Methods 54 (2011) 274–283

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Methods
journal homepage: www.elsevier.com/locate/ymeth

Review Article

Imaging of nucleic acids with atomic force microscopy
Yuri L. Lyubchenko a,⇑, Luda S. Shlyakhtenko a, Toshio Ando bab

Department of Pharmaceutical Sciences, University of Nebraska Medical Center, Omaha, NE 68198-6025, USA
Department of Physics, Kanazawa University, Kakuma-machi, Kanazawa 920-1192, Japana r t i c l e

i n f oArticle history:
Available online 16 February 2011
Keywords:
Atomic force microscopy
AFM
DNA dynamics
DNA nanostructures
Holliday junctions
RNA assembly
RNA nanostructures
High-speed AFM

a b s t r a c t
Atomic force microscopy (AFM) is a key tool of nanotechnology with great importance in applications to
DNA nanotechnology and to the recently emerging ﬁeld of RNA nanotechnology. Advances in the methodology of AFM now enable reliable and reproducible imaging of DNA of various structures, topologies,
and DNA and RNA nanostructures. These advances are reviewed here with emphasis on methods utilizing
modiﬁcation of mica to prepare the surfaces enabling reliable and reproducible imaging of DNA and RNA
nanostructures. Since the AFM technology for DNA is more mature, AFM imaging of DNA is introduced inthis review to provide experience and background for the improvement of AFM imaging of RNA. Examples of imaging different structures of RNA and DNA are discussed and illustrated. Special attention is
given to the potential use of AFM to image the dynamics of nucleic acids at the nanometer scale. As such,
we review recent advances with the use of time-lapse AFM.
Ó 2011 Elsevier Inc. All rights reserved.

1. Introduction
Nucleic acid nanotechnology is a branch of nanotechnology thatutilizes the ability of nucleic acids to self-assemble into complex
three-dimensional (3D) structures. The ﬁeld of DNA nanotechnology was established in 80’s by pioneering works of N. Seeman inwhich he demonstrated the possibility to assemble 3D DNA
structures such as a cube from DNA oligomers of the well-deﬁned
structures (reviewed in [1]). The key elements for the DNA nanotechnology are three-way and four-way DNA junctions. These arebiologically important DNA structures formed during DNA replication and recombination, respectively. Unlike DNA, which exists
primarily as a double helix, RNA molecules are single stranded,
so the formation of branched RNA molecules is a rule rather than
exception. Loops in branched RNA molecule interact with eachother to form a three-dimensional biologically important RNA
structure. Unraveling principles by which such structures areformed, stabilized and transit from one into another is a problem
of great importance for molecular biology. In addition, understanding the self-assembly principles of RNA open the prospects forengineering of RNA molecules with deﬁned structural propertiesand desired functionality. This is the area of interest of the recently
emerging ﬁeld of RNA nanotechnology [2–4]. Among different
nanoimaging techniques atomic force microscopy (AFM) is thevery attractive due to its versatility in imaging of various DNA
and RNA nanostructures and the ability to directly image dynamics
of the DNA and RNA nanostructures.
Immobilization of the sample is a key step in imaging biologicalmolecules AFM. A number of sample preparation techniques weredeveloped for reliable AFM imaging of nucleic acids based primarily on their inherent negative charge [5–13]. The surface modiﬁcation methods are focused on preparing the surface to compensate
for the DNA (or RNA) charge, or forming a positively charged
substrate to hold the nucleic acids electrostatically. The mostcommon substrates for deposition of biological objects are mica
[5,6,8,10,12–17], gold ﬁlms [18–23], and glass [24–26]. Highly oriented pyrolytic graphite (HOPG) is used as well [27–31], although
its inherent high hydrophobicity complicates routine use of thissubstrate for AFM studies of nucleic acids due to their hydrophilic
nature. The sample preparation procedure depends on the imaging
type. The sections below describe various methods to prepare theAFM substrates for each type of imaging. A special emphasis is given to AFM studies of RNA nanoassembles [2]. Historically the procedures were developed for imaging of duplexes of DNA and RNA[6,14]. Later they were applied to branched DNA nanostructures(reviewed in [13,32–34]).

2. Experimental design⇑ Corresponding author. Address: Department of Pharmaceutical Sciences,
College of Pharmacy, COP 1012, University of Nebraska Medical Center, 986025
Nebraska Medical Center, Omaha, NE 68198-6025, USA. Fax: +1 402 559 9543.
E-mail address: ylyubchenko@unmc.edu (Y.L. Lyubchenko).
1046-2023/$ - see front matter Ó 2011 Elsevier Inc. All rights reserved.
doi:10.1016/j.ymeth.2011.02.001

The sample preparation methods described below were developed for DNA, but they are fully applicable for imaging of RNA
nanostructures as shown in [2,35].

Y.L. Lyubchenko et al. / Methods 54 (2011) 274–283

2.1. Mica properties and preparation as an AFM substrate
Mica is a layered mineral with potassium cations located between the hydroxyl groups of the layers [36]. The attractive featureof mica as a substrate for AFM studies is the smoothness of the surface. Mica, primarily muscovite mica, is the most widely used substrate for AFM imaging of various biological objects, nucleic acids
in particular. All commercially available mica sheets (green or rubymica) Asheville-Schoonmaker Mica Co. (Newport News, VA) arethick and large (more than 5  7 cm) sheets suitable for making
substrates of different sizes.
Mica when cleaved along the layers using either razor blades or
Scotch tape is ready for use. Freshly cleaved mica is an atomicallyﬂat surface several microns thick. After cleavage along the layers,
potassium cations are distributed on both sides, leaving an uncompensated negative charge on both surfaces of mica. Thus, freshly
cleaved negatively charged mica is a perfect AFM substrate for positively charged molecules. The adsorption of negatively charged
molecules, such as nucleic acids, requires special surface and/or
sample preparation procedures. These procedures can be divided
into two classes: treatment of mica with divalent cations [37–42]
and chemical modiﬁcation of mica [42–48]. In a few applications,mica is coated with polylysine and cationic lipids; the protocols
can be found in [43] and [11], respectively. The sections below describe the procedures for the two major methods.
2.2. Treatment of mica with divalent cations
Initially, Brack [44] suggested the pretreatment of mica with
Mg2+ cations to prepare the sample for transmission electron
microscopy. This idea was eventually implemented into the procedure for preparing the mica surface for DNA imaging [5,8].
2.2.1. Overview of the cation-assisted procedure
In this approach, mica is incubated in a solution containing 5–
10 mM Mg2+, where the resulting substrate holds DNA molecules
onto the surface, enabling reliable imaging with AFM. Later experiments have shown that pretreatment of mica with cations is notnecessary, but Mg2+ cations should be present in the depositionbuffer [5,8,10,16,45–47]. Also, later studies extended the repertoireof metal cations, so that divalent cations such as Ni2+, Mg2+, Co2+ or
Zn2+ could be used to hold the DNA molecules on the surface for
imaging with AFM [37,40], and the addition of a monovalent salt
to the deposition buffer would release the DNA molecules fromthe surface [48]. The optimum cation concentration is 1 mM,
with no DNA bound at 0.1 mM and 50% of the maximal DNA
binding at 10 mM. The DNA binding efﬁciency depends not only
on the concentrations of the metal cations,but also on their radii0[37]. When the ionic radius is 0.74 ÅA or less, DNA binds tightly tothe mica and can be imaged with AFM directly in solution. Increasing the ionic radii of the cations weakens the DNA binding. Despite
the wide use of the technique (e.g., [10,12,16,37,40,45–47]), themechanism of DNA adsorption onto a mica surface in the presenceof divalent cations is not entirely clear [48–51]. The counterions
decrease the electrostatic repulsion between the DNA and mica
surface and may also function as bridges that keep the negatively
charged DNA near the mica surface. The distribution and balanceof counterions near each of these surfaces are also driving forcesof the adsorption process [48–52].
The application of this procedure for imaging of the RNA nanostructures is described in Section 3.3.
2.2.2. Methodology for the cation-assisted procedure
The protocol for the sample preparation is straightforward. Below is the protocol provided in [37] in which various DNA samples
were imaged with AFM:

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(1) Cleave mica immediately before use.
(2) Prepare DNA solution 1–20 ng/lL (depends on the DNA type
and the length) in buffer containing 40 mM HEPES buffer(pH 7.0) and 10 mM MgCl2.
(3) Place 1 lL of the DNA solution on the mica surface, incubate
for 1–3 min, and rinse with 2–4 mL of deionized water.
(4) Dry the specimens under compressed air and further dry in a
desiccator over P2O5. Image with the Tapping mode AFM
method.
2.2.3. Major features of the cation-assisted procedure and potential
pitfalls
The simplicity of the preparation of the DNA sample is the most
attractive feature of cation-assisted technique. The surface remains
smooth, enabling high-resolution AFM studies. At the same time, anumber of issues with this method need to be taken into consideration. The major one is the change of DNA structure and morphology by divalent cations. For example, Zn2+ induces DNA kinking[53,54]. DNA remains kinked in mixed Mg2+/Zn2+ electrolytes until
the Zn2+ concentration is reduced below 100 lM. Replacing one
type of cation with another usually alleviates the problem. Mg2+
cations are used preferentially, although for imaging protein–
DNA complexes, Ni2+ cations are often used. Regardless of the divalent cation used for the deposition, DNA appears stiffer in theimages than it should be in solution with the persistence length
larger than in solution [61–67]. In addition, sequence can affectthe ﬂexibility of the DNA. This effect should be taken into account
if the DNA bending is analyzed at scales below 100 nm [55]. Finally, cation-assisted techniques complicate AFM imaging of supercoiled DNA. These are circular DNA molecules with plectonemic
geometry due to uncompensated overall twist. The overall geometry of supercoiled DNA is rather labile and sensitive to environmental conditions. In initial AFM experiments, with the use of
the Mg-assisted approach [5,8], supercoiled DNA molecules appeared on the images as fully relaxed circular molecules. Further
development of the Mg-assisted method made it possible to visualize plectonemic superhelices, but the sizes of the supercoiled
loops widely varied. In addition, large loops and tightly twisted
segments of the plectonemic superhelix appeared between theloops (reviewed in [56]). Such DNA morphology is in contrast with
the images of supercoiled DNA obtained with electron microscopy,
along with numerous physical chemical data and computer modeling (paper [57] and references therein). The problem was eased inlater experiments in which a modiﬁed procedure was used. In
these experiments, DNA was deposited onto freshly cleaved mica
from the solution containing divalent cations. This methodology
enabled to preserve the plectonemic morphology of supercoiled
DNA molecules [58]. Thus, a number of potential problems in theprocedure for DNA preparation can interfere with the quality of
AFM imaging and should be taken into account.
2.3. Chemical modiﬁcation of mica
Alternative approaches chemically modify mica with silanes
that controllably change the surface charge to yield a positively
charged mica surface [6,13,14,59]. Two major techniques are described in the sections below. Both methods are applied to AFM
imaging of DNA and RNA including DNA [32,33] and RNA nanostructures [60].
2.3.1. Chemical modiﬁcation of mica with APTESTreatment of mica with 3-aminopropyltriethoxy silane (APTES),
termed AP-mica, was developed simultaneously with the Mg-type
methods for imaging DNA [6,14] and was applied later to various
protein–DNA complexes [6,14,59,61]. The reaction of APTES isshown schematically in Fig. 1. The reaction leads to covalent276

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Fig. 1. Scheme of mica reaction with 3-aminopropopyltriethoxy silane (APTES).

attachment of aminopropyl groups that are positively charged dueto the protonation of amino groups.
2.3.1.1. Methodology for the AP-mica procedure. The vapor deposition procedure prepares smooth mica surfaces with a roughness
similar to untreated mica. The procedure has been described in detail [13]. Brieﬂy:
 Freshly cleaved mica sheets are mounted on top of a 3L glassdesiccator.
 A plastic container with 30 lL of distilled APTES is placed at thebottom of the desiccator.
 The lid is closed to allow the chemical to evaporate, yielding APmica in 2 h.
 The container with APTES is removed and the desiccator is ﬁlledwith Ar gas.
 AP-mica retains its DNA binding activity for 2 weeks with noaccumulation of visible contamination.
 The procedure produces a surface with uniform distribution of
DNA molecules in no preferred orientation.
2.3.1.2. Major features of the AP-mica procedure and drawbacks.
 AP-mica remains protonated (positively charged) over a broad
range of pH values up to pH 10. This important feature provides
an enormous ﬂexibility in using APTES-treated surfaces.
 The sample can be deposited on the AP-mica surface in a broad
range of pH and ionic strengths, without the presence of divalent cations.
 The samples prepared on AP-mica are stable over months without any accumulation of impurities. Thus, a large number ofimages can be acquired without fear of sample deterioration.
 The deposition onto AP-mica preserves the plectonemic geometry of supercoiled DNA.
This last important feature of AP-mica is illustrated by Fig. 2,
which shows AFM images of supercoiled DNA. The plectonemic

morphology of the DNA molecules is clearly seen on all moleculesin the ﬁeld. Moreover, the sensitivity of the geometry of supercoiled DNA to ionic strength was tested in the AP-mica approach[15,62]. DNA binds to AP-mica primarily due to electrostatic interactions between the protonated amino groups of the AP-mica substrate and the negatively charged DNA backbone. Also note thatthe geometry of immobilized supercoiled DNA molecules resembles two-dimensional projections of unperturbed DNA molecules
onto a plane because of the low surface charge density of AP-mica.
The latter feature suggests that the AP-mica technique is appropriate for studying global DNA conformations [63].
This approach has a drawback. During time-lapse AFM imaging
in aqueous solution, rather large aggregates appeared on the surface, presumably due to the hydrolysis of APTES molecules andtheir fast aggregation. These observations suggest that in addition
to covalent binding of APTES, molecules adsorb loosely to the surface and form aggregates after hydrolysis [64].
2.3.2. Mica functionalization with 1-(3-aminopropyl)silatrane – APSmica method
As a useful alternative to APTES, 1-(3-aminopropyl)silatrane(APS) alleviates the problems outlined above [17,32,33]. Similar
to APTES, this reagent binds covalently to hydroxyl groups of silicon surfaces (Fig. 3).
2.3.2.1. Methodology for the APS-mica procedure.
 Prepare a 50 mM APS stock solution in water and store it in arefrigerator. The shelf life of the stock solution is greater than
6 months.
 Prepare a working APS solution for mica modiﬁcation, dissolving the stock solution at 1:300 in water; it can be stored at room
temperature for several days.
 Cleave mica sheets of needed sizes (typically 1  3 cm) at 0.05–
0.1 mm thickness, place them in appropriate plastic tubes, pourworking APS solution to cover the sheets, and leave them on thebench for 30 min.
 Remove the mica sheets, wash with deionized water and drythem under an argon stream. The strips are ready for the sample
preparation. The APS mica sheets remain active for several days.

2.3.2.2. Major features of the APS-mica procedure. Unlike APTES,
which is commonly used for silica surface modiﬁcations, APS is lessreactive and extremely resistant to polymerization at neutral pH. A

Fig. 2. AFM images of supercoiled DNA deposited onto AP-mica. After deposition onto AP-mica, the samples were rinsed with water and argon dried. The images wereacquired in air with Multimode AFM (Nanoscope III, Veeco) operating in Tapping mode.

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277

suggested mechanism of the surface modiﬁcation is shown in
Fig. 3. The ﬁnal product APS-mica (III) is obtained after partialhydrolysis of the intermediate (II) on the surface. Apparently, thestabilizing triethanolamine unit remains attached to the Si atom
in APS-mica, so the linkage to the surface is more easily hydrolyzed
than the surface functionalized with APTES. A procedure has beendeveloped for preparing APS-mica with characteristics similar tothat of AP-mica (smoothness, stability, etc.) [17,32,33].
APS is not commercially available, but can be synthesized using
standard organic chemistry lab equipment [17,32]. The preparation of APS-mica for AFM is described in detail in [32].

has been trademarked by the AFM manufacturer, Veeco. Therefore,
other manufacturers utilize this methodology under different
names such as alternating contact (AC) mode. In the contact mode,
the tip–sample distance is maintained via measuring the deﬂectionof the tip cantilever determined by the van der Waals repulsion
forces, but the detection principle for the intermediate mode is different. In AC/IC/TM-AFM, a cantilever is deliberately vibrated at afrequency close to the cantilever resonant frequency by a piezoelectric modulator with very small amplitude. As the tip approaches a surface, the van der Waals attractive force between
the tip and the sample changes both the amplitude and the phaseof the cantilever vibration. These changes are monitored by a Zservo system feedback loop to control the tip–sample distance.
The ability of the AC/IC/TM mode to gently image biological samples was the major attractive feature of this operating mode.
Although acoustic excitation is primarily used to drive the tip oscillation, magnetically coated cantilevers can be driven with an oscillating magnetic ﬁeld [53]; this useful principle is implemented in
the Agilent AFM instruments. Currently, many companies manufacture AFM instruments that are quite comparable in theircharacteristics.

2.4. Other substrates for DNA immobilization

3. AFM study of branched DNA and RNA molecules

This section provides a brief overview of techniques that use
substrates other than mica. A highly oriented pyrolytic graphite(HOPG) [27,28,30,31] and epitaxially grown gold ﬁlms [19,22,23]
can be used for AFM imaging of DNA and its complexes. Both substrates are rather inert and need additional chemical activation for
adsorption of nucleic acids. Gold substrates can be modiﬁed by organic thiols or bifunctional disulﬁdes to increase their afﬁnity. Nucleic acids can then be covalently attached to the functionalized
surface [18–23,65].
An attractive property of HOPG and gold is that they are conductive substrates and can be used for scanning tunneling microscopy (STM) studies of DNA of various sizes and nucleotide
composition [66–69]. Similar to mica, HOPG has a layered structure that can be cleaved with sticky tape. However, structural defects and the ﬁne structure in step edges of pure graphite often
resemble the expected structure of DNA and thus can be misleading during DNA imaging [70–73]. Note that HOPG is a hydrophobic
substrate, which complicates its use for imaging hydrophilic samples such as DNA. HOPG can be modiﬁed for binding DNA throughmultiple steps of chemical treatment. The ﬁnal formation of thiolgroups on the surface allows binding of mercurated DNA for imaging with a scanning probe microscope [74]. Recently, a simpliﬁed
procedure for modifying HOPG was proposed [30]. The authors
used a graphite modiﬁer composed of a hydrocarbon–peptide pairterminating with an amine group: (CH2)n-(NCRH2CO)m-NH2[30,75,76]. The application of this graphite modiﬁer to the hydrophobic HOPG surface results in the formation of a thin (0.7 nm)
monolayer of positively charged polymer on the graphite surface,which promotes DNA binding. The surface is rather smooth, allowing the authors [75] to image triple-stranded poly (dG)-poly (dG)poly (dC) molecules in addition to poly (dG)-poly (dC) duplexes.
Interestingly, single-stranded regions within these polymers weredetected as well.

Although DNA primarily exists as a linear double helical polymer, during certain stages of the cell cycle, it adopts more complex
branched conﬁgurations. For example, three-way (three-arm)junctions are the models for replicating DNA, whereas a structurewith four arms linked together at a four-way junction is formed
during DNA recombination. Structural characterization of such
DNA structures is important for understanding the molecularmechanisms of the corresponding genetic processes. Comparedto DNA, branching of RNA molecules is the rule rather than the
exception. RNA molecules are primarily single stranded, but intramolecular base pairing due to complementary short regions resultsin the formation of hairpins. Using imaging to identify hairpin regions and their interaction with each other is important for understanding the functions of these 3D RNA structures [78]. For
example, many RNAs act as ‘‘ribozymes’’ and their 3D structureis critical for this function. AFM imaging would be the method of
choice for such an analysis. Unfortunately, progress in this area isstill modest; however, the advances in AFM resolution and in sample preparation will likely spur further progress in the AFM imaging of RNA.

Fig. 3. Scheme of mica modiﬁcation by 1-(3-aminopropyl)silatrane. APS (I) reactingwith the hydroxyl groups of mica, forming a transient molecule (II), and yielding a
product (III).

2.5. AFM instrumentation
Additional improvements of AFM instrumentation were needed
for routine use of AFM in biological studies. An intermittent contact mode (IC mode) has been an efﬁcient modiﬁcation of the
instrument for AFM imaging in numerous biological applications.
Initially, this mode was termed Tapping mode [77], but the name3.1. AFM imaging of three-way DNA junctions
Successful imaging of three-way DNA junctions was performed
with AP-mica methodology [79,80]. Three-way DNA junctions
were obtained by annealing two DNA strands of different lengths.
One contained an extra self-homology section capable of forming
hairpins in the middle. Except for this palindrome region, the restof the sequence of the long single strand was fully complementary
to the second strand. After annealing, these regions formed long
duplexes with hairpins in the middle. The sample was deposited
onto AP-mica, rinsed and dried as speciﬁed for the AP-mica protocol, [13] and imaged in air. AFM images for the sample with a 27 bp
hairpin are shown in Fig. 4 (see paper [80] for more images andexperimental details). Hairpin duplexes located in the middle ofthe construct are indicated with arrows in the image. The striking
feature of these three-way junctions is a large variability of the angle between the arms. For example, molecule 1 has a very smallinterstrand angle, but molecule 2 has an angle of 180o. Such variability in the interarm angle indicates a highly dynamic propertyof three-way junctions. This variability of the angle does not278

Y.L. Lyubchenko et al. / Methods 54 (2011) 274–283

Fig. 4. AFM images of the three-way DNA junctions obtained by annealing two DNA
strands with the 27 bp hairpin structure in one of the strands indicated witharrows. The images were obtained by scanning in air using Multimode AFM(Nanoscope III, Veeco) operating in Tapping mode. The ﬁgure was reprinted frompublication [80]; copyright Ó 2000 with permission from the Oxford University
Press.

Fig. 5. AFM image of dried HJs deposited onto APS-mica in Mg-containing buffer(TNM). Unambiguously classiﬁed molecules in trans (extended) and cis conformations are labeled 1 and 2, respectively. The images were obtained by scanning in air
using Multimode AFM (Nanoscope III, Veeco) operating in Tapping mode. The ﬁgure
was reprinted from publication [85]; copyright Ó 2003 with permission from theAmerican Society for Biochemistry and Molecular Biology.

depend on the orientation of the molecule relative to the scan
direction, which rules out a possible tip orientation effect. Thisconclusion is perfectly in line with the results obtained in solution[81,82], providing additional support for the suitability of the APmica procedure for analyzing dynamic 3D DNA structures.

regime, the data were acquired at a rate of 1 frame per minute.
The time-lapse AFM imaging unambiguously showed that DNA
conformation with parallel orientation exchanging arms (initially
proposed as a model for the HJ branch migration) does not support
branch migration. Unfolding of the HJs is required for branchmigration. An alternative procedure using divalent cations was
not applicable in these studies, as divalent cations bind strongly
to the junction vertex and prevent branch migration. Thus, Mg2+
cations had to be removed in these experiments to start the branchmigration process.

3.2. AFM imaging of four-way DNA junctions
Four-way DNA junctions are the model system for the DNA
intermediates. These structures consist of four DNA duplexes attached to each other at the ends. The biological signiﬁcance of
these structures is that four-way DNA junctions are formed during
DNA recombination. Termed Holliday junctions (HJs) after R. Holliday who proposed the model for homologous DNA recombination[83]. Due to the enormous biological importance of HJs, various
methods have been applied to characterize structure of HJs. Thesestudies, including recent X-ray crystallography analysis, show thatin the presence of multivalent cations, the junction adopts an antiparallel orientation in which the four helices stack in pairs to form
two double-helical domains (reviewed in [84]). Interestingly, thestudies also revealed that enzymes only interact with folded conformations of HJs to resolve the junctions, yet these conformations
do not allow branch migration to occur. Indirect data show that afully unfolded conformation of the junction is required for branchmigration [80].
AFM imaging was able to resolve this controversy in [85]. This
study designed junctions capable of branch migration and used
time-lapse imaging in solution to directly image the branch migration process. The molecules containing HJs were gel puriﬁed fromunreacted molecules or fully migrated HJs and prepared for AFM
imaging with the APS-mica methodology. Fig. 5 shows the AFM image for a sample deposited onto APS-mica and argon-dried. In thetime-lapse experiments, the sample was imaged in solution. The
DNA solution was injected into the ﬂuid AFM cell and was imaged
immediately after the AFM tip engaged the surface. The DNA drying step was omitted in these studies. In the continuous scanning

3.3. Imaging of RNA arrays
As mentioned above, unlike DNA, RNA primarily exists as a single-stranded polynucleotide, which is capable of folding into complex branched structures, depending on the sequence.
Nanoimaging techniques, such as AFM, could provide important
information on the tertiary structure of RNA. Another area of greatinterest is the use of short DNA and RNA molecules of welldesigned sequence for self-assembly of nanostructures withvarious shapes and potential novel functions. The ﬁeld was established by the pioneering works of N. Seeman [86] and is currently
being applied for designing multifunctional nanostructures. We
focus here on a few recent works involving application of AFM
imaging to study nanoassembly of RNA molecules.
In a series of works led by Jaeger [87,88], AFM was used to demonstrate the critical importance of the loop–loop interaction (kissing loop arrangement) in the assembly of hairpin-looped RNA
molecules (tecto RNAs) into square-shaped structures. The nanosquares’ assembly is stabilized by Mg2+ cations; therefore, AFM
imaging was performed in the presence of 15 mM Mg2+ cations
and samples were prepared with the cation-assisted procedure.
Note that in addition to imaging dry sample, imaging in 15 mM
Mg acetate solution was used to observe the assembly process[88]. The sample was placed on the mica surface, and the formationof nanostructures via kissing-loop formation was observed by slow

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cooling from 50 to 4 °C. The structures appeared on the AFM
images during repetitive scanning in the buffer.
Another interesting application of the RNA-based nanodesign isa series of works by P. Guo [78,89,90]. His group found that a small
viral procapsid RNA (pRNA) plays an essential role in packaging
viral DNA into the procapsid of the bacteriophage phi29. The
DNA-packaging motor of phi29 contains six copies of pRNA molecules that together form a hexameric ring. It binds to the connector
of procapsids (the unique site on procapsids where DNA enters andexits) as the ﬁrst step in DNA packaging. A loop/loop interaction ofpRNA molecules forms the hexameric ring, which contains two regions: a helical region and a self-folded region, including right andleft loops as described in [90]. The interactions between pRNA molecules can result in the formation of not only a hexameric ring, butalso other oligomers like dimers or trimers.
The interactions of these pRNA molecules were studied with acryo-AFM instrument (operates under cryogenic temperatures,
80–90 K), which was developed in the lab of Z. Shao [91]. The
major feature of cryo-AFM is an improved resolution due to theelimination of capillary forces [91,92]. The samples of pRNA forcryo-AFM were prepared by a direct deposition of solution of
RNA molecules. The samples were prepared with the use either
Mg-assisted procedure and freshly cleaved mica or mica pretreatedby spermidine. All unadsorbed molecules were removed by buffer
rinse and then by water [90]. To avoid dissociation of RNA associates, rinsing samples with water was performed quickly (<1 s).
Experimental details for the use of the cryo-AFM instrumentation
are described in [91,92].
AFM imaging of pRNA molecules [90] conﬁrms that pRNA canform a variety of structures and shapes, including dimers, tetramers, rods, triangles, and 3D arrays via interaction of programmed
helical regions and loops (Fig. 6). For example, the pRNA monomers folded into a checkmark shape 16.7 ± 0.9 nm long (Fig. 6A),

279

dimers displayed a rod shape 30.2 ± 2.5 nm long and 11.6 ±1.4 nm wide (Fig. 6B), trimers exhibited a triangle shape 30.3 ±2.4 nm on each side (Fig. 6C), and arrays displayed as bundles several microns in size (Fig. 6D) [35,90]. It should be noted that pRNA
arrays form only in the presence of 1 M of monovalent ions such as
NaCl, although the buffer already contains 5 mM of divalent ions,such as MgCl2 or CaCl2 [35,90]. Such arrays are more stable andresistant to the temperature, salt concentration, and pH than linear
120-nucleotide RNA. This is due to the tight folding in the pRNA
structure and the intertwining of the RNA molecule in the arrays.
One of the limiting factors of cryo-AFM is that the sample must
be imaged just after its preparation. Prolonged storage of a sample
in the preparation box results in sample deterioration [92,93].
Note, however, that similar resolution has been achieved for analogous RNA designs recently [60]. One set of images from this paper
is shown in Fig. 7. A good correlation between expected shapes forthe constructs (plates A and B) and the AFM images (plates C–F) areseen. Note almost a coincidence between the model (plate B) and
AFM images for this construct in plate F. The In this paper theimages were obtained at room temperature using a regular AFM.
The APS-mica sample preparation procedure was used in thoseexperiments.

4. High-speed AFM imaging of DNA and its complexes
Conventional atomic force microscopes operate rather slowly.
Typically, it takes at least 30–60 s to capture an image. Several
groups developed designs with higher data acquisition speed.
The ﬁrst fast scanning AFM was made by the group of AFM coinventor Quate [94]. This design integrated self-actuation cantilevers and used cantilever arrays to achieve high scanning speed.
However, the stiffness of the active probes did not allow imaging

Fig. 6. Cryo-AFM images of pRNA monomers (A), dimers (B), trimers (C), and arrays (D). The frame sizes of images A and B are identical to the frame size of image C. The ﬁgure
was reprinted from publication [90]; copyright Ó 2004 with permission of the American Chemical Society.

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Fig. 7. Atomic force microscopy images showing hand-in-hand and foot-to-foot dimer nanoparticles of nonmodiﬁed pRNA and 2 = FC/U pRNA, respectively. Plates A and B
are the model for the constructs. Plates C–F are the AFM images for both constructs with zoomed images are shown to the left of the corresponding large scale images. The
images were acquired in air with TM AFM (Veeco). APS-mica procedure was used for the preparation of samples. The ﬁgure was reprinted from publication [60]; copyright Ówith permission of the American Chemical Society.

of soft biological systems. A breakthrough in the development of ahigh-speed AFM (HS-AFM) for biological application was madeafter implementing the short cantilever idea proposed by Hansma[95]. The initial HS-AFM design of the Hansma group provided dataacquisition at a rate of 0.5 frames/s, which was 100 times fasterthan the rate of a conventional AFM. T. Ando’s group further developed the short cantilever idea by improving the design of the scanner and by developing the fast data acquisition electronics [96].
With this new design, the Ando team could acquire images inaqueous solution with a sub-second data acquisition rate and observe the dynamics of myosin at 12.5 frames/s for a 240-nm2 areawith 100  100 pixel resolution. Later improvements led to an
instrument operating at video rate [96–98], and its implementation in other labs made it possible to observe molecular dynamics
on a millisecond time scale [99–103]. The sections below describe
the overview and performance of the instrument and the data on
imaging segmental dynamics of DNA.

so the specimen is supported at the bottom of the sample stage andinverted to the level of the cantilever probe. The angle between the
tip and the sample is achieved by the angular position of the scanner with a sample stage.
Three key factors provide the high-speed imaging capability of
the instrument [97,98]: (i) high bandwidth of the feedback control
to maintain the tip–sample distance, (ii) active damping to suppress the scanner’s mechanical vibrations, and (iii) low-invasive
techniques to minimize the tip–sample interaction force. To
achieve the fast feedback operation, almost all devices of AFM
are optimized for fast response. For more comprehensive descriptions of HS-AFM techniques, see paper [98]. An older version of
the HS-AFM apparatus is commercially available only in Japan,
but the most recent version is not yet available. We expect that
it will be commercially available world-wide within 2 years.

4.2. Performance of HS-AFM
4.1. Overview of tapping-mode HS-AFM
A scheme of the HS-AFM setup is depicted in Fig. 8. The opticalmethod is used to measure deﬂection of the cantilever (3). The
optical beam deﬂection (OBD) detector was developed according
to the idea of Schaffer et al. [104,105]. Cantilevers for fast imaging
are approximately 10-times smaller than conventional ones (i.e.,
6–10 lm long, 2 lm wide, and 90 nm thick), and the resonant frequency and the spring constant are 600 kHz-1.2 MHz in water and
0.1–0.2 N/m, respectively [106]. Small cantilevers with a resonant
frequency of 600 kHz in water are now commercially available(AC-10; Olympus, Tokyo, Japan). Because of the small size ofhigh-speed cantilevers, the optics of the OBD detector must generate a very small incident beam spot. The objective lenses (Fig. 8, labeled 9) focus the laser spot on the rear side of the cantilever (3);the reﬂected laser beam is collected and collimated with the same
objective lenses (9). The incident and reﬂected light beams are separated using a polarization beam splitter (12) and a quarter-wave
plate (11). Then, they are reﬂected onto the split photodiode(15). The cantilever (3) is attached to a transparent glass plate
via a short cantilever holder glued to the glass plate. The rear sideof the cantilever faces downward and the tip faces upward. The
scanner (1) with a sample stage (2) is mounted facing downward,

The highest possible imaging rate of the most recent version of
the HS-AFM has reached 25 frames/s at a 250  250 nm2 scan
range, 100 scan lines, and 0.1/nm spatial frequency of sample
topography. Importantly, this high imaging rate is compatible with
low-invasive imaging. That is, the tip–sample interaction does not
disturb weak protein–protein and protein–DNA interactions. The
mechanical quantity that affects the sample is not the force itself
but the force impulse, i.e., the product of force and the time overwhich the force acts. In tapping-mode HS-AFM, the duration of
the force action is short (100 ns), and therefore, a relatively large
peak force (30 pN) would not affect the sample signiﬁcantly.
At a high imaging rate, the sample stage is moved quickly in thez-direction to maintain the cantilever oscillation amplitude (andhence the tip–sample interaction force) constant. This quick displacement generates a large hydrodynamic pressure, which could
affect the sample as well as the dynamics of cantilever oscillation.
The effects from this hydrodynamic pressure become negligible by
the following means: First, a small sample stage 2 mm in diameter is used [107]. Second, the cantilever tip is placed at an edge region of the small sample stage so that the whole cantilever chip is
minimally overlapped with the sample stage. Third, the tip lengthis adjusted to longer than 2.5 lm.

Y.L. Lyubchenko et al. / Methods 54 (2011) 274–283

281

Fig. 8. Schematic drawing of the HS-AFM head integrated with an inverted type of optical microscope. Z-piezo scanner (1), sample stage (2), cantilever (3), counter weight (4),x-piezo scanner (5), illuminator (6), stepper motor (7), piezo for exciting cantilever (8), objective lens (9), dichroic mirror (10), quarter wave plate (11), polarization beam
splitter (12), laser diode (13), focusing lens (14), split photodiode (15), and CCD camera (16).

The above-mentioned capabilities of HS-AFM have made it possible to directly visualize dynamic processes involving protein andnucleic acid molecules in millisecond time scales [96,98,108,109].
For example, the dynamic behavior of myosin V molecules walking
along actin ﬁlaments was successfully captured on video [109]. For
this observation, partially biotinylated actin ﬁlaments were immobilized onto a planar lipid bilayer surface containing a biotin–lipid,
and myosin V molecules were nearly free on the surface. The walking velocity detected therein was very similar to that measured by
ﬂuorescence microscopy, indicating negligible effects of the tip–
sample and surface–sample interactions. The molecular movies
simultaneously showed known and unknown behaviors of functioning myosin V, leading to a comprehensive understanding of
its motor mechanism. Another example is the time-lapse imaging
of bacteriorhodopsin (bR) dynamics within native purple membranes. The proteins dynamically change their structure in response to light stimuli. Upon light illumination, part of each bR
molecule is displaced outward from the trimer center. As a result,three nearest-neighbor bR monomers, each of which belongs to adifferent adjacent trimer, are brought into contact with each other.
This transient bR–bR interaction elicits both positive and negative
cooperative effects on the decay kinetics as the initial bR recovers.
4.3. Imaging of DNA segmental dynamics with HS-AFM
Successful imaging of DNA dynamics or the dynamics of otherbiological systems requires the ability of the system to move over
the AFM substrate. At the same time, the sample should remainbound within the scanning area during imaging. Selection of an
appropriate sample preparation procedure can accommodate these
conﬂicting conditions. For example, the cation-assisted methodology with Mg2+ was used [100] to observe the dynamics of DNA incomplexes with the EcoP151 restriction enzyme. This study observed ATP-dependent translocation of the enzyme. A similar
immobilization technique was applied in [101] to image the interaction of biotinylated DNA with streptavidin. Similar approachwith the use of Ca2+ was applied to image the interaction of EcoRII
restriction enzyme with DNA [102]. The use of HS AFM made it
possible to directly image various pathways of the search mechanism employed by this protein. In a recent study [103], mica was
modiﬁed by spermidine before sample deposition. With this modiﬁcation, negatively charged biological molecules can be directly
adsorbed onto the positively charged mica surface without adding
cations. The speciﬁcs for each sample preparation procedure can befound in the papers cited above. We will specify some generalfeatures of the methods. The solution, containing the biologicalsample (2 ll), is deposited onto the surface of freshly cleaved mica
(1–2 mm in diameter) for 1–15 min. After that, the sample is rinsed
with buffer to remove unattached molecules; it is then scanned in
the imaging buffer without a drying step. Generally, the rinsing
and imaging buffers are the same.
To illustrate the power of the fast scanning AFM for studies ofnucleic acids, we brieﬂy describe the results of imaging the segmental dynamics of DNA. A solution (2 ll) of the DNA fragment(491 bp, 167 nm) in 10 mM HEPES buffer containing 5 mM Mg2+
cations was placed on the surface of freshly cleaved mica, left for
2 min, rinsed with the same buffer, and imaged with an HS-AFM
instrument (RIBM Co., Ltd.). The data acquisition rate was threeframes per second. One frame is shown in Fig. 9A. The shape of
the DNA molecule is consistent with images obtained with traditional AFM, and the curvy morphology reﬂects DNA ﬂexibility.
The DNA paths on this and other images were traced and superimposed by aligning the top left ends of the molecules. Fig. 9B–D
show the superimposing of 20, 40 and 60 frames, respectively.

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Fig. 9. (A) Single frame of an HS-AFM image obtained at three frames per second. DNA strands were traced in successive frames and the traced images were overlaid. (B) 20
frames; (C) 40 frames; (D) 60 frames.

These data demonstrate that over time, DNA undergoes substantial
segmental motion that leads to a change in the entire morphologyof the molecule.
Time-lapse imaging is the most attractive mode of AFM, as the
dynamics of biological systems can be observed with nanometerspatial resolution. The critical issue with this mode is its potential
tip dragging effect due to the inevitable tip–sample interaction.
Tapping mode decreases this effect because the tip is in contact
with the sample for only a short time. The tip–sample interaction
is further decreased by the drive amplitude of the tip oscillation,
as the energy that the tip transmits to the sample depends quadratically on this parameter [63]. The drive amplitude issue isespecially important for HS-AFM due to its elevated scanningspeed. Therefore, this instrument was designed to operate at low
drive amplitude since it operates with amplitudes an order of magnitude lower than traditional AFM. To investigate a potential dragging effect [101], the movement rates of DNA segments along thetip scanning direction and perpendicular to the HS-AFM instrument were analyzed. A difference between scanning directions
has not been found, suggesting that the lateral force applied tothe sample does not increase with increasing scan rate. Additional
studies recently analyzed the elastic bending energies and ﬂuctuations of single-molecule conformations induced by the AFM probe[103]. The ﬁndings demonstrate that the AFM tip does not contribute to the whole value of DNA thermal energy and the drag effects
are negligible.
5. Conclusions
AFM has emerged from the development stage and is currently
being applied to various molecular biology problems, which often
require direct imaging of nucleic acids. Among the various imaging
modes of AFM, time-lapse nanoimaging is the most attractive, as
biological processes can be imaged at nanometer resolution.
Instrument development combined with advances in the sample
preparation techniques has made it possible to study the molecularmechanisms of such processes as homologous DNA recombinationand chromatin dynamics. Moreover, the instrumentation and thenecessary methodology for high-speed AFM have already passed
the development stage. As a result, an instrument capable of imaging the dynamics of molecules at video rate has become available
to the biomedical community. Currently, several groups have applied the technology to the most critical biological problems andhave already made a number of important discoveries including
those in RNA nanotechnology.
Acknowledgments
We are grateful to E. Menshikov and I. Menshikova for their
help with the review preparation, Alex Lushnikov, E. Menshikov,
and Alex Portillo for providing useful suggestions, J. Mercer forediting of the manuscript and other current and former membersof the group for their contribution to works incorporated in the
manuscript. The work was supported by grants to YLL from NIH(GM 62235), NSF (EPS-0701892, PHY-0615590), DOE (DE-FG0208ER64579), NATO (SfP 983204), and the Nebraska Research Initiative (NRI).
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