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NEGATIVE STAINING
Recent Developments in Negative Staining
for Transmission Electron Microscopy
J. Robin Harris 1, David Bhella 2 and Marc Adrian 3 1. Institute of Zoology, University of Mainz, Germany
2. MRC Virology Unit, Glasgow, UK 3. Department of Biology, University of Lausanne, Switzerland
BIOGRAPHY
Robin Harris obtained his
PhD from the University
of Edinburgh, Scotland.
After holding a lectureship in physiology at the
University of St Andrews
and a readership in biology at the North East London Polytechnic,
he worked for the UK Blood Transfusion Service, at the German Cancer Research Centre
in Heidelberg and at the Max-Planck-Institute for Biochemistry in Munich. Throughout his career, he has been involved with
TEM, with emphasis upon the development
of negative staining techniques and their
application to a wide range of biological,
medical and polymer science samples.
ABSTRACT
We describe some of the negative staining
procedures that are currently available for
the TEM study of biological and physical science samples. Some emphasis is placed upon
the use of ammonium molybdate as a negative stain. The benefit to be gained from
including trehalose is also discussed, as is the
use of holey or perforated carbon support
films for air-dried specimens. Cryonegative
staining can provide the best negative stain
data, with the advantages of improved
specimen preservation in vitreous ice and
the enhanced contrast of the embedding
medium versus the biological particle. The
application of negative staining to aqueous
and organic solvent polymer solutions,
dynamic or time-resolved systems, the formation of 2D crystals and higher order
assemblies, epitope and affinity labelling,
and to nanotechnology and nanobiology
samples are all discussed.
KEYWORDS
transmission electron microscopy, negative
staining, cryonegative staining, ammonium
molybdate, uranyl acetate, trehalose, holey
carbon support film, bacteria, viruses, particles, polymers, nanobiology
INTRODUCTION
Negative staining has been a useful specimen
preparation technique for biological and medical electron microscopists for almost 50 years,
following its introduction as an established
procedure by Robert (Bob) Horne [1]. During
this period of time the technique has slowly
undergone extensive modification and
improvement, now extending well beyond the
simple use of aqueous uranyl acetate as a negative stain for samples adsorbed to a relatively
thick carbon-plastic film or to a thinner but
more fragile carbon support film. Several scientists have been responsible for this progress,
including Bob Horne, with the development
of the mica-spreading ‘negative staining-carbon film’ procedure (reviewed in [2]), which
has been useful for the preparation of twodimensional crystals of viral particles and protein molecules.
Apart from the cationic and slightly acidic
1% or 2% w/v aqueous uranyl acetate, solutions of several other anionic heavy metal-containing salts have been routinely used for negative staining (e.g. tungstate, phosphotungstate, silicotungstate, molybdate and
vanadate), usually at neutral pH. By varying
the stain concentration, different levels of
stain density can be achieved. However, it will
be found that after drying, the molybdate and
vanadate negative stains impart lower mass
density around the biological material than
uranyl acetate. The anionic negative stains
generally have a finer granularity and have
less direct (charge-dependent) interaction
with the biological material (there are, however, notable exceptions, such as when an
anionic stain produces haemocyanin dissociation). Inclusion of the disaccharide trehalose
as a biological protectant during negative
staining has been shown to be beneficial [3].
This di-glucose is known to have remarkable
properties as a protectant during the drying of
biological material and its exposure to high
temperatures and UV irradiation, possibly by
replacing or retaining bound water; this property may well be beneficial at the level of the
air-dried TEM specimen grid and when it is
exposed to complete dehydration in vacuo,
and also in the electron beam. In this short
review we will present a survey of several
recent technical developments in negative
staining, all of which have led to
routine/established procedures and have the
potential for wide application in the biomedical and physical sciences.
The following recent developments will be
discussed: 1. Negative staining in the presence
of trehalose on continuous carbon films. 2. Airdry negative staining on holey carbon support
films. 3. Cryonegative staining on holey carbon support films.
Only a few relevant applications of negative
staining will be presented here; these represent areas where negative staining is currently
useful and likely to be of increasing importance for future studies: negative staining of
aqueous and organic solvent polymer solutions and colloidal suspensions; dynamic or
time-resolved negative staining; 2D crystallization of viruses and proteins, and formation
of higher-order supramolecular assemblies
during negative staining; epitope-specific
antibody and site-specific affinity labelling
revealed by negative staining; and negative
staining of nanotechnology and nanobiology
specimens.
For technical details and protocols the
reader should see the literature and a recent
review [27]. The impact and expansion of digital image processing and 3D reconstruction of
macromolecular electron optical images at
near-to-atomic resolution continues apace,
but will not be expanded upon. Some negative staining approaches, such as freeze-fracture negative staining and the negative staining-carbon film technique, will also not be
dealt with here.
Figure 1:
Haemoglobin from the marine
annelid Nereis virens negatively
stained with 5% ammonium
molybdate with 1% trehalose
(pH 7.0) on a carbon support
film. Note the presence of intact
hexameric molecules (arrowheads) and dissociating molecules, with smaller sub-components on the background.
Scale bar = 100 nm.
A U T H O R D E TA I L S
Prof. J. Robin Harris, Institute of Zoology,
University of Mainz, D-55099 Mainz,
Germany.
Tel: +44 (0)1277 210163 (UK Home)
Email: [email protected]
Microscopy and Analysis 20(3):17-21 (UK), 2006
MICROSCOPY
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17
Figure 2a:
Micronemes from Cryptosporidium parvum negatively stained with 5% ammonium molybdate with 1% trehalose after spreading across a holey carbon support film. Note the clustering of the micronemes to the edge of
the hole. Intact micronemes appear as electron transparent rods, whereas micronemes with a damaged surface
membrane allow stain entry. Scale bar = 100 nm.
Figure 2b:
The metalloendopeptidase meprin-a negatively stained on a holey carbon support film with 5% ammonium
in 0.1% trehalose (pH 7.0). Scale bar = 100 nm.
N E G AT I V E S TA I N I N G I N T H E
PRESENCE OF TREHALOSE
A I R - D R Y N E G AT I V E S TA I N I N G
ON HOLEY CARBON SUPPORTS
With the accumulating knowledge that trehalose is uniquely beneficial for the preservation of biological materials during dehydration, cold-storage, freezing, UV irradiation,
etc., it was a natural extension to prepare negatively stained specimens in the presence of
this disaccharide for TEM study. The most satisfactory combination of negative stain with
trehalose is to use ammonium molybdate
[3,4], in this instance with a 1% w/v trehalose
and 5% w/v ammonium molybdate, neutralized with NH4OH or NaOH, with the biological
sample adsorbed to a glow-discharged continuous carbon support film. The increased concentration of ammonium molybdate, above
the more usual 2% w/v, is required because of
the reduction in net mass density due to the
presence of 1% trehalose. The immediate benefit to be gained is that the film of dried stain
and trehalose is somewhat thicker than is
often the case with stain alone, allowing the
biological material to be subjected to a
reduced flattening force. In addition, ammonium molybdate does have a tendency to
release adsorbed particles from the carbon;
these can then beneficially adopt varying random orientations within the relatively deep
stain-trehalose solution, prior to drying
(Figure 1).
Although some electron-beam sensitivity of
the stain film may be encountered, conventional electron doses at minimal conventional
beam intensity do not rapidly influence the
trehalose, unlike the situation with glucose
and sucrose that more rapidly bubble in the
electron beam. If, however, a low electrondose system is available on the TEM, it should
be used. Uranyl acetate and trehalose mixtures have been found to be workable, but the
granularity of this stain and some increased
sensitivity of the sample to the electron beam
is more apparent that with the ammonium
molybdate-trehalose combination. Other negative staining salts, such as the phosphotungstate and silicotungstate can also be used
successfully in combination with trehalose.
Although many researchers may have appreciated the fact that to study biological samples
that are suspended in a thin layer of negative
stain alone (i.e. without an underlying carbon
support) could offer technical advantages,
until fairly recently this was not established as
a routine procedure. The chance spreading of
an aqueous stain and sample film across the
corner of a grid square undoubtedly provided
18
MICROSCOPY
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a
b
c
A N A LY S I S • M AY 2 0 0 6
the first indications that this approach can succeed, but considerable reproducibility and
mechanical stability problems were usually
encountered. Significant progress came when
samples were spread across glow-discharged
holey carbon support films and stained with
negative stain and trehalose [5]. Although 5%
w/v ammonium molybdate with 1.0% w/v trehalose is acceptable, some instability in the
electron beam will be encountered unless low
electron doses are used. However, in the presFigure 3:
Comparison of conventional
negative staining and cryonegative
staining.
(a) Measles virus ribonucleoprotein
imaged in 2% w/v ammonium
molybdate adsorbed to a continuous
carbon support film.
(b) Similar sample imaged in ammonium molybdate cryonegative stain
suspended across a hole in the
carbon support. Note the superior
image detail and specimen
preservation.
Scale bars = 100 nm.
(c) 3D reconstruction of Echovirus
type 12 bound to a two-domain
fragment of its cellular receptor,
CD55, calculated at 16 Å resolution
from cryonegative stain TEM data.
Docking of crystallographic coordinates for component molecules
to the EM map produces a quasiatomic resolution model of the virusreceptor complex.
NEGATIVE STAINING
ence of 5% w/v ammonium molybdate and
0.1% w/v trehalose, the stability is superior,
particularly when the holes contain an even,
thinly spread film of sample embedded in negative stain. For negative staining across holes,
a relatively high sample concentration should
be applied to the grid (~1.0 mg ml-1) as subsequent washing to remove salts and addition of
negative stain reduces the final concentration.
The strict maintenance of sample and stain on
one side only of the grid is critical for the success of this technique.
A representative example of the successful
use of the holey carbon negative staining technique, showing isolated micronemes [6] from
the apicomplexan parasite Cryptosporidium
parvum, is given in Figure 2a. Some clustering
of the micronemes towards the edge of the
hole, within a slightly thicker film of stain, is
characteristic of this technique, which indicates the freedom of the organelles immediately prior to drying of the stain. A macromolecular example is given in Figure 2b. Here the
endopeptidase meprin-a is shown; the protein
particles, some of which form curving chains,
are freely spread in the ammonium molybdate-trehalose film.
When samples are spread across holes in the
presence of 1% w/v trehalose alone, the thin
film of dried trehalose is remarkably stable in
the electron beam [5]. This approach, which
avoids the use of negative stain, has potential
for biological, polymer science and nanotechnology samples where the inherent sample
density is greater than that of the surrounding
trehalose layer.
C R Y O N E G AT I V E S TA I N I N G
Computerised 3D image reconstruction was
initially developed as a technique for the
analysis of negatively stained biological
macromolecules [7,8]. Recently, however, the
use of negative stain has largely been superseded by cryoelectron microscopy: imaging of
unstained hydrated specimens embedded in
vitreous ice [9]. The advent of cryoelectron
microscopy combined with developments in
TEM technology, such as the field-emission
gun (FEG), has permitted microscopists to
attain close-to-atomic resolution data. Imaging of unstained vitreous specimens does present significant difficulties however, not least
that image data are very low contrast, requiring often significant levels of defocus and consequent contrast transfer function correction,
to image smaller macromolecules (300-500
kDa). Vitreous specimens are also highly susceptible to radiation damage and furthermore
require specialised and expensive electron
microscopes to achieve high resolution. The
recent development of cryonegative staining
by Adrian et al. [10] abrogates some of these
difficulties while retaining the enhanced specimen preservation obtained through imaging
material embedded in vitreous ice.
As with negative staining of biological material embedded in the presence of trehalose,
cryonegative staining requires a higher concentration of stain to attain adequate contrast, typically a solution of 16-20% w/v ammonium molybdate at neutral pH is used. Prepa-
Figure 4:
A liquid crystalline 2D array of the amphiphilic poly(dimethylsiloxane)-bpoly(ethylene oxide) diblock copolymer spread across a hole and negatively stained with 5% ammonium molybdate in 0.1% trehalose (pH
7.0). Scale bar = 100 nm.
ration of cryonegatively stained material is
performed in an essentially similar manner to
that for cryoelectron microscopy of unstained
vitrified specimens. Approximately 5 µl of
sample suspension at a concentration of 0.20.5 mg ml-1 is applied to a freshly glow-discharged or gold-sputtered holey carbon support film. The sample is then washed for 5-60
seconds (depending on sample stability) in the
stain solution. Finally the sample is blotted for
1-2 seconds to produce a thin aqueous film of
sample in stain across the holey support film,
which is allowed to thin by air drying for a further 1-2 seconds before the grid is plunged
into a bath of liquid nitrogen-cooled ethane
slush. The vitrified sample is then transferred
to the microscope and imaged under low
electron-dose conditions at liquid-nitrogen
temperatures.
Cryonegatively stained specimens have been
shown to contain 30% water [10] and many
studies have demonstrated improvements in
sample preservation when compared to conventionally negative stained material. Figure
3a shows measles virus ribonucleoprotein
imaged in 2% w/v ammonium molybdate
adsorbed to a continuous carbon support film,
compared with the same sample imaged in
a
cryonegative stain (Figure 3b), suspended
across a hole in the carbon support, clearly
demonstrating significant improvement in
preservation in the frozen-hydrated stained
material [11]. The enhanced contrast and
preservation of cryonegative stain brings considerable advantages to 3D reconstruction
studies, particularly when access to a modern
top-of-the-range electron microscope is limiting. It is routinely possible to collect data on a
120 kV cryoelectron microscope, in which
structure information can be measured out to
10 Å resolution (as determined by the presence of Thon rings in incoherently averaged
power spectra, calculated from single micrograph data sets). Several 3D reconstructions
have been published at resolutions of 12-14 Å
using this technique (Figure 3c), allowing the
fitting of crystallographic data to produce
quasi-atomic resolution models [12]
While such resolutions are occasionally
attainable by cryoelectron microscopy of
unstained vitrified specimens at 120 kV, such
data are difficult to generate, requiring optimal ice-thickness, sample and imaging conditions. The paucity of published reconstructions
from 120 kV electron microscopes at greater
than 20 Å resolution is testament to the
advantages that cryonegative staining offers
such investigations. Indeed the practical resolution limits of this technique are still subject
to debate. While it has been suggested that
cryonegative stain is limited by the grain size
of the stain and is also restricted to analysis of
the stain-excluded surface regions of the sample, it is likely that higher resolution internal
features are retained, although more weakly
imaged. Such data may be accessible by development of appropriate image processing
methods.
Cryonegative staining may also prove useful
for tomographic 3D image reconstruction. The
process of collecting a tilt series of 60-70
images from a single frozen-hydrated object
incurs significant loss of data due to radiation
damage. Furthermore, cryonegative stain
samples have recently been shown to be less
susceptible to radiation damage [13]. Combined with the advantage of enhanced con-
b
Figure 5:
Cholesterol microcrystals with attached Vibrio cholerae cytolysin (VCC) oligomers. (a) After 15 min. incubation the pore-like oligomers are attached
only at the bilayer edges of the planar cholesterol crystals. (b) Following 1h incubation the whole of the surfaces of the microcrystals are coated with
oligomers [21]. The samples were negatively stained with 2% ammonium molybdate following adsorption to a carbon support film.
MICROSCOPY
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19
trast, leading to improved alignment of the
tilt series, cryonegative stain would seem to
have much to offer in this field.
a
b
N E G AT I V E S TA I N I N G O F
P O LY M E R S A N D C O L L O I D S
Despite the fact that polymer chemists use a
variety of physical techniques to assess the size
and shape of their synthetic particles, the
exploitation of TEM negative staining has
been rather slow, even though aqueous polymer solutions behave in an essentially similar
manner to biological macromolecules and
subcellular organelles. Aqueous suspensions
of gas-filled n-butyl-2-cyanoacrylate microcapsules, termed cavisomes, have been studied using negative staining on carbon support
films [14]. In this instance the globular surface
of the cavisomes was revealed, an interpretation supported by metal shadowing. Further
application of negative staining to copolymer
particles, in both aqueous and organic solvents [15,16] showed that uranyl acetate can
be utilized as dimethylformamide, tetrahydrofurane and dimethylsulphoxide solutions,
selected for miscibility with the copolymer
solutions. Furthermore, the aqueous polymer
solutions and colloidal suspensions can also be
usefully imaged by negative staining across
holes, with ammonium molybdate and trehalose, as shown in Figure 4, or in trehalose
alone [5]. Others are gradually appreciating
the potential and simplicity of air-dry negative
staining for the study of polymers and it can
be predicted that it will be increasingly used
for polymer samples, alongside cryonegative
staining and unstained cryoelectron
microscopy.
DYNAMIC AND TIMER E S O LV E D N E G AT I V E S TA I N I N G
The use of transmission electron microscopy
for the study of slow and rapid time-dependent events has always presented considerable possibilities. With negative staining, the
drying of the thin layer of stain solution occurs
over 1-2 minutes, and this might generally be
thought to impose a minimum time period for
dynamic studies, but this is not the case under
conditions where the direct action of the negative stain or a fixative has the ability to
rapidly trap biological material in a defined
metabolic state. This latter approach has been
particularly successful for the dynamic study
of flexible molecules and myosin filaments
[17,18]. For any system where the time-dependent changes occur over a period in excess of
a few minutes, conventional negative staining on a continuous carbon support film or
across holey carbon films can be successfully
utilized. With cryonegative staining, sample
pretreatment would again generally be somewhat slow, but the possibility to treat a sample
immediately before plunge freezing, such as
by suddenly changing the pH, adding a
metabolite or drug, or exposure to a temperature change, lighting conditions or gaseous
environment, could reduce the interaction
time to the millisecond range.
There is great interest in the many peptides
that spontaneously form fibres in aqueous
20
MICROSCOPY
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Figure 6:
(a) 2D arrays of the ring-like decameric peroxiredoxin from Thermus aquaticus (courtesy of Stephen G. Mayhew) spread across a holey carbon support
film in the presence of 5% ammonium molybdate and 1% trehalose (pH 7.0).
(b) Dodecahedral supramolecular assemblies formed from the decameric erythrocyte peroxiredoxin-2 in the presence of 5% ammonium molybdate,
0.1% trehalose and 0.2% PEG (Mr 1000) (pH 6.5), when spread across a holey carbon support film.
and physiological solutions, in particular the
amyloid-b and tau peptides involved in
Alzheimer’s disease. Dynamic negative staining performed over a period of minutes, hours
and days provides a system by which peptide
oligomerization, protofibril and fibre formation, and fibre aggregation can be assessed.
When combined with studies on potentiating
compounds and drugs that inhibit fibrillogenesis this negative staining approach can immediately be readily seen to have even further
possibilities [19,20].
The time-dependent interaction of bacterial
pore-forming toxins with biomembranes and
artificial lipid systems can likewise be investigated using negative staining. We found that
over a period of a few minutes the cytolysin
from Vibro cholerae formed oligomers
attached to the bilayer edges of cholesterol
microcrystals, but over a longer period of time
(1 h) the planar surfaces of the microcrystals
also became coated with oligomers [21], as
shown in Figure 5.
2D CRYSTALLIZATION OF VIRUSES
AND PROTEINS AND FORMATION
OF MACROMOLECULAR ASSEMBLIES
Induction of 2D crystal formation by viruses
and protein molecules in the presence of
ammonium molybdate and polyethylene glycol (PEG) is the underlying formative principle
of the mica-spreading negative staining-carbon film technique [2]. Intermolecular forces
A N A LY S I S • M AY 2 0 0 6
at the fluid-air interface and in solution,
rather than at the fluid-mica interface are
considered to be of importance for the production of ordered arrays [4]. On transferring
this approach to samples spread across holey
carbon support films, it has been found that,
again, viruses and protein molecules have a
tendency to produce 2D crystals (Figure 6a)
[5]. Furthermore, the time-dependent creation of higher-order macromolecular assemblies in the staining solution spread across the
holes of holey carbon support films can also
occur (Figure 6b), indicating that the negative
staining procedure can actually be utilised to
induce experimental changes. Similarly, the
cryonegative stain procedure can incorporate
the presence of PEG to induce 2D crystallization, prior to specimen freezing [10].
N E G AT I V E S TA I N I N G A N D
IMMUNOLABELLING
The combination of negative staining with
immunolabelling has been available since the
early days of the technique, but in comparison
to pre- and postembedding immunogold
labelling of thin sectioned biological material,
it has received relatively little attention. However, by negative staining, immunogold
labelling can even reveal the location of an
internalised C. parvum microneme antigen,
but only when the microneme surface membrane is damaged [22]. With the increasing
availability of peptide sequence-specific polyFigure 7:
Decamers, didecamers and multidecamers of keyhole limpet hemocyanin
type 2 (KLH2), linked with a monoclonal
IgG specific for an epitope on the functional unit h, located at the collar edge
of the decamers (smaller arrowheads).
Decamers (larger arrowheads) are
always located at the ends of the antibody-linked molecular chains. Negatively stained with 5% ammonium
molybdate in 1% trehalose (pH 7.0)
after adsorption to a carbon support
film.
NEGATIVE STAINING
clonal and monoclonal IgGs, Fab’ fragments
and single-chain variable (scFv) cloned antibody fragments, it is possible to perform
immunolabelling at the molecular level, in an
attempt to define the location of defined and
accessible epitopes on the surface of macromolecules [23]. The macromolecular linkage
pattern induced by bivalent IgG can be particularly useful (Figure 7), but a higher level of
definition can be achieved (albeit with
greater technical difficulty) by defining the
location of bound Fab’ [24].
Site-specific affinity labelling is being
increasingly used in molecular studies. The
biotin-streptavidin system is particularly powerful, because of the high affinity between
these two reagents, and the fact that biotinylated proteins and nucleic acids can readily be
produced. His-tagged proteins can be labelled
with his-specific antibodies or with nickellinked gold probes. Labelling with small gold
probes (e.g. 2-5 nm colloidal gold, nanogold
and undecagold) can readily be performed in
combination with negative staining and cryoelectron microscopy of unstained vitrified
specimens. The density of the negative stain
needs to be minimized, to avoid masking the
small gold probes, thus sodium vanadate or a
low concentration of ammonium molybdate
should be employed.
Protein-protein interactions often can be
studied by negative staining, significantly
when one of the proteins possesses a fibrous
nature and binds a soluble protein on to its
surface. The amyloid-b peptide, composing
the fibres within Alzheimer amyloid plaques,
binds several different proteins with both saltand peptide sequence-specific affinity. Of
interest is the binding of the antioxidant
enzyme catalase to amyloid fibres, as this
could be of physiological significance in relation to combating the putative oxidant activity of the amyloid-b peptide. Figure 8 shows
the binding of human erythrocyte catalase to
fibres formed from the amyloid-b 17-32 fragment. The complimentary peptide sequences
on catalase and amyloid-b have been defined
biochemically, thus introducing the possibility
of probing structurally the interaction site of
the two proteins by TEM.
NANOSTRUCTURES
Alongside atomic force microscopy, TEM has
much to contribute within the disciplines of
nanotechnology and nanobiology. Negative
staining can often be utilized, in some
instances to complement the study of
unstained samples. Particulate chemical and
materials science samples as well as synthetic
and biological polymers and lipids (nanoparticles, nanovesicles, nanotubules and nanofibres) can all be studied by negative staining,
as long as aqueous or organic solvent suspensions or solutions are available at an appropriate concentration (i.e. overloading of an
EM grid is worse than underloading!) [25].
The pharmaceutical industry, with its wideranging emphasis on drug delivery systems
and biodegradable nanoparticles could have
much to gain from the use of TEM during
product development and quality assessment.
Figure 8:
Fibres of the amyloid beta 17-40 fragment following incubation with
human erythrocyte catalase, negatively stained with 2% uranyl acetate
[19]. The catalase molecules decorate the surface of the fibres.
Figure 9:
A biotinylated DNA nanotubule decorated with streptavidin [26]. The
sample was spread across a holey carbon support film and negatively
stained with 5% ammonium molybdate with 0.1% trehalose (pH 7.0).
An example of synthetic DNA nanotubules
incorporating a biotinylated nucleotide, and
subsequently labelled with streptavidin [26], is
given in Figure 9, prepared by the holeycarbon negative-staining technique.
12. Bhella, D. et al. The structure of echovirus type 12 bound to
a two-domain fragment of its cellular attachment protein
decay-accelerating factor (CD 55) J. Biol. Chem. 279:83258332, 2004.
13. De Carlo, S. et al. cryonegative staining reduces electronbeam sensitivity of vitrified biological particles. J. Struct.
Biol. 138:216-226, 2002.
14. Harris, J. R. et al. The structure of gas-filled n-butyl-2-cyanoacrylate (BCA) polymer particles. Micron 26:103-111, 1995.
15. Harris, J. R. et al. Application of the negative staining
technique to both aqueous and organic solvent solutions
of polymer particles. Micron 30:289-298, 1999.
16. Maskos, M. and Harris, J. R. Double-shell vesicles, strings of
vesicles and filaments found in crosslinked micellar
solutions of poly(1,2-butadiene)-block-poly(ethylene oxide)
diblock copolymers. Macromol. Rapid Commun. 22:271273, 2001.
17. Burgess, S. A. et al. Use of negative stain and single-particle
image processing to explore dynamic properties of flexible
macromolecules. J. Struct. Biol. 147:247-258, 2004.
18. Zhao, F.-Q. and Craig, R. Capturing time-resolved changes
in molecular structure by negative staining. J. Struct. Biol.
K141:43-52, 2003.
19. Harris, J. R. In vitro fibrillogenesis of the amyloid a 1-42
peptide: Cholesterol potentiation and aspirin inhibition.
Micron 33:609-626, 2002.
20. Bohrmann, B. et al. Self-assembly of b-amyloid 42 is retarded by small molecular ligands at the stage of structural
intermediates. J. Struct. Biol. 130:232-246, 2000.
21. Harris, J. R. et al. Interaction of the Vibrio cholerae cytolysin
with cholesterol, some cholesterol esters and cholesterol
derivatives: a TEM study. J. Struct. Biol. 139:122-135, 2002.
22. Petry, F. and Harris, J. R. Ultrastructure, fractionation and
biochemical analysis of Cryptosporidium parvum
sporozoites. Int. J. Parasitol. 29:1249-1260, 1999.
23. Harris, J. R. Immunonegative staining: epitope localization
on macromolecules. Methods: A companion to Methods in
Enzymology 10:234-246, 1996.
24. Tulloch, P. A. et al. Single-molecule imaging of human
insulin receptor ectodomain and its Fab complexes. J.
Struct. Biol. 125:11-18, 1999.
25. Hartgerink at al. Peptide-amphiphile nanofibers: A versatile scaffold for the preparation of self-assembling
materials. Proc. Natl. Acad. Sci. USA 99:5133-5138, 2002.
26. Mitchell, J. C. et al. Self-assembly of chiral DNA nanotubes.
J. Amer. Chem. Soc. 126:16342-16343, 2004.
27. Harris, J. R. Negative Staining of Thinly Spread Biological
Samples, in: Electron Microscopy Methods and Protocols,
Methods in Molecular Biology. Vol. 117, 2nd edition. Edited
by John Kuo, Humana Press, In Press.
CONCLUDING COMMENTS
We have reviewed recent developments in
negative staining for transmission electron
microscopy. We have tried to present a range
of newer negative staining techniques and
applications, to show that negative staining
has progressed significantly over the past
decade, thereby advancing considerably from
the initial technique [1]. We believe that the
availability of the different negative stains
and negative staining approaches offers much
to both the biological and physical sciences.
REFERENCES
1. Brenner, S. and Horne, R. W. A negative staining method
for high resolution electron microscopy of viruses. Biochim.
Biophys. Acta 34:60-71, 1959.
2. Harris, J. R. The Preparation of Negatively Stained 2 Dimensional Crystals. Microscopy and Analysis, July (UK), 13-16,
1992.
3. Harris, J. R. et al. Keyhole limpet hemocyanin (KLH): Negative Staining in the presence of trehalose. Micron 26:2533, 1995.
4. Harris, J. R. Negative Staining and Cryoelectron Microscopy:
the thin film techniques. RMS Handbook No. 35, Bios
Scientific Publishers, Oxford, UK, 1997.
5. Harris, J. R. and Scheffler, D. Routine preparation of airdried negatively stained and unstained specimens on holey
carbon support films: a review of applications. Micron
33:461-480, 2002.
6. Harris, J. R. et al. Structure of the Cryptosporidium parvum
microneme: a metabolically and osmotically labile apicomplexan organelle. Micron 34:65-78, 2003.
7. DeRosier, D. J. and Klug, A. Reconstruction of three dimensional structures from electron micrographs. Nature
217:130-134, 1968.
8. Crowther, R. A. et al. Three dimensional reconstructions of
spherical viruses by Fourier synthesis from electron micrographs. Nature 226:421-425, 1970.
9. Adrian, M. et al. Cryo-electron microscopy of viruses.
Nature 308:32-36, 1984.
10. Adrian, M. et al. Cryonegative staining. Micron 29:145-160,
1998.
11. Bhella, D. et al. Conformational flexibility in recombinant
measles virus nucleocapsids visualised by cryonegative stain
electron microscopy and real-space helical reconstruction.
J. Mol. Biol. 340:319-331, 2004.
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