Journal of Biomaterials and Nanobiotechnology, 2011, 2, 8-17
doi:10.4236/jbnb.2011.21002 Published Online January 2011 (http://www.SciRP.org/journal/jbnb)
Soluble Structure of CLIC and S100 Proteins
Investigated by Atomic Force Microscopy
Stella M. Valenzuela1*, Mark Berkahn2, Alexander Porkovich1, Thuan Huynh1, Jesse Goyette3, Donald
K. Martin1,4, Carolyn L. Geczy3
1Department of Medical and Molecular Biosciences, University of Technology Sydney, NSW, Australia; 2Microstructural Analysis
Unit, University of Technology Sydney, NSW, Australia; 3School of Medical Sciences, University of New South Wales, Sydney,
Australia; 4Fondation RTRA Nanosciences, TIMC-IMAG, Université Joseph Fourier, Grenoble, France.
[e-mail], [e-mail], [e-mail]
Received November 9th, 2010; revised December 2nd, 2010; accepted December 17th, 2010
ABSTRACT
The ability to visualise proteins in their native environment and discern information regarding stoichiometry is of criti-
cal importance when studying protein interactions and function. We have used liquid cell atomic force microscopy
(AFM) to visualise proteins in their native state in buffer and have determined their molecular volumes. The human
proteins S100A8, S100A9, S100A12 and CLIC1 were used in this investigation. The effect of oxidation on the protein
structure of CLIC1 was also investigated and we found that CLIC1 multimerisation could be discerned by AFM, which
supports similar findings by other methods. We have found good correlation between the molecular volumes measured
by AFM and the calculated volumes of the individual proteins. This method allows for the study of single soluble pro-
teins under physiological conditions and could potentially be extended to study the structure of these proteins when
located within a membrane environment.
Keywords: CLIC Proteins, S100 Proteins, Atomic Force Microscopy
1. Introduction
A major challenge facing the biological sciences today is
characterization of function(s) of numerous proteins iden-
tified in this, the post-genomic era. Integral to this task is
the need to view proteins as dynamic, highly plastic
structures, in which shape and form dictate function. This
is facilitated by spatial-temporal studies, which may ulti-
mately reveal networks and multi-ligand interactions be-
tween various biomolecules. To begin addressing such
complex studies, it is pertinent to establish simple, robust
systems for the ready recognition and imaging of discrete
proteins within their native milieu. Towards this end,
atomic force microscopy (AFM) is proving a useful tool
that allows imaging of proteins in their native environ-
ment.
AFM has the potential to provide information con-
cerning conformations of proteins that are appropriate to
their natural function because they can be imaged under
conditions that most likely reflect their physiological
counterparts. This technique also has the advantage of
requiring only small quantities of protein (as low as pi-
comolar levels, but commonly in the order of ng-mg)
and manipulation of the sample environment is rela-
tively easy to achieve in order to potentially observe
structural changes in the presence of metals, lipids, co-
factors and other proteins. Importantly, AFM opens the
way for directly observing biological samples which had
previously proven extremely difficult to image.
A number of studies have used AFM to image protein
structure and stoichiometry [1-4]. AFM imaging has also
been employed to visualize the organization of bacterial
photosynthetic membranes and has provided the first
view of a multi-component membrane, revealing the rel-
ative positions and associations of the photosynthetic
complexes [5]. This technique has also been used to pro-
vide insights into aberrant protein polymerisation into
amyloid fibrils that occurs in diseases such as Alz-
heimer’s and type II diabetes [6,7].
In the current study, we used liquid cell AFM to image
protein samples in an aqueous environment, thereby
capturing them in a native soluble structure. The proteins
used were from two distinct protein families, the S100
and CLIC family of proteins, specifically CLIC1, S100-

Soluble Structure of CLIC and S100 Proteins Investigated by Atomic Force Microscopy 9
A8, S100A9 and S100A12. These proteins share certain
similarities including their expression by cells of the
immune system (although not exclusively), their rela-
tively small sizes, 10-27 kDa, and their known associa-
tion with cell membranes (except for S100A12).
1.1. Background
The CLIC family consists of 6 proteins in humans (des-
ignated CLIC1-6), which form part of the glu-
tathione-S-transferase (GST) superfamily [8,9]. They are
highly conserved in vertebrates, with related proteins in
invertebrates [10]. They have a conserved C-terminal
240 residue module and one major transmembrane do-
main. The distribution and intracellular localisation of
these proteins is distinct for each family member [11].
All members can invoke chloride ion channel activity,
and it is proposed that the proteins themselves form ion-
conducting chan nels in membranes [8,10-19]. I n order to
perform this function, these proteins must move from
their soluble form to a transmembrane complex. It re-
mains unclear how this translocation is regulated in vivo
and to date, no transmembrane structures of these pro-
teins have been elucidated.
Structural studies on CLIC proteins have focused on
CLIC1 and CLIC4. Structures of the soluble forms were
determined by X-ray crystallography [9,10]. Soluble
CLIC1, first isolated from activated macrophage cells
[20], is a monomer and has a GST fold with a covalent
binding site for glutathione [9]. Oxidation causes a major
conformational change that is stabilized by non-covalent
dimerization [15]. Soluble CLIC4 is also a monomer and
is structurally similar to CLIC1 [10] but oxidation does
not stabilize the radical conformational change that is
observed in CLIC1 [10].
The S100 proteins are highly homologous, low mo-
lecular weight (10-14 kDa), calcium modulated proteins
belonging to the EF hand superfamily [21]. Several are
over-expressed in tumour cells and have been used as
markers for tumour classification [22]. S100 proteins are
involved in vital intra- and extracellular processes [23,24]
including; modulation of cell growth, migration and dif-
ferentiation, regulation of intracellular signal transduc-
tion/phosphorylation pathways, energy metabolism, cy-
toskeletal-membrane interactions [25,26], fatty acid trans-
port [23,27] and modulation of ion channels [28].
The three “myeloid-associated” S100 proteins S100A8,
S100A9, S100A12 (A8, A9, A12 respectively) are ex-
pressed constitutively in large amounts by neutrophils
(together they constitute approximately 45% of total
neutrophil cytoplasmic protein) and are induced in mono-
cytes/macrophages [29], endothelial cells [30], keratino-
cytes [31,32] and fibroblasts [33], by a variety of media-
tors that regulate inflammation. A8 and A9 form a non-
covalent complex known as calprotectin, which is impli-
cated in neutrophil defence, by virtue of its anti- micro-
bial activity, which is dependent on the Zn2+-binding
capacity of A9 [34]. The complex also causes apoptosis
of lymphocytes by uptake of extracellular Zn2+ [35]. The
A8/A9 complex is lipophilic and intracellularly is a ma-
jor transporter of unsaturated fatty acids and arachidonic
acid [27]. At low levels of intracellular Ca2+ typical of
resting cells, A8 and A9 are primarily located in the cy-
tosol but following elevation of [Ca2+]i generated by cell
activation, they translocate to membranes and cytoskele-
tal components such as vimentin, in neutrophils and
monocytes [26,36].
The crystal structure of these S100 proteins has been
determined [37-39]; all appear to exist in cells as their
preferred structure of either non-covalently attached ho-
mo- or heterodimers [24]. In A12, the crystal structure of
the Cu2+-bound form indicates a complex of 3 dimers [40]
and multimer A12-Zn2+ complexes are found in human
atherosclerotic plaque [41], A8 and A9 form heterocom-
plexes in the presence or absence of calcium and the qu-
atramer may be the preferred functional form [42]. The
crystal structure of the A8/A9 heterocomplex is not re-
ported. We recently described the first AFM liquid cell
images of the A8/A9 heterocomplex, along with images
of its interaction with artificial lipid membranes [43]. An
earlier AFM study of interactions between the soluble
A8/A9 complex and cytochrome b558 was performed us-
ing dried samples, imaged in air [44]. The S100 proteins
have properties similar to the CLIC proteins in terms of
their oligomerisation and interactions with lipid mem-
branes.
In the current study we successfully imaged four dis-
tinct proteins by liquid cell AFM-S100A8, S100A9,
S100A12 and CLIC1. Their dimensions determined by
AFM measurement were found to be in agreement with
calculated molecular volumes and X-ray crystallography
structural dimensions. In addition, the effect of oxidation
on the oligomeric state of the protein, CLIC1, was inves-
tigated and shown to be redox sen sitive.
2. Methods
2.1. General Reagents and Protein Production
Reagents and chemicals used were analytical grade
(Sigma, Bio-Rad). Recombinant S100A8, S100A9 and
S100A12 were produced and purified using the pGEX-
4T-1 expression system as previously described [45,46].
Recombinant CLIC1 protein was produced and purified
using the pGEX-4T-1 vector expression system as pre-
viously described [15] CLIC protein was stored in buffer

Soluble Structure of CLIC and S100 Proteins Investigated by Atomic Force Microscopy 10
containing 1 mM of the reducing agent dithiothreitol
(DTT).
2.2. Atomic Force Microscopy
All images were acquired in buffer at room temperature
using a Nanoscope IIIA MultiMode AFM equipped with
an Extender™ electronics module (Veeco Instruments,
Santa Barbara, CA, USA). An E type scanner was used,
having a maximum scan area of 12.5 µm2 and vertical
height range of 3.4 µm. The NP-S series of narrow-
legged, V-shaped, 100 µm long oxide-sharpened silicon
nitride cantilevers, with integrated tips (Veeco Instru-
ments) and a nominal spring constant, k, of 0.36 N/m
were used. The AFM was driven in ‘Tapping mode™’
(TM-AFM) at the typical cantilever resonance frequency
(near 9 kHz) in a liquid cell environment at ambient
temperature. The piezo Z range was reduced to around
500 nm whilst scanning. The scan speeds ranged from 1
to 4 Hz with high gains deployed throughout. The Mul-
tiMode AFM is isolated from acoustic noise by an isola-
tion chamber and from mechanical vibration by bungee
shock cord suspension.
For imaging, proteins were diluted in working buffer
(140 mM KCl, 10 mM Hepes, 0.1 mM CaCl2, pH 6.5) to
a final concentration of 1 mg/mL. The protein (50 µL)
was then spotted onto a circular disc of freshly cleaved
mica (ProSciTech) that had been super-glued to hydro-
phobic waxy parafilm (ProSciTech), which was in turn
super-glued to a 12 mm magnetic metal disc (which at-
taches to the sample holde r of the Multimode AFM). The
protein samples were then placed under a small Petri dish
for 15 minutes with distilled water spread around the
perimeter in order to saturate the environment and mini-
mise evaporation. The protein sample was then rinsed 3
times with 100 µL of working buffer, with a final 50 µL
of working buffer added prior to mounting the disc on
the AFM.
2.3. Oxidation Experiments
To examine the effect of hydrogen peroxide treatment on
CLIC1, a 20 mM stock solution of hydrogen peroxide
was prepared immediately before use then added to give
a final concentration of 2 mM to typically 20-50 ul of the
purified protein (2 mg/ml) in 140 mM KCl, 10 mM
Hepes, 0.1 mM CaCl2, pH 6.5 and incubated overnight at
+4˚C. After in cubation the protein wa s prepared for AFM
imaging as described above.
2.4. Image Analysis
Measurements of protein “spots” on mica substrate were
carried out using Nanoscope IIIA software (Veeco In-
struments, Santa Barbara, CA, USA). The cross-section
tool was used to measure the height (h) and the diameter
(d) at half-maximal height of the proteins. This was done
in order to compensate for the artificially induced
overestimation of the protein width. Samples with
dimensions <0.5 nm were disregarded as they were
deemed too small to represent intact protein. The Experi-
mental Molecular Volume (VExp) of the proteins was
calculated using the following formula which calculates
the volume of a spherical cap [1]:
VExp = (hπ/6)(3r2 + h2) (1)
Where h and r are the height and radius of the protein
particle, respectively.
The Molecular Volume based on molecular weight
(VCal) was calculated using the following equation:
VCal = (M/NA)(V1 + dV2) (2)
Where M is the molecular weight of the protein, NA is
Avogadro’s Number (6.022 × 1023), V1 and V2 are the
partial specific volumes of particle and water (0.74 cm3
g–1 and 1 cm3 g–1 water, respectively), and d is the extent
of protein hydration (0.4 mol H2O/mol protein) [1].
3. Results
3.1. AFM Imaging of the S100 Proteins in Buffer
Imaging of the S100 proteins was undertaken in buffer
using tapping-mode, liquid-cell AFM. The 3 proteins
were imaged individually under the same conditions at
room temperature. Images collected were from at least 3
independent preparations of the protein samples spotted
onto mica on different days. All 3 proteins gave unique
error mode images and protein heights and widths were
easily discernable from the cross section images (Figures
2-4). It was found that the populations of protein parti-
cles were heterogeneous in size. Molecular volumes were
calculated for each protein {S100A8 (n = 44); S100A9 (n
= 24); S100A12 (n = 52)}, using Equation 1 (Methods)
and a frequency distribution of volumes was produced
(see Figure 1). Molecular volumes representing large
aggregates of protein were excluded from the frequency
distribution data.
S100A8 (comprises 93 amino acids; mass, 10834.51
Da) adsorbed onto mica formed spherical cap-shaped
spots, of heterogeneous size, ranging from 1.2 to 3.5 nm
in height and 7.0 to 16.2 nm diameter (at half height).
Figure 2, panel B, is the image of a representative
S100A8 spot with dimensions measuring 8.2 nm diame-
ter (at half height) and a height of 1.5 nm. Using these
dimensions, the experimental molecular volume is 41.4
nm3. The X-ray crystal structure reported by Ishikawa et
al (2000) demonstrated that the protein in solution exists

Soluble Structure of CLIC and S100 Proteins Investigated by Atomic Force Microscopy 11
Figure 1. Frequency distributions of the experimental molecular volumes of the proteins (A)S100A8; (B)S100A9 and
(C)S100A12. The molecular volumes are determined from the dimensions taken from individual protein “spots”. In the case
of the S100A9 protein, the dimensions were taken from one lobe of the tetramer structure when it appeared in this configura-
tion and reported as the volume of each individual lobe.
as a non-covalently linked dimer [37]. Assuming that
the spot imaged represented a dimer, then the experi-
mental molecular volume for the monomer would be
20.7 nm3, similar to the calculated molecular volume of
S100A8, which is 21 nm3, calculated from its molecular
weight.
S100A9 has a molecular mass of 13241.99 Da and
comprises 114 amino acid residues. The X-ray crystal
structure [38] of soluble S100A9 carried out in the pres-
ence of Chaps detergent indicates a tetramer structure of
four S100A9 dimers. Our AFM imaging resolved these
four dimers, which came together in a distinct clover-
leaf-like structure (Figure 3, Panels (a)- (c)).
The dimensions of a single S100A9 dimer within the
tetramer structure was approximately 11.8 nm wide by
0.95 nm high, giving an experimental molecular volume
for the monomer as 26.2 nm3, which agrees favourably
with the calculated molecular volume of S100A9 mono-
mer of 25 nm3.
AFM images of S100A12 (Figure 4), also revealed a
range of spherical structures ranging in size from 0.6 to
2.53 nm in height and 9.0 to 27.5 nm in diameter. The X-
ray crystal structure of this protein, determined in 2001
by Moroz et al. [47], indicates that soluble S100A12
(mass, 10575.04 Da comprising 92 amino acids) also
exists as a homodimer. However, it also forms quatram-

Soluble Structure of CLIC and S100 Proteins Investigated by Atomic Force Microscopy 12
Figure 2. AFM imaging in liquid of S100A8 protein on mica substrate. Panel (a) AFM error mode of S100A8 protein spots at
low resolution; (b) AFM error mode of a single S100A8 protein spot; (c) AFM height mode image in 3D of S100A8 proteins;
(d) PyMOL generated ribbon representation of the dimer form of S100A8 (PDB structure-1MR8) [37], showing bound
calcium ions and transparent molecular surface; (e) Cross-section analysis of a S100A8 protein spot.
Figure 3. AFM imaging in liquid of S100A9 protein on mica substrate. Panel (a) AFM error mode of S100A9 protein spots; (b)
AFM height mode of a single tetramer structured S100A9 protein spot; (c) AFM height mode image in 3D of S100A9 protein;
(d) PyMOL generated ribbon representation of the tetramer form of S100A9 dimers (PDB structure-1IRJ) [38], showing
bound calcium ions and transparent molecular surface; (e) Cross-section analysis of a single S100A9 protein spot.

Soluble Structure of CLIC and S100 Proteins Investigated by Atomic Force Microscopy 13
Figure 4. AFM imaging in liquid of S100A12 protein on mica substrate. Panels (a) & (b) AFM error mode of S100A12 protein
spots; (c) AFM height mode image in 3D of S100A12 proteins; (d) PyMOL generated ribbon representation of the hexamer
form of S100A12 (PDB structure-1GQM) [38,48], showing bound calcium ions and transparent molecular surface; (e)
Cross-section analysis of a single S100A12 protein spot.
ers and hexomers, particularly in the presence of Zn2+
[41,48]. The hexamer structure determined by X-ray cry-
stallography, is reported to be arranged as a trimer of
dimers in a spherical assembly, with an external diameter
of about 5.5 nm with a central hole of 1 nm diameter [48],
and represented in Figure 4, Panel (d).
Figure 4, Panel (e), shows the dimensions of a repre-
sentative protein spot with dimensions of 14.7 nm di-
ameter (at half height) and height of 1.4 nm. Using these
dimensions, the experimental molecular volume would
be 120.25 nm3. Assuming a hexamer structure, the S100-
A12 monomer has a volume of 20.0 nm3; the calculated
molecular volume determined from the mass of S100A12
monomer is 20 nm3.
3.2. AFM Imaging of CLIC1 in Buffer
CLIC1 (mass, 26922.73 Da, comprising 241 amino acids)
was imaged in buffer in its reduced form or following
oxidation with hydrogen peroxide, as described in the
methods (Figure 5). The height and diameter at half-
height for the individual protein particles was measured
and molecular volumes calculated {reduced CLIC1 (n =
96); oxidised CLIC1 (n = 136)}. The population of parti-
cles was found to be heterogeneous in size. A frequency
distribution of the molecular volumes of reduced CLIC1
particles (Figure 6(a)) demonstrated that >50% had a
molecular volumes between 50-100 nm3; approximately
36% of the particles had a size of 50 nm3 (smallest parti-
cle sizes). Molecular volumes representing large aggre-
gates of protein were excluded from the frequency dis-
tribution data. Under oxidising conditions, the frequency
distribution of the protein particle sizes (Figure 6(b))
changed dramatically, with <5% having a molecular
volume of 50 nm3. Using the mass of CLIC1, the calcu-
lated molecular volume of a monomer is 51 nm3.
4. Discussion
We have shown that the calculated molecular volume of
4 distinct proteins using AFM measurements, correspond
to the volume of the proteins calculated from their re-
spective molecular weight. The images of the soluble
oligomeric structures of these proteins was also found to
correspond to structures determined by X-ray crystallog-
raphy for each of these proteins [9,15,37-39,48] and re-
ported in the protein database, confirming that AFM can
reliably interpret quaternary protein structure. Based
upon the measured height and diameter values obtained
from the AFM images, single subunit volumes could be
discerned, along with multimer configurations of the
proteins.
Because CLIC1 protein structure has been shown to be
redox sensitive [15], we also imaged soluble CLIC1 un-
der oxidising conditions. SDS-PAGE and western blot-
ting experiments also confirm that CLIC1 is a monomer
under reduced conditions and forms oligomers under
oxidising conditions (results not shown). Our current
data clearly demonstrate by independent means, the oli-
gomerisation of CLIC1 following its exposure to the
strong oxidant, H2O2. From the X-ray crystallography
studies [15] it is known that the dimer is produced as a
result of a major co nformational chan ge in the N-domain
of the protein, which includes the formation of an in-
tramolecular disulfide bond between Cys-24 and Cys-59.
This rearrangement in the N-domain results in the loss of
the -sheet found in the monomer. The dimers interact
principally through 2 hydrophobic flat sheets, consisting
of 4-helices [15]. This dimer structure may act as an
intermediary in the soluble milieu, which stabilises the

Soluble Structure of CLIC and S100 Proteins Investigated by Atomic Force Microscopy 14
Figure 5. AFM imaging in liquid of CLIC1 pro tein on mica substrate. Panel (a) AFM error mode of CLIC1 protein spots in
buffer; (b) AFM height mode image in 3D of CLIC1 proteins; (c) PyMOL generated ribbon representation and transparent
molecular surface of the monomer form of CLIC1 (PDB structure–1KOM) [9]; (d) AFM error mode of CLIC1 protein spots
in buffer following overnight oxidation with H2O2; (e) AFM height mode image in 3D of oxidised CLIC1 proteins; (f) PyMOL
generated ribbon representation and transparent molecular surface of the dimer form of CLIC1 (PDB structure–1RK4) [15].
A. Frequency distribution of molecular volumes of CLIC1 reduced protein
0
10
20
30
Molecular Volume (nm3)
B. Frequency distribution of molecular volumes of CLIC1 oxidised protein
0
10
20
30
Molecular Volume (nm3)
Figure 6. Frequency distribution of molecular volumes of CLIC1 protein particles under (A) reduced or (B) oxidised condi-
tions.

Soluble Structure of CLIC and S100 Proteins Investigated by Atomic Force Microscopy 15
protein by allowing the exposed hydrophobic faces of the
monomers to interact. However, in the presence of a
membrane, it is postulated that this hydrophobic domain
of CLIC1 may interact directly with, and lead to mem-
brane insertion of the protein [15].
We have now also established a technique for incor-
porating CLIC1 into artificial membranes by incubating
liposomes with CLIC1 under oxidising conditions (data
not shown here). Using a combination of such methods
along with the AFM imaging, it is envisaged that the
transmembrane form of CLIC1 can be deduced, along
with its oligomeric state within the membrane. Similarly,
the membrane forms of the S100 proteins can similarly
be investigated, in order to shed further light on the role
and regulation of these proteins when located at cellular
membranes.
5. Acknowledgements
JG held an Australian Postgraduate award; CG was sup-
ported by a grant from the National Health & Medical
Research Council of Australia for this work. We thank
Prof Paul M. G. Curmi, School of Physics, The Univer-
sity of New South Wales, Australia, for helpful discus-
sions.
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