Transworld Research Network
37/661 (2), Fort P.O., Trivandrum-695 023, Kerala, India

DNA Structure, Chromatin and Gene Expression, 2006: 17-26 ISBN: 81-7895-228-9
Editors: Ryoiti Kiyama and Mitsuhiro Shimizu

2

Biochemical and microscopic
analyses of DNA conformations
Mikio Kato
Department of Biological Science, Osaka Prefecture University Graduate
School of Science, 1-1 Gakuencho, Naka-ku, Sakai 599-8531, Japan

Abstract
In addition to the well-known right-handed double
helix (B-DNA), a number of alternative DNA
structures can be formed at defined, ordered DNA
sequences. For example, inverted repeats may form
DNA cruciforms, pyrimidine/purine-biased sequences
may form intra- and intermolecular DNA triplexes,
alternating purine-pyrimidine sequences may form ZDNA, and so on. These characteristic sequence tracts
occur often within the genomes. And to understandtheir function, it is necessary to understand theirpotential to form alternative DNA structures. In thispaper, we review several methods for analyzing theCorrespondence/Reprint request: Dr. Mikio Kato, Department of Biological Science, Osaka Prefecture
University Graduate School of Science, 1-1 Gakuencho, Naka-ku, Sakai 599-8531, Japan
E-mail: mkato@b.s.osakafu-u.ac.jp

18

Mikio Kato

structures of DNA and chromatin, and discuss some of the biologicalimplications of the observed DNA structures.

Introduction
In response to changes in environmental conditions, the structure of DNA
may vary in a sequence-dependent manner. Any DNA conformation otherthan the right-handed double helix (B-DNA) is classified as an “unusual DNA
structure,” and a variety of DNA sequences with the potential to form unusual
structures have been characterized [1-3]. For example, pyrimidine/purinebiased sequences (hereafter, Pyr/Pur sequences) may form inter- andintramolecular triplexes and their derivatives; inverted repeats may form DNA
cruciforms; alternating purine-pyrimidine sequences may form Z-DNA (lefthanded double helix); the occurrence of phased oligoadenine tracts causes
curvature of the DNA axis; direct repeat sequences may cause slipped
misalignment; and telomeric guanine-rich repeats may form quadruplexes.
Some of these sequences are widespread within genomes, occurring more
often than random sequences [4,5], and the unusual (alternative) DNA
structures that result are thought (or expected) to affect processes by whichgene expression is regulated by modulating global and local DNA topology orby serving as target sites for specific proteins [6-10]. In addition, some
unusual DNA structures that appear to have no effect on genome function
nevertheless may have a big impact on gene structure and expression. For
instance, in the case of triplet repeats and some other simple repeat sequences,
the numbers of tandem repeats are unstable during transmission [11].
Although the instability is, itself, not known to affect genome function,
expansion of the repeat regions may cause congenital anomalies. Models for
expansion and contraction during DNA replication have been proposed basedon unusual DNA structures that form at the template or nascent strands, andthe mechanisms underlying the diseases caused by expansion of repeat tracts
have been discussed [12,13].
In order to clarify how B-DNA transitions into unusual DNA structures, inparticular how it occurs in vivo, one needs to establish methods forcharacterizing DNA structures under defined sets of conditions. In this paper,
the current methodologies of topological fractionation using gelelectrophoresis, chemical and enzymatic probing, and microscopic imaging of
DNA and chromatin are reviewed with an emphasis on alternative DNA
structures forming at inverted repeats and Pyr/Pur sequences.

Electrophoretic mobility of supercoiled DNA
DNA’s electrophoretic mobility is strongly dependent on both the size andshape of the molecule. Larger molecules migrate slower in agarose gel, while

Unusual DNA structures

19

more compact molecules (e.g., supercoiled DNA) migrate faster. Because theformation of intramolecular triplexes results in relaxation of negative
supercoils (as does cruciform formation), two-dimensional agarose gelelectrophoresis enables one to identify topoisomers at which structuraltransitions have occurred.
E. coli plasmid DNA molecules are mixtures of topoisomers, and each
topoisomer yields an individual spot on two-dimensional gel electrophoresis[14]. In the first dimension, a set of topoisomers is fractionated underconditions in which structural transition may occur in some; in the second
dimension they are fractionated under conditions in which no structuraltransitions occur. Because intercalation of chloroquine reduces the twist angle
between neighboring base pairs, electrophoretic buffer containing chloroquine
is used in the second dimension to prevent supercoiling-dependent structuraltransition. We have used this approach to detect formation of unusual DNA
structures occurring on supercoiled DNA containing short inverted repeats[15]. The plasmid pBan1 is a pUC19 derivative containing a short inverted
repeat made up of 17-bp complementary sequences. The electrophoretic
profile of pBan1 DNA exhibiting native bacterial superhelicity is shown in Fig.
1. A structural transition that consumes about 3.5 superhelical turns occurs in

Figure 1. Two-dimensional agarose gel electrophoresis of pBan1 DNA. Plasmid
pBan1 exhibiting native bacterial superhelicity was two-dimensionally fractionated on
1.2% agarose gel. In the first dimension, standard TAE buffer was used; 6 µg/ml
chloroquine was added in the second dimension. OC represents nicked circular pBan1,
and the numbers of negative supercoils present in the samples are given at therespective spots. Structural transition has occurred in the DNA molecules having 10 ormore negative supercoils (indicated by arrow), consuming ~3.5 helical turns.
Reproduction from Kato et al. [15] with permission of the Biophysical Society.

20

Mikio Katoa small population of pBan1 DNA molecules having 10 superhelical turns(indicated by the arrow in the Fig. 1), and in up to 50% of pBan1 DNA
molecules having more than 10 superhelical turns. Thus, the negative
superhelicity required for structural transition and the numbers of superhelicalturns consumed by that transition can be identified using two-dimensional gel
electrophoresis.
Pyr/Pur sequences arranged in mirror symmetry have the potential to form
DNA triplexes that are induced and stabilized by acidic conditions [1-3].
Moreover, long (more than 100 bp) Pyr/Pur sequences, which appear frequently
in eukaryotic genomes [16,17], can form other unusual DNA structures underacidic conditions. Because the center of symmetry within long Pyr/Pur
sequences can vary, alternative DNA structures forming at long Pyr/Pur tracts
are heterogeneous, enabling triplex stems of various size and multiple triplexes
to be seen. Consistent with such heterogeneity, supercoiled DNA molecules
show a smeary mobility, rather than discrete bands (or spots), when subjected toelectrophoresis under acidic conditions [18-20]. As an example, agarose gelelectrophoresis of another pUC19 derivative, pTIR10, which contains a rat
genomic DNA fragment with a ~230-bp Pyr/Pur insert is shown in Fig. 2. Note

Figure 2. Electrophoretic profiles of plasmid pTIR10 under neutral and acidic
conditions. Left panel, electrophoresis on 1% agarose in 40 mM Tris-acetate/ 5 mM
sodium acetate/1 mM EDTA (pH7.5); right panel, electrophoresis on 1% agarose in 30
mM sodium acetate/1mM EDTA (pH 4.6). Lane 1, pUC19 DNA; lane 2, pTIR10
DNA; lane 3, pTIR10 DNA linearized by HindIII digestion; lane M, HindIII-digested
lambda phage DNA size marker. Fully supercoiled molecules, linear molecules, open
circles and supercoiled dimeric molecules of pUC19 are labeled a, b, c and d,respectively, in the left panel. The faint DNA band at the bottom of lanes 2 and 3(marked with an asterisk) might be the supercondensed structure reported previously[25, 26]. Reproduction from Kato et al. [20] with permission of FEBS.

21

Unusual DNA structures

the smeary profile produced by electrophoresis in acidic buffer, and that theinserted region is sensitive to S1 nuclease when under superhelical tension [18].
But whereas analysis of electrophoretic mobility is useful for detecting how
structural transitions affect the global topology of DNA, it is not designed toshow what type of local conformational change has occurred.

Chemical andconformation

enzymatic

probingofDNA

A number of chemical compounds that react with bases or with the sugarphosphate backbone are used to define the tertiary structures of DNA andchromatin. Because susceptibility to these chemicals is strongly dependent on
the accessibility of their target sites, the sites attacked by chemicals are thought
to be exposed to the environment. The nature of these chemical reactions andthe results of probes of several DNA structures have been described [2,3].
The Pyr/Pur sequence in the promoter region of the human epidermal
growth factor receptor gene (EGFR) can be regarded as possessing either
direct repeat or mirror symmetry. The region is sensitive to S1 nuclease [21]and, at acidic pHs, the 5’-half of the polypurine strand is susceptible to attackby diethylpyrocarbonate (Fig. 3), which mainly reacts with unstacked adenine
and less with guanine. Acidic pH makes unpaired cytosines more likely to be
protonated, thereby promoting formation of C+:G:C base triads, which isindicative of the potential of the promoter region of EGFR to form
intramolecular triplexes [22]. The role of the potential triplex region in
regulating EGFR expression is unclear, however. This region was found to beunnecessary for maximum transcriptional activity of the promoter in vivo andin vitro [23,24], but Johnson et al. [21] showed that mutations in this region
reduce promoter activity in transient expression assays.

Figure 3. Nucleotide sequence of Pyr/Pur stretch at the human EGFR promoter
region. Shown are positions -374 to -337; the numbering is according to Johnson et al.
[21]. The region can be regarded as possessing either direct repeat or mirror symmetry.
The center of symmetry is boxed, and the sites attacked by diethylpyrocarbonate areunderlined and italicized. S1 nuclease mainly cleaved the 3’ side of the boxed nucleotides(between A and T in the top strand, and between T and G in the bottom strand). The
results indicate that the region may produce triplexes occurring at the mirror symmetry
rather than DNA slippage occurring at direct repeats. See ref. 22 for details.

22

Mikio Kato

It is generally accepted that DNA containing short inverted repeats may
assume a cruciform structure [1]. On the other hand, Ohta et al. [27] and Katoet al. [28] have proposed an alternative DNA structure that is distinct but
derived from the DNA cruciform. In that structure, the two stem-loops of the
cruciform are arranged in parallel and, possibly, one stem-loop melts to yield athird strand able to form a triple-stranded structure. We tested that idea using
potassium permanganate, which efficiently attacks unpaired thymine bases[15]. We found that permanganate reacted with the thymines in the 3’-half of
the inverted repeat as well as with those at the middle of the inverted repeat(Fig. 4), which is consistent with the hairpin triplex model proposed by Kato etal. [28]. This structure is also formed in E. coli cells. Although thepermanganate reacted with DNA more efficiently under acidic conditions,the profiles of permanganate oxidation were essentially identical under bothneutral (lanes 1, 2, and 3 of Fig. 4) and acidic (lanes 4, 5, and 6 of Fig. 4)

Figure 4. Primer extension assay of permanganate-treated DNA. The products ofprimer extension were electrophoresed along with control sequencing ladders (lanes T, C,
G and A). Permanganate oxidation was carried out in vitro (panels A and B) and in situ
(panels C and D). Panels A and C show results obtained with the M13 forward primer,while panels B and D show those obtained with the M13 reverse primer. Asterisks andarrows at the left of the panels indicate the positions of the inverted repeat symmetry. A
summary of sites susceptible to permanganate oxidation (indicated by arrowheads) isshown at the bottom. Panels A and B: lanes 1 and 4, 1 min permanganate reaction; lanes 2
and 5, 2 min permanganate reaction; lanes 3 and 6, 4 min permanganate reaction. The
reactions in lanes 1, 2 and 3 were in neutral buffer; those in lanes 4, 5 and 6 were in acidic
buffer. Panels C and D: lane 1, permanganate reaction with mid-log growth phase E. coli
cells; lane 2, permanganate reaction with late-log growth phase E. coli cells. Reproduction
from Kato et al. [15] with permission of the Biophysical Society.

Unusual DNA structures23

conditions, suggesting that cytosine protonation is not involved in formation of
hairpin triplex, but parallel G:G:C and A:A:T triads may be. Such parallel
triplexes have been described previously [29,30].

Microscopic imaging of DNA and chromatin
Because biochemical measurements generally describe the average
structure of the molecules in a DNA sample, it is difficult to characterizeheterogeneous populations of molecules. For example, because long Pyr/Pur
sequences may form triplex stems of various size or cause formation ofmultiple stems in one molecule, electrophoretic profiles are smeary and mostof the potential target sites are attacked by chemicals in an uninterpretablemanner. In such cases, microscopic imaging is a powerful tool with which totake a snapshot of individual molecules.
We have used atomic force microscopy (AFM) to characterize theheterogeneity of triplex formation in pTIR10 plasmid [20]. AFM imaging
of supercoiled pTIR10 revealed the presence of various triplex stems, which
was expected based on the biochemical observations (Fig. 5). AFM has the

Figure 5. AFM images of the most representative families of irregularities foundin pTIR10 DNA. Panel a, short triplex-like stem; panel b, long curved stem; panel c,twin stems; panel d, Π-shaped structure; panel e, Ψ-shaped structure. Scale bar (100nm) is given at the bottom of panel e and is common to all panels. Potential triplex
stem regions are indicated by arrows. Structures shown in panels c, d and e are
multiple triplex formations. A model showing a conventional intra-molecular triplex isgiven at the right bottom. Reproduction with modification from Kato et al. [20] withpermission of FEBS.

24

Mikio Katoadvantage that no staining is required for observation, but there is the
disadvantage that, due to a convolution effect [31] caused by the probe tip, theapparent width of the DNA is greater than the actual size. Electron
microscopy with heavy metal staining gives sharp (high contrast) images ofDNA fiber. Plasmids pBan1 and pTIR10 visualized using electron microscopy
with uranyl acetate staining are shown in Fig. 6.

Figure 6. Electron microscopic images of alternative DNA structures. Panel A,
supercoiled DNA containing short inverted repeat (pBan1); panel B, supercoiled DNA
containing a Pyr/Pur sequence (pTIR10). The stem part is indicated in each panel.
Reproduction from Kato et al. [15] with permission of the Biophysical Society.

Conventional electron microscopy usually requires artificial fixation of
specimens followed by heavy metal staining, so the images obtained should be
evaluated cautiously. However, procedures for observing nearly native(unfixed and unstained) samples have been developed with the aim of avoiding
chance artifactual deformation of the specimen that can be caused by staining.
Bednar et al. characterized the structures of nucleosome [32] and transcribing
RNA polymerase on the chromatin template [33], and Bouchet-Marquis et al.
[34] analyzed nuclear architecture by cryo-electron microscopy. In addition,
Kaneko et al. recently used a novel type of electron microscopy, Hilbertdifferential transmission electron microscopy, to analyze the subcellular
structure of cyanobacteria in a near-living state [35,36]. These techniques will
likely prove effective for characterizing alternative DNA structures occurringin vivo.

Conclusion
Many sequence tracts within genomes have the potential to form
alternative DNA structures; the formation of such structures has been observedin vitro, and the kinetics of their structural transition have been studied. For
instance, long Pyr/Pur sequences are scattered abundantly within genomes andmay assume a variety of alternative DNA structures, including multiple

Unusual DNA structures25

intramolecular triplexes. Indeed, probing with anti-triplex DNA antibody
revealed the presence of numerous triplexes within the interphase nucleus [37].
In addition, intermolecular interaction of two intramolecular triplexes has beenreported for the (GAA/TTC)n triplet repeat in the first intron of the humanfrataxin gene, which yields sticky DNA [38], and formation of sticky DNA
directly correlates with inhibition of transcription [39]. Thus, certain Pyr/Purloci may offer triplex stems for inter- and intrachromosomal interactions tomodulate transcription and replication. That said, little is known about thefunction of alternative DNA structures within the genome. To define thephysiological role of Pyr/Pur sequences and other characteristic DNA
sequences, it will be necessary to directly observe alternative DNA structures
at work in vivo (or in situ). To that end, improved microscopic imaging
techniques may enable us to detect the in vivo dynamics of DNA andchromatin.

References
1.
2.
3.
4.
5.
6.
7.
8.
9.
10.
11.
12.
13.
14.
15.
16.
17.
18.
19.

Sinden, R. R., 1994, DNA Structure and Function. Academic Press, San Diego.
ISBN 0-12-645750-6
Wells, R. D. and Harvey, S. C. (editors), 1988, Unusual DNA Structures, SpringerVerlag, New York. ISBN 0-387-96631-5
Soyfer, V. N. and Potaman, V. N., 1995, Triple-Helical Nucleic Acids. SpringerVerlag, New York. ISBN 0-387-94495-8
Schroth, G. P., Chou, P.-J. and Ho, S., 1992, J. Biol. Chem. 267, 11846-11855.
Schroth, G. P., and Ho, S., 1995, Nucleic Acids Res. 23, 1977-1983.
Kohwi Y and Kohwi-Shigematsu, T., 1991, Genes Dev. 12, 2547-2554.
Kato, M and Shimizu, N., 1992, J. Biochem. (Tokyo) 112, 492-494.
Brukner, I., Belmaaza, A. and Chartrand, P., 1997, Proc. Natl. Acad. Sci. USA 94,
403-406.
Chen, S., Supakar, P. C., Vellanoweth, R. L., Song, C. S., Chatterjee, B. and Roy,A. K., 1997, Mol. Endocrinol. 11, 3-15.
Zain, R. and Sun, J-S., 2003, Cell. Mol. Life Sci. 60, 862-870.
Sinden, R. R., 1999, Am. J. Hum. Genet. 64, 346-353.
Pelletier, R., Krasilnikova, M. M., Samadashwily, G. M., Lahue, R. and Mirkin, S.
M., 2003, Mol. Cell. Biol. 23, 1349-1357.
Krasilnikova, M. M., Mirkin, S. M., 2004, Mol. Cell. Biol. 24, 2286-2295.
Hanai, R., and Rocca, J., 1999, Methods Mol. Biol. 94, 19-27.
Kato, M., Hokabe, S., Itakura, S., Minoshima, S., Lyubchenko, Y. L., Gurkov, T.
D., Okawara, H., Nagayama, K. and Shimizu, N., 2003, Biophys. J. 87, 402-408.
Straus, N. A. and Birnboim, H. C., 1976, Biochim. Biophys. Acta 454, 419-428.
Birnboim, H. C., 1978, J. Mol. Biol. 121, 541-549.
Kato, M., 1993, Mol. Biol. Reports 18, 183-187.
Blaszak, R. T., Potaman, V., Sinden, R. R. and Bissler, J. J., 1999, Nucleic Acids
Res. 27, 2610-2617.

26Mikio Kato

20. Kato, M., McAllister, C. J., Hokabe, S., Shimizu, N. and Lyubchenko, Y. L., 2002,
Eur. J. Biochem. 269, 3632-3636.
21. Johnson, A. C., Jinno, Y. and Merlino, G. T., 1988, Mol. Cell. Biol. 8, 4174-4184.
22. Kato, M., Kudoh, J. and Shimizu, N., 1990, Biochem. J. (London) 268, 175-180.
23. Johnson, A. C., Ishii, S., Jinno, Y., Pastan, I. and Merlino, G. T., 1988, J. Biol.
Chem. 263, 5693-5699.
24. Kageyama, R., Merlino, G. T. and Pastan, I., 1988, J. Biol. Chem. 263, 6329-6336.
25. Huang, X. and Chen, X., 1990, J. Mol. Biol. 216, 195-199.
26. Kato, M. and Furuno, A., 1992, Res. Microbiol. 143, 665-669.
27. Ohta, T., Nettikadan, S., Tokumasu, F., Ideno, H., Abe, Y., Kuroda, M., Hayashi,
H. and Takeyasu, K., 1996, Biochem. Biophys. Res. Commun. 226, 730-734.
28. Kato, M., Matsunaga, K. and Shimizu, N., 1998, Biochem. Biophys. Res. Commun.
246, 532-534.
29. Shchyolkina, A. K., Borisova, O. F., Minyat, E. E., Timofeev, E. N., Il'icheva, I.
A., Khomyakova, E. B. and Florentiev, V. L., 1995, FEBS Lett. 367, 81-84.
30. Shchyolkina, A. K, Timofeev, E. N., Lysov, Y. P., Florentiev, V. L., Jovin, T. M.
and Arndt-Jovin, D. J., 2001, Nucleic Acids Res. 29, 986-995.
31. Yang, J., Mou, J., Yuan, J. Y. and Shao, Z., 1996, J. Microsc. 182, 106-113.
32. Bednar, J., Horowitz, R. A., Grigoryev, S. A., Carruthers, L. M., Hansen, J. C.,
Koster, A. J. and Woodcock, C. L., 1998, Proc. Natl. Acad. Sci. USA 95, 1417314178.
33. Bednar, J., Studitsky, V. M., Grigoryev, S. A., Felsenfeld, G. and Woodcock, C.
L., 1999, Mol. Cell 4, 377-386.
34. Bouchet-Marquis, C., Dubochet, J. and Fakan, S., 2006, Histochem. Cell Biol. 125,
43-51.
35. Kaneko, Y., Danev, R., Nitta, K. and Nagayama K., 2005, J. Electron Microsc.
(Tokyo) 54, 79-84.
36. Kaneko, Y., Danev, R., Nagayama, K. and Nakamoto H., 2006, J. Bacteriol. 188,805-808.
37. Ohno, M., Fukagawa, T., Lee, J. S. and Ikemura, T., 2002, Chromosoma 111, 201213.
38. Sakamoto, N., Chastain, P. D., Parniewski, P., Ohshima, K., Pandolfo, M.,
Griffith, J. D. and Wells, R. D., 1999, Mol. Cell 3, 465-475.
39. Sakamoto, N., Ohshima, K., Montermini, L., Pandolfo, M. and Wells, R. D., 2001,
J. Biol. Chem. 276, 27171-27177.