and E. Kellenberger, J. Supramol. Struct. 1 (1973) 498. [44] J.C. Troncoso, J.L. March, M. H~iner and U. Aebi, J. Struct. Biol. 103 11990) 2. [45] S. Hisanga and N. Hirokawa, J. Mol. Biol. 202 (1988) 297. [46] U. Ael0i, W.E. Fowler, P. Rew and T.-T. Sun, J. Cell Biol. 97 (1983) 1131.

110

A. Bremer et al. / Negatil'e staining

[47] P.A. Janmey, S. Hvidt, G.F. Oster, J. Lamb, T.P. Stossel and J.H. Hartwig, Nature 347 (1990) 95. [48] P.A. Janmey, Curt. Opin. Cell Biol. 3 (1991) 4. [49] A. Bremer and U. Aebi, Curr. Opin. Cell Biol. 4 (1992) 20. [50] J. Lepault, I. Erk, G. Nicolas and J.-L. Ranck, J. Microscopy 161 (1991)47. [51] C.L. Woodcock and W. Baumeister, Eur. J. Cell Biol. 51 (1990) 45. [52] J. Brink, J.F.L. Van Breemen, W. Keegstra and E.F.J. Van Bruggen, Ultramicroscopy 27 (1989) 79. [53] T.S. Baker, G.E. Sosinsky, D.ED. Caspar, C. Gall and D.A. Goodenough, J. Mol. Biol. 184 (19851 81. [54] K.C. Holmes and W. Kabsch, Curr. Opin. Struct. Biol. 1 (1991) 270. [55] W. Kabsch, H.G. Mannherz, D. Suck, E. Pai and K.C. Holmes, Nature 347 (1990) 37. [56] K.C. Holmes, D. Popp, W. Gebhard and W. Kabsch, Nature 347 (19901 44. [57] R.A. Milligan, M. Whittaker and D. Safer, Nature 348 (1990) 217. [58] E.H. Egelman, J. Muscle Res. Cell Motil. 6 (19851 129. [59] U. Aebi, R.C. Millonig, H. Salvo and A. Engel, Ann. NY Acad. Sci. 483 (1986) 100. [60] U. Aebi, B. Ten Heggeler, L. Onorato, J. Kistler and M.K. Showe, Proc. Natl. Acad. Sci. USA 74 (1977) 5514. [61] R.A. Milligan and P.F. Flicker, J. Cell Biol. 105 (19871 29. [62] K.A. Stauffer, A. H6nger and A. Engel, J. Mol. Biol. 223 (1992) 1155. [63] J. Kistler, U. Aebi, L. Onorato, B. Ten Heggeler and M.K. Showe, J. Mol. Biol. 126 (1978) 571. [64] E.L. Buhle, Jr. and U. Aebi, J. Ultrastruct. Res. 89 (19841 165. [65] E. Kubalek, S. Ralston, J. Lindstrom and N. Unwin, J. Cell Biol. 105 (1987) 9. [66] B. Dahlmann, F. Kopp, L. Kuehn, B. Niedel, G. Pfeiffer, R. Hegerl and W. Baumeister, FEBS Lett. 251 (19891 125. [67] A. Grziwa, W. Baumeister, B. Dahlmann and F. Kopp, FEBS Lett. 290 (19911 186. [68] G. Pithier, S. Weinkauf, L. Bachmann, S. Miiller, A. Engel, R. Hegerl and W. Baumeister, EMBO J. 11 (19921 1607. [69] J.F. Hainfield, in: Colloidal Gold, Principles, Methods, and Applications, Vol. 2, Ed. M.A. Hayat (Academic Press, San Diego, 1989) pp. 413-431. [70] R.R. Schroeder, W. Hofmann and J.-F. Men6tret, J. Struct. Biol. 105 (1990) 28. [71] K.H. Downing, A.J. Koster and D. Typke, Ultramicroscopy 46 (1992) 189. [72] J. Frank, A. Verschoor, M. Radermacher, T. Wagenknecht, in: W.E. Hill, P.B. Moore, A. Dahlberg, D. Schlessinger, R.A. Garrett and J.R. Warner, Eds., The Ribosome: Structure, Function and Evolution (American Society for Microbiology, Washington, DC, 19991 pp. 107-113.

[73] R. Reichelt, A. Holzenburg, E.L. Buhle, Jr., M. Jarnik, A. Engel and U. Aebi, J. Cell Biol. 110 (1990) 883. [74] D. DeRosier, Nature (London) 357 (1992) 196. [75] A. Mattevi, G. Obmolova, E. Schulze, K.H. Kalk, A.H. Westphal, A. de Kok and W.G.J. Hol, Science 255 (1992) 1544. [76] R.N. Perham, Biochemistry 30 (19911 8501. [77] M.A. Robin, G.M. Clore, J.G. Omichinsky, R.N. Perham, E. Appella, K. Sakaguchi and A.M. Gronenborn, Biochemistry 31 (1992) 3463. [78] J.M. Baldwin, R. Henderson, E. Beckmann and F. Zemlin, J. Mol. Biol. 202 (1988) 585. [79] P.A. Bullough and R. Henderson, Biophys. J. 58 (1990) 705. [80] D.N. Wang and W. Kiihlbrandt, J. Mol. Biol. 217 (1991) 691. [81] W. Kfihlbrandt and D.N. Wang, Nature 350 (19911 13(t. [82] B.K. Jap, P.J. Walian and K. Gehring, Nature 350 (1991) 167. [83] H.J. Sass, G. Biildt, E. Beckmann, F. Zemlin, M. van Heel, E. Zeitler, J.P. Rosenbusch, D.L. Dorset and A. Massalski, J. Mol. Biol. 209 (1989) 171. [84] T.-W. Jeng, W. Chiu, F. Zemlin and E. Zeitler, J. Mol. Biol. 175 (1984) 93. [85] D.L. Stokes and N.M. Green, J. Mol. Biol. 213 (1990) 529. [86] J.M. Valpuesta, R. Henderson and T.G. Frey, J. Mol. Biol. 214 (1990) 237. [87] R.A. Grant, M.F. Schmid, W. Chiu, J. Mol. Biol. 217 (19911 551. [88] T.-W. Jeng, R.A. Crowther, G. Stubbs and W. Chiu, J. Mol. Biol. 205 (1989) 251. [89] A. Saito, M. Inui, M. Radermacher, J. Frank and S. Fleischer, J. Cell Biol. 107 (19881 211. [90] M. Radermacher, T. Wagenknecht, R. Grassucci, J. Frank, M. lnui, C. Chadwick and S. Fleischer, Biophys. J. 61 (1992) 936. [91] T. Wagenknecht, R. Grassucci, J. Frank, A. Saito, M. Inui and S. Fleischer, Nature 338 (1989) 167. [92] H. Cohen and W. Chiu, J. Mol. Biol. 169 (19831 235. [93] D.L. Stokes and N.M. Green, Biophys. J. 57 (1990) 1. [94] J. Brink, F.P. Booy and E.F.J. van Bruggen, Ultramicroscopy 27 (1989) 91. [95] P.N.T. Unwin and A. Klug, J. Mol. Biol. 87 (1974) 641. [96] J. Lepault, N. Martin and K. Leonard, J. Bacteriol. 168 (19861 303. [97] J. Lepault and T. Pitt, EMBO J. 3 (1984) 101. [98] G. Zampighi and P.N.T. Unwin, J. Mol. Biol. 135 (1979) 451. [99] P.N.T. Unwin and P.D. Ennis, Nature 307 (1984) 609. [100] P.N.T. Unwin and G. Zampighi, Nature, 283 (1980) 545. [101] E.F.J. van Bruggen, E.H. Wiebinga and M. Gruber, Biochim. Biophys. Acta 42 (1960) 171. [102] J.E. Mellema, E.F.J. van Bruggen and M. Gruber, Biochim. Biophys. Acta 140 (1967) 180. [103] R. Leberman, J. Mol. Biol. 13 (19651 606.

A. Bremer et al. / NegatizJe staining

[104] L.F. Estis, R.H. Haschemeyer and J.S. Wall, J. Microscopy 124 (1981) 313. [105] S. Brenner and R.W. Home, Biochim. Biophys. Acta 34 (1959) 103. [106] M.B. Sherman, E.V. Orlova, S.S. Terzyan, R. Kleine and N.A. Kiselev, Ultramicroscopy 7 (1981) 131. [107] T.M. Terry, J. Ultrastruct. Res. 41 (1972) 519. [108] R.H. Haschemeyer, in: Advances in Enzymology, Vol. 33, Ed. P.E. Nord (Wiley, New York, 1970) p. 71.

111

[109] R.M. Oliver, Methods Enzymol. 27 (1973) 616. [110] A.C. Faberge and R.M. Oliver, J. Microscopie 20 (1974) 244. [111] P.J. Shaw and G.J. Hills, J. Cell Sci. 68 (1984) 271. [112] J. Bohonek, Colloid Polym. Sci. 252 (1974) 417. [113] C.A. Manella and J. Frank, Ultramicroscopy 13 (1984) 93. [114] W. Kiihlbrandt, Ultramicroscopy 7 (1982) 221.