Chain and conformation stability of solid-state DNA: implications for room temperature storage.
ABSTRACT There is currently wide interest in room temperature storage of dehydrated DNA. However, there is insufficient knowledge about its chemical and structural stability. Here, we show that solid-state DNA degradation is greatly affected by atmospheric water and oxygen at room temperature. In these conditions DNA can even be lost by aggregation. These are major concerns since laboratory plastic ware is not airtight. Chain-breaking rates measured between 70 degrees C and 140 degrees C seemed to follow Arrhenius' law. Extrapolation to 25 degrees C gave a degradation rate of about 1-40 cuts/10(5) nucleotides/century. However, these figures are to be taken as very tentative since they depend on the validity of the extrapolation and the positive or negative effect of contaminants, buffers or additives. Regarding the secondary structure, denaturation experiments showed that DNA secondary structure could be preserved or fully restored upon rehydration, except possibly for small fragments. Indeed, below about 500 bp, DNA fragments underwent a very slow evolution (almost suppressed in the presence of trehalose) which could end in an irreversible denaturation. Thus, this work validates using room temperature for storage of DNA if completely protected from water and oxygen.
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ABSTRACT: A new procedure for room-temperature storage of DNA was evaluated whereby DNA samples from human tissue, bacteria, and plants were stored under an anoxic and anhydrous atmosphere in small glass vials fitted in stainless-steel, laser-sealed capsules (DNAshells(®)). Samples were stored in DNAshells(®) at room temperature for various periods of time to assess any degradation and compare it to frozen control samples and those stored in GenTegra™ tubes. The study included analysis of the effect of accelerated aging by using a high temperature (76°C) at 50% relative humidity. No detectable DNA degradation was seen in samples stored in DNAshells(®) at room temperature for 18 months. Polymerase chain reaction experiments, pulsed field gel electrophoresis, and amplified fragment length polymorphism analyses also demonstrated that the protective properties of DNAshells(®) are not affected by storage under extreme conditions (76°C, 50% humidity) for 30 hours, guaranteeing 100 years without DNA sample degradation. However, after 30 hours of storage at 76°C, it was necessary to include adjustments to the process in order to avoid DNA loss. Successful protection of DNA was obtained for 1 week and even 1 month of storage at high temperature by adding trehalose, which provides a protective matrix. This study demonstrates the many advantages of using DNAshells(®) for room-temperature storage, particularly in terms of long-term stability, safety, transport, and applications for molecular biology research.Biopreservation and Biobanking 06/2014; 12(3):176-183. · 1.58 Impact Factor
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ABSTRACT: The authors report the unusual thermal stability of deoxyribose nucleic acid (DNA) origami when adhered to a solid substrate. Even when heated to 150 °C for 45 min, these DNA nanostructures retain their physical and chemical integrity. This result suggests that DNA origami could be integrated into applications requiring moderate substrate heating, such as photoresist baking or chemical vapor deposition processes.Journal of vacuum science & technology. B, Microelectronics and nanometer structures: processing, measurement, and phenomena: an official journal of the American Vacuum Society 07/2014; 32(4):040602-040602-4. · 1.36 Impact Factor
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ABSTRACT: Molecular dynamics simulation and biophysical analysis were employed to reveal the characteristics and the influence of ionic liquids (ILs) on the structural properties of DNA. Both computational and experimental evidence indicate that DNA retains its native B-conformation in ILs. Simulation data show that the hydration shells around the DNA phosphate group were the main criteria for DNA stabilization in this ionic media. Stronger hydration shells reduce the binding ability of ILs' cations to the DNA phosphate group, thus destabilizing the DNA. The simulation results also indicated that the DNA structure maintains its duplex conformation when solvated by ILs at different temperatures up to 373.15 K. The result further suggests that the thermal stability of DNA at high temperatures is related to the solvent thermodynamics, especially entropy and enthalpy of water. All the molecular simulation results were consistent with the experimental findings. The understanding of the properties of IL-DNA could be used as a basis for future development of specific ILs for nucleic acid technology.Physical Chemistry Chemical Physics 06/2014; · 4.20 Impact Factor
Chain and conformation stability of solid-state DNA:
implications for room temperature storage
Jacques Bonnet1,2,*, Marthe Colotte3, Delphine Coudy3, Vincent Couallier4,
Joseph Portier5, Be ´ne ´dicte Morin6and Sophie Tuffet7
1Universite ´ de Bordeaux-plateforme Ge ´nomique Fonctionnelle,2Institut Bergonie ´-INSERM U916 VINCO,
Bordeaux,3socie ´te ´ IMAGENE-Recherche et De ´veloppement, plateforme Ge ´nomique Fonctionnelle, Bordeaux
Pessac,4Universite ´ de Bordeaux-Institut de Mathe ´matiques de Bordeaux UMR 5251,5Universite ´ de
Bordeaux-Institut de la Chimie de la Matie `re Condense ´e de Bordeaux,6Universite ´ de Bordeaux-UFR de Chimie,
Institut des Sciences mole ´culaires, Bordeaux and7Socie ´te ´ IMAGENE-Plateforme de production, Genopole,
Received April 10, 2009; Revised November 1, 2009; Accepted November 2, 2009
There is currently wide interest in room temperature
storage of dehydrated DNA. However, there is insuf-
ficient knowledge about its chemical and structural
stability. Here, we show that solid-state DNA degra-
dation is greatly affected by atmospheric water
and oxygen at room temperature. In these condi-
tions DNA can even be lost by aggregation. These
are major concerns since laboratory plastic ware
is not airtight. Chain-breaking rates measured
Arrhenius’ law. Extrapolation to 25?C gave a degra-
dation rate of about 1–40 cuts/105nucleotides/
century. However, these figures are to be taken as
very tentative since they depend on the validity of
the extrapolation and the positive or negative effect
of contaminants, buffers or additives. Regarding the
showed that DNA secondary structure could be
except possibly for small fragments. Indeed, below
about 500bp, DNA fragments underwent a very slow
evolution (almost suppressed in the presence of
trehalose) which could end in an irreversible denat-
uration. Thus, this work validates using room tem-
perature for storage of DNA if completely protected
from water and oxygen.
There is currently a wide consensus for the need to
preserve DNA. Among other aims, DNA samples need
to be stored, sometimes in huge numbers, for constituting
biorepositories and preserving genetic diversity, for
large-scale genetic studies and for medical, clinical, phar-
maceutical and forensic applications. In view of the expo-
nential increase in the number of samples to be stored,
classical storage in freezers appears cumbersome, costly
and not without risk of failure. Therefore, room temper-
ature storage is currently an alternative gaining an
However, dehydration cannot really be considered as
an effective room-temperature conservation procedure
unless more information about the stability and storage
life of dehydrated DNA is acquired. In this introduction,
we review relevant data regarding DNA degradation.
Since degradation kinetics have mostly been based on
measuring chain breaks, we pay particular attention
to DNA alterations which are not directly related to or
not causing DNA chain breaks. We refer to work con-
ducted in vivo and in solution, leaving aside events
related to alterations due to light, ionizing radiation and
xenobiotics. For reviews about DNA degradation see for
instance refs. 1–5.
In vivo studies
In vivo, DNA is degraded mainly by water and reactive
oxygen species (ROS). Hydrolysis mainly leads to
single-strand breaks following depurination. It also
causes base deamination which is not followed by chain
breakage [for instance cytosine can be converted into
uridine (6)]. Other pathways including removal of
pyrimidines (7) or phosphodiester hydrolysis (8) are
much slower. Oxidation concerns both sugar and bases:
attacks on deoxyribose generally lead to chain breaks or
to formation of abasic sites; attacks on the bases generate
a wide variety of modifications; some of them lead to the
*To whom correspondence should be addressed. Tel.: 33 5 56 33 04 22; Email: email@example.com; firstname.lastname@example.org
Published online 7 December 2009Nucleic Acids Research, 2010, Vol. 38, No. 51531–1546
? The Author(s) 2009. Published by Oxford University Press.
This is an Open Access article distributed under the terms of the Creative Commons Attribution Non-Commercial License (http://creativecommons.org/licenses/
by-nc/2.5), which permits unrestricted non-commercial use, distribution, and reproduction in any medium, provided the original work is properly cited.
rupture of the N-glycosidic or phosphodiester bond, but
the major modifications (8-oxo-gua and thymine glycol)
do not. For reviews see refs. 4,9,10.
In vitro studies (solution)
Numerous studies on DNA degradation in solution
have demonstrated that the main degradation pathway
is an acid-catalyzed, water-independent, cleavage of the
N-glycosidic bond, followed by water addition on the
2?C of the sugar at least at acidic or moderate pH.
Subsequently, base-catalyzed b-elimination removes the
phosphate (11–14). Direct elimination of the base has
also been observed for guanine, but not for adenine (15).
Oxidation also exists in solution. It results from ROS
generated from triplet oxygen and water in the presence
of catalyzing impurities such as metal ions or lipids
(16,17). Base oxidation (16,18) and direct strand breaks
(11,19) have also been observed. However, as in vivo,
oxidation of bases is generally not immediately followed
by the rupture of the phosphodiester or N-glycosidic
linkages (4,20,21). This is especially the case with the
major oxidation product, 8-oxoguanine (9). 8-oxoguanine
can further react with lysine to give DNA–protein
cross-links (22). In some rare instances, DNA–DNA
cross-links induced by oxidation have been reported
The fact that DNA is more stable in a solid state
(‘solid state’ defines DNA that is no longer dissolved
without necessarily being totally anhydrous) than in
solution is widely accepted, although a few studies have
reported that desiccation induces strand breaks and
cross-links in spores (25). There are many examples of
DNA preservation by various desiccation procedures
(precipitation in ethanol, air drying or lyophilization)
(26). Spectacular examples of this stability are the
retrieval, polymerase chain reaction amplification and
sequencing of DNA from up to 80000-year-old biological
remnants (27–30). It was found that besides being heavily
fragmented, this ancient DNA contained large amounts of
oxidized (28) or hydrolyzed bases (31) and interstrand
also been reported to be a secondary consequence of
These observations are strong support for the possibil-
ity of room-temperature conservation of DNA since,
unlike ancient DNA, purified dehydrated DNA can be
protected from water and oxygen. It should not suffer
from either early post-mortem damage (through autolysis
and microbial attacks) or contaminants that catalyze oxi-
dation and lead to cross-links through the Maillard
Solid-state DNA degradation mechanisms and kinetics
There are few data on the mechanism(s) of DNA degra-
dation in more controlled conditions. Depurination has
been observed and depurination rates were measured
depurination and rapid subsequent chain cleavage in
oligodeoxynucleotides were observed by using various
mass spectrometry techniques (39–41).
Atmospheric oxygen has been found to accelerate
DNA decay in lyophilized rat liver, but not if the tissue
was delipidated before storage. Likewise, there is no
evidence for increased breakage from oxidation in dried
purified lambda DNA (42). Working on aged desiccated
colonies of Nostoc communae, Shirkey et al. (33) observed
cross-links produced by Maillard products, but no detect-
able increase in the level of oxidized bases. It has also
been reported that dehydration leads to DNA–protein
cross-links in spores (25). In lyophilized pharmaceutical
plasmid preparations, ROS production and probably
oxidation could be modulated by addition or complex-
ation of metal ions, but practically no effect on
chain-breaking was observed (17,43).
In vacuo, the bases themselves are quite stable since it
has been shown in pyrolysis experiments that while DNA
is completely destroyed in a few minutes above 180?C (44),
bases can be recovered after several hours of heating
at 250?C (45) or 5min at 450?C (46). Various thermal
decomposition experiments indicate that, in gas phase,
nucleosides are much less stable than bases and that
N-glycosidic linkage is the most fragile site (5,45,47,48).
More generally, studies on oligonucleotide fragmentation
have revealed multiple possible mechanisms of chain
breakage, most of them involving a preliminary base
loss, for a review see ref. 41.
Solid-state reaction kinetics have been thoroughly
studied, for instance on pharmaceutical preparations (for
reviews, see refs 49 and 50). Several general observations
are relevant to our topic. First, the reaction mechanisms
occurring in solution are generally maintained in the
solid state. In particular, acid-catalyzed reactions in
solution still occur in the solid state. This is related to
‘pH memory’ due to the fact that when protonated in
solution, a chemical group remains protonated in the
solid state (51,52). Second, establishment of a solid state
leads to a strong decrease in molecular mobility and con-
sequently in chemical reactivity. This effect can be
increased by using additives such as trehalose, which
creates a vitreous matrix (53,54). Third, residual water
can act either as a reactant or as a ‘plasticizer’ by
increasing molecular mobility (50,53–55).
Regarding DNA, information from samples extracted
from fossils or museum specimens are difficult to use
because of the immediate post-mortem degradation
events. Cherng et al. (56) reported that desiccated DNA
could be stored for 10 months without visible degradation
at room temperature. On the contrary, Shirkey et al. (33)
showed that dried DNA stored over P2O5 underwent
strong degradation in <72h unless trehalose was added,
and that even in the presence of the latter, degradation
was quite evident after 17 days. However, the assay of
Cherng et al. (genomic DNA sizing by nondenaturing
agarose electrophoresis) is much less sensitive to degrada-
tion than that of Shirkey et al. (33; relaxation of a 12-kb
plasmid). In the same paper, it was reported that DNA
underwent aggregation in the absence of trehalose.
Lyophilized plasmid DNA has also been used to study
rates of chain breakage at room temperature, 40?C
1532 Nucleic Acids Research, 2010,Vol.38, No. 5
and 60?C (17,26). Strangely enough, these rates were
almost insensitive to temperature (Figures 10 and 11).
This apparent discrepancy is discussed in the section
‘Temperature dependence of chain-breaking rate: effect
of residual water’. On the contrary, another study (38)
showed that the kinetics of depurination of ‘vacuum-
dried’ DNA followed Arrhenius’ law between 97?C
Solid-state DNA secondary structure
Another aspect of the storage of dried DNA concerns the
preservation of its secondary structure. Since water
strongly contributes to the secondary structure of DNA
(for a review, see ref. 57), the consequences of dehydration
should be considered. It is well known that upon dehydra-
tion [between 92% and 70% relative humidity (RH)],
DNA undergoes conformational changes from form B
to form A (58) (for reviews see refs 59 and 60).
Moreover, below 70–50% RH at room temperature,
natural DNA undergoes denaturation. Destacking of the
bases and rupture of hydrogen bonds has been observed
by ultraviolet (UV) and infrared (IR) spectroscopy
(61–63). Mass spectroscopy experiments have shown that
oligodeoxynucleotide duplexes can still exist in vacuo (64)
but that they are thermodynamically unstable and have
a short lifetime (40). On the contrary, double-stranded
homopolymers do not denature (65–67). This difference
could be due to differences in electrostatic interactions
between double helices induced by removal of water
(68–70) (for a review see refs 57 and 71).
Dehydration-induced denaturation is entirely reversible
upon rehydration unless the dried DNA has been heated
(44). This loss of reversibility indicates that there exists a
possibility of evolution over time of the DNA secondary
structure; however, no study has yet been carried out on
the evolution of solid-state DNA secondary structure.
heating is reversible (67). It has also been reported that
trehalose has a strong stabilizing effect on DNA secondary
structure. In the presence of trehalose, solid-state natural
DNA, even heated to 120?C, does not denature (72). This
stabilization effect has been explained by a screening of
the negative charges by trehalose binding to the phos-
phates (water replacement hypothesis) or a network built
by hydrogen bonding between trehalose and DNA, which
reduces the structural fluctuations of DNA (vitrification
hypothesis) (72,73). This will be discussed further in the
‘Results and discussion’ section in the section on second-
ary DNA structure.
Finally, a molecular dynamic simulation has been used
to study DNA double helix in the gas phase. Rueda et al.
(74) concluded that in vacuo, the DNA helix is highly
distorted with a very elongated conformation. Part of
the base pairing is lost, the stacking being maintained
with the bases protruding outside the helix. However,
these conclusions strongly depend on how the DNA
molecule is considered to be charged. If the DNA is
fully charged, no helical structure is maintained.
In summary, removal of water increases DNA chemical
stability first by inhibiting hydrolysis and oxidation and
second by decreasing molecular mobility. Conversely,
residual water may act as a plasticizer as well as a
reactant or a catalyst. The fastest degradative event in
the solid state seems to consist in an acid-catalyzed,
water-independent base loss followed by or simultaneous
to chain breakage via a variety of mechanisms not neces-
sarily involving water. However, oxidation, cross-linking
and the action of many agents, some possibly stabilizing
(for instance, Tris or trehalose) (33,75,76) or more gener-
ally destabilizing (such as metallic ion contaminants),
cannot be neglected. Finally, dehydration of natural
DNA leads to a strong distortion of its secondary struc-
ture, probably ending in reversible denaturation.
Here, we report an empirical study that evaluates
DNA at low hydration with the final aim of establishing
and validating conditions for long-term room-temperature
MATERIALS AND METHODS
We purposely used standard DNA extraction procedures
in order to work with the DNA quality generally used in
Plasmids (pBAD and pcDNA3, 5.4kb or pEGFP,
4.9kb) were produced in Escherichia coli. The transformed
bacteria were cultivated in Luria Bertani (LB) broth con-
taining 50mgml?1ampicillin. The plasmids were isolated
via alkaline lysis according to (77) and purified using a
silica column (Qiagen or Promega). The pEGFP plasmid
was desalted on Sephacryl?S-400 before use. They were
stored at 4?C or –20?C.
Equine genomic DNA was extracted from blood using
the Puregene?kit (Gentra) according to the manufac-
turer’s instructions or a standard phenol:chloroform
protocol. Briefly, red blood cells were lysed by a 0.16M
NH4Cl, 10mM KHCO3 and 1mM EDTA solution at
room temperature. Then white blood cells were lysed
overnight at 37?C in 5mM NaCl, 2mM EDTA, 1%
SDS and 100mgml?1proteinase K. Two phenol:chloro-
form:isoamyl alcohol (25:24:1) extractions were per-
formed and followed by a chloroform:isoamyl alcohol
(24:1) extraction. To the aqueous phase were added
2vol. of absolute ethanol and 0.1vol. of 5M NaCl for
precipitating the DNA and the pellet was washed with
70% ethanol. DNA was finally dissolved in TE (10mM
Tris–HCl pH 8.0, 1mM EDTA) and stored at –20?C or
Agarose gel electrophoresis and determination
of supercoiled plasmid content
Plasmid DNA (250ng) or genomic DNA samples
(250ng to 2mg) were loaded onto 0.8–1% agarose gels
and submitted to electrophoresis in TAE 0.5? (40mM
Tris acetate, 1mM EDTA) electrophoresis buffer. Gels
were stained 30minin 0.5mgml?1ethidium bromide or
in 1/10,000 SybrGreen?I (Molecular Probes). Gels were
photographed with a Visiomic digital Imaging apparatus
(Genomic, Archamps, France) equipped with an ethidium
Nucleic Acids Research,2010, Vol.38, No. 51533
bromide or Sybrgreen?fluorescence filter (555nm). The
images were analyzed with the Kodak Digital Science
1D Image analysis software. The fluorescence inten-
sity of supercoiled plasmid and the sum of the fluores-
cence intensities of relaxed and linear forms were
The proportion of supercoiled plasmid (SC content) in
the samples was calculated either directly from the fluo-
rescence intensity values with Sybrgreen?stained gels,
or by correcting (for the ethidium bromide-stained gels)
the fluorescence intensity of the supercoiled form by
1.4 factor as previously described (78). Alternatively,
(for both dyes) SC content was calculated by reference
to standards (supercoiled, relaxed and/or linear plasmid)
run in the same gel. In our conditions, differences in rate
constants between these methods were negligible. For an
example, see Supplementary Data S1 (Plasmid degrada-
tion in solution). The kinetic constants were classically
determined by fitting the curves to data points assuming
an exponential decrease (11,79,80) Supplementary Data
S2 (Statistical analysis)].
DNA melting was monitored by UV absorption on a
Uvikon 940 spectrophotometer with a thermostated cell
holder connected to a Huber cryothermostat driven
by a Huber PD410 programmer, as described by (81).
The temperature was measured by means of a Pt probe
immersed in a water cuvette placed in the sample holder.
Samples (6mg) were diluted in cacodylate 20mM sodium,
1mM EDTA buffer, pipetted into 1-cm path length quartz
cuvettes and preincubated in the cell holder at 25?C
for 30min. The temperature was raised 0.5?C/min from
25?C to 95?C.
A260nm of samples was measured before (native) and
after heat denaturation and immediate cooling in ice
(denatured). The relative hyperchromicity was defined
as either: (A260, denatured – A260, native)/A260, native when
looking for cross-links or (A260,denatured–A260,
A260, denaturedwhen looking for denaturation.
We checked the linearity and the sensitivity of this assay
by applying it to a series of mixtures of native and
previously denatured DNA (Figure 1).
Detection of apurinic (AP) sites by cleavage with APE-1
Preparation of a partially depurinated DNA control. Using
the method described in (12), pBAD plasmid DNA
(0.5mMnt) was partially depurinated by heating 15min
at 70?C in 0.1M NaCl, 0.01M sodium citrate and 0.01M
Tris–HCl at pH 5.2. Depurinated DNA was then
precipitated 2h at –20?C with 2vol. of absolute ethanol,
0.1vol. of 5M NaCl and 1ml of 20mgml?1glycogen.
After a 20-min centrifugation, the pellets were washed
three times with 70% ethanol and suspended in deionized
Cleavage at apurinic sites. Human apurinic/apyrimidinic
endonuclease Ape-1 (New England Biolabs) was used to
cleave the phosphodiester backbone immediately 5 to an
AP site to generate a single-strand DNA break according
to the manufacturer’s instructions. The samples were
analyzed by agarose gel electrophoresis and ethidium
bromide staining. The number of cleaved sites was
estimated from SC content as described.
Measurement of 8-oxodG
For a 155-ml final volume, DNA (15–60mg) was incubated
with 21ml of nuclease P1 buffer (300mM sodium acetate
and 1mM ZnSO4pH 5.3) and 10ml (10 units) of nuclease
P1 at 37?C for 2h. Dephosphorylation of samples was
achieved by the addition of 23ml of 500mM Tris–HCl
and 1mM EDTA pH 8.0 buffer and 1ml (1 unit) of
alkaline phosphatase (Roche) at 37?C for 1h. About
100ml of the hydrolysate were analyzed by HPLC-EC.
High-pressure liquid chromatography/electrochemical
detection (HPLC-EC) measurement of 8-oxodG was per-
formed with a Beckman Series pump system equipped
with a pulse damper, a cooling autosampler and a
spectrophotometric detector connected to a Kontron
amperometric detector. The electrochemical cell was
operating at 650mV versus an Ag/AgCl reference elec-
trode. The system was operated at 0.5-nA full-range detec-
tion. The deoxynucleosides was detected at 254nm. HPLC
separation was obtained on an Uptisphere ODB C18
column (5-mm particle size, 250?4.6mm) equipped with
an Ultrasphere ODB C18 guard column (5-mm particle
size, 50?4.6mm) (Interchim,). The mobile phase used
for isocratic elution of 8-oxodG was composed of
12.5mM citric acid, 15mM sodium acetate, 30mM
NaOH pH 5.3 with 10% methanol at a flow rate of
0.8ml/min. The injection volume was 100ml. The dG con-
centration was estimated from the UV peak and the
8-oxodG concentration from the electrochemical signal.
Results are expressed as the number of 8-oxodG
residues per 106dG.
Figure 1. Hyperchromicity: linearity and sensitivity of the assay.
Relative hyperchromicity (A260, denatured–A260, native)/A260, denatured) of
mixtures of native and previously denatured DNA samples for
high-molecular-weight and 400-bp DNA. The straight lines drawn
through data points are y=0.1958x (R2=0.981) and y=0.953x
(R2=0.9847) for high-molecular-weight and 400-bp DNA, respectively.
1534 Nucleic Acids Research, 2010,Vol.38, No. 5
To calculate the 8-oxodG formation rate per nucleotide,
we first corrected the measured value by the dG content of
mammalian DNA (we took 21.5% for all of them). Then,
considering that the level of 8-oxodG was always below
1% of the amount of dG, meaning that we were in the
linear part of the kinetics, we calculated oxidation rates
by simply dividing the values obtained (after subtracting
the corresponding control) by the incubation time in
For addition of ferric ions, we prepared 35-mg DNA
aliquots in 118ml TE containing 40 molecules of trehalose
per nucleotide and a large excess (89 equivalents per
nucleotide) of Fe3+ions. The aliquots were vacuum-dried
as described below.
DNA dehydration, incubation conditions and rehydration
Dehydration and rehydration. About 10–20ml of a plasmid
DNA solution (1–5mg of DNA) were deposited on the
bottom of 0.2-ml cylindrical glass inserts. All samples
were vacuum-dried for 30min to 1h (Thermo Savant)
heating, DNA was dried in the presence of trehalose
(40 molecules per nucleotide). Trehalose was included
because we noticed that when DNA was stored in air,
most of the material was occasionally impossible to
redissolve, probably because of irreversible aggregation
or adsorption, as previously noted by others who also
showed that trehalose could prevent this phenomenon
(33,82); Supplementary Data: S3 (Aggregation and inhibi-
tion by trehalose) and also the section ‘Evolution of DNA
secondary structure of dehydrated DNA’.
For rehydration, vials or tubes were opened and inserts
were placed in 1.5-ml microtubes. Samples were then
rehydrated with water or TE buffer at room temperature
or by two successive 15-min incubations at 37?C.
(i) Open air (generally around 50% RH): the inserts
(ii) Open air at 11%, 28%, 50% and 75% RH: the
inserts were placed in uncapped tubes inside closed
glass bottles containing saturated solutions of LiCl,
MgCl2, NaBr and NaCl (prepared and equilibrated
at incubation temperature).
(iii) Air at low RH: the inserts were placed in tubes
which were subsequently put into ‘trap-bottles’
(Duran?ISO bottles containing P2O5with vacuum
(iv) Anhydrous argon atmosphere: 2-ml vials with
inserts containing vacuum-desiccated DNA were
equilibrated in a dry box containing an anhydrous
(H2O<1p.p.m.) and anoxic (O2<0.1p.p.m.) argon
atmosphere, and the silicon stoppers and tear-off
aluminum seals were crimped on the top of the
vials with a Wheaton hand-operated crimper.
(v) Incubation in the presence of CaCl2. DNA samples
were kept in a dessiccator with CaCl2. This proved
to be ineffective probably because of the large
volume of the dessiccator.
Gravimetric control of air tightness of containers. The
various containers were crimped glass vials (Wheaton,
VWR, operated as described above), 1.5-ml plastic tubes
(‘Easy fit’ and ‘Click fit’ Treff microtubes (Treff AG),
Simport O-Ring screw cap microtubes) and glass bottles
(Duran?ISO) with or without greased caps. The small
containers received ?1g of CaCl2 and were regularly
weighed. Air tightness of the bottles was checked
with open 1.5-ml tubes loaded with CaCl2 (Figure 2).
Figure 2. Control for air tightness of containers. The containers were filled with 1g CaCl2and regularly weighed. The lines serve as visual guides.
Nucleic Acids Research,2010, Vol.38, No. 51535
Empty containers were weighed in parallel as controls
(data not shown).
Use of P2O5. Phosphorus pentoxide was effective at room
temperature. However, we occasionally noticed a rapid
degradation of DNA samples (data not shown). This
was probably due to P2O5dust, because this effect was
prevented by covering the insert with aluminum foil.
This probably explains why P2O5 seemed to accelerate
DNA degradation, as reported in the literature (33).
When heated to 90?C in closed bottles, we witnessed
a browning of the plastic tubes and an acceleration of
DNA degradation. This was probably due to traces of
vapor emitted from heated P2O5 or phosphoric acid
(boiling point: 158?C).
Accelerated degradation studies
Vacuum-dried samples were heated at temperatures
ranging from 70?C to 140?C either in an oil bath or in
an oven equipped with a sand bath. A thermometer was
placed in an empty insert to control the inner temperature
of the samples. Samples were sequentially removed from
the oven or water or oil bath, quickly cooled on ice, and
stored at 4?C until rehydration and DNA analysis. The
first time point (for time 0) was taken after allowing the
sample toreach temperature
Preparation of different-sized DNA
obtained by sonication (Branson sonifier 150) at power
3. DNA fragment sizes were checked by agarose gel
DNAfragments of differentsizeswere
RESULTS AND DISCUSSION
Initially, the work reported here was aimed at estimating
the lifetime of dehydrated DNA. Considering that
the DNA degradation rate is too slow to be conveniently
measured at room temperature, we ran DNA chain-
breaking kinetics at temperatures ranging from 70?C to
140?C. Chain-breaking was monitored by measuring
the relaxation of a supercoiled plasmid. However, the
chain-breaking rate is not the only parameter of the deg-
radation rate since many base modifications do not
generate chain breaks. Most of these modifications are
caused by chemicals and are not relevant here. However,
some others, mainly cross-links or oxidized bases, can
either bias or prevent further DNA analysis, for
instance, by impeding the action of DNA polymerases
because of cross-linking and oxidative events. We have
designed experiments to estimate the level of these events.
To be able to detect cross-links in the context of degrada-
tion, we worked on two different-sized DNA. Calf thymus
DNA samples (average single-strand sizes: 2000nt and
500nt) were heated in air at 110?C in the presence of
trehalose. At each time point, the absorbance of the
sample was measured immediately after rehydration
(native) and then after heating to 95?C and rapid
Figure 3A shows the relative hyperchromicity for
samples of both sizes as a function of heating time at
110?C, while Figure 3B shows the average fragment size
for the longer fragment. This size decreased at a rate in
agreement with the rate calculated from the plasmid relax-
ation data (Figure 10) (plasmid relaxation should not
be affected by a moderate number of cross-links). This
indicated that cross-links were not as abundant as chain
breaks. However, in both samples, there was a decrease in
hyperchromicity with a stronger decrease in the shorter
Figure 3. Control
2000-nt and 500-nt average sizes of fragments after vacuum drying
and heating kinetics at 110?C. Some error bars are smaller than the
symbols. The lines serve as visual guides. (B) Average size of the large
fragment population measured by denaturing gel electrophoresis. The
continuous line was calculated by curve-fitting (giving a corresponding
degradation rate of 5.5?10?9s?1nt?1). The dotted line represents the
decrease in average fragment size calculated from plasmid degradation
data (degradation rate: 3.5?10?9s?1nt?1at this temperature). Some
error bars are smaller than the symbols.
native) of DNA populations of
denatured – A260,
1536 Nucleic Acids Research, 2010,Vol.38, No. 5
fragments. This was not due to cross-linking but rather to
the appearance in the samples of denatured DNA, with a
larger proportion in the smaller samples. This is dealt with
in greater details in the section devoted to the evolution of
the secondary structure of DNA.
DNA oxidation was assessed by measuring the amount of
8-oxodG in DNA. Figure 4 shows the results obtained
from controls and from DNA samples stored or incubated
in various conditions.
First, at 100?C, in air, the production rate of 8-oxodG
in DNA dried from a TE solution was 1.2?10?10s?1nt?1,
which is about 200-fold lower than the solution rate
interpolated from (16) for this temperature. Second, tre-
halose seemed to have an inhibitory effect on 8-oxodG
formation at 100?C, (decreasing from 8.1?10?10s?1nt?1
to 2.1?10?10s?1nt?1and from 1.5?10?10s?1nt?1to
2.4?10?11s?1nt?1in Tris or phosphate buffers respec-
tively) but not at room temperature. Third, partially
removing oxygen had only a moderate effect (decreasing
from 1.2?10?10s?1nt?1to 5.3?10?11s?1nt?1). This
could mean that oxidation necessitated only a low concen-
tration of oxygen to occur. Fourth, atmospheric humidity
accelerated the rate of oxidation 100-fold and 30-fold
between 1.4% RH to 75% RH in TE and phosphate
buffer, respectively. Addition of a saturating amount of
Fe3+ions also seemed to enhance the rate of oxidation.
In summary, on one hand, DNA oxidation did occur in
air, and, on the other hand, the presence of water
accelerated the rate of oxidation.
However, while in vivo, 8-oxodG formation is widely
considered as a major DNA oxidation product, the situa-
tion could be quite different in the solid state. Future work
will therefore adopt more general approaches such as the
use of enzymes (e.g. Endo III/EndoVIII, UNG, etc.) able
to detect oxidized or modified bases and the chemical
analysis of the DNA degradation products.
Control for the absence of accumulation of abasic sites
during DNA heating at low hydration
It may be thought that acid-catalyzed depurination is still
present in the solid state. Indeed, mechanisms occurring in
solution have generally been shown to be maintained in
the solid state (for a review see ref. 49), and depurination
has been observed in vacuo or in solid-state DNA (38–40).
It is possible that at low hydration, subsequent chain
breakage does not occur or occurs very slowly leading
to accumulation of abasic sites. To estimate the extent
of accumulation, we used the Ape enzyme to reveal
apurinic sites in heated DNA. Figure 5, ‘control
solution’, shows that a partially depurinated plasmid
(positive control) treated with Ape enzyme lost 28%
(0.91–0.63) of its SC content, while the undepurinated
control lost only 2% (0.96–0.94) of it. The same was
true if the depurinated plasmid and the undepurinated
control were dried, immediately rehydrated and treated
with Ape (Figure 6, ‘dried’, ‘nonheated’): the losses
were respectively 30% (0.92–0.63) and 3% (0.89–0.86).
Figure 4. Level and rate of 8-oxodG formation as a function of incubation conditions.afrom (11);bfreshly extracted;capproximate value;d1mM
Fe3+before drying; w: weeks; y: years; sol: in solution; c rv: crimped vials; P: phosphate buffer; PE: phosphate buffer + EDTA; T: Tris buffer;
TE: Tris buffer + EDTA. The term nt?1is used only to normalize the rates to one nucleotide to make the reaction rates independent of the size of
Nucleic Acids Research,2010, Vol.38, No. 51537
These data showed first that the positive control had been
significantly depurinated and that the enzyme treatment
detected the depurinated positions, and second, that mere
drying did not induce breaks at abasic sites.
When the plasmids were incubated at 118?C for 1 and
2h, the difference between Ape- treated or untreated
depurinated plasmid disappeared almost completely.
Therefore, upon heating all the previously depurinated
sites had been cut. Moreover, the heated undepurinated
plasmid lost 60% and 61% of SC content, respectively
before and after Ape treatment, suggesting that no
abasic sites had accumulated during the treatment at
118?C. The same was true for the depurinated plasmid,
the difference before and after Ape treatment being insig-
nificant. This meant that any existing abasic site had been
cut upon heating and that no accumulation was detectable
in the conditions we used for our kinetics.
Chain-breaking in air at different relative humidities, at
70?C or room temperature
We first compared DNA degradation rates at 70?C.
Figure 6A shows that at 75% RH, the SC content
than inopen air
respectively). As a control, we ran a kinetic in TE buffer
solution)], giving a k70?C=4.42?10?9s?1nt?1. In a
separate experiment, we measured the degradation rates
as a function of RH to check for an eventual threshold
below which the degradation rate would no longer depend
on water content. RH was varied from 75% to an
estimated 5%. Indeed, at room temperature and in open
air, increasing temperature from 25?C to 70?C increases
10-fold the saturating water vapor pressure, leading con-
sequently to a 10-fold RH decrease. In usual laboratory
conditions, this leads to an RH decrease from 50% to 5%
and a concomitant decrease in DNA-bound water con-
tent according to the isotherms of binding water
to DNA (44,84). Such a water release was confirmed in
Figure 6. DNA degradation as a function of RH. (A) Plasmid DNA
samples containing trehalose were incubated at 70?C in capped bottles
containing an NaCl saturated solution (giving a RH of 75%). DNA
samples were then rehydrated and the plasmid SC content was deter-
minedon Sybrgreen?-stained agarose
(‘Materials and methods’ section). The kinetic run in open air is
redrawn from the 70?C experiment in Figure 10. (B) Closed circle:
same experiment with additional RH values done with another
plasmid (which consistently gave higher degradation rates than that
used for the other experiments). The numbers refer to Figure 11. The
dotted straight line serves as a visual guide. Closed diamonds: 8-oxodG
rate formation at 100?C. The continuous line is an exponential fit
through the circles. The 8-oxodG formation rates were determined
from single measurements and should be viewed more as comparisons
than absolute values. The numbers refer to Figure 4.
Figure 5. Control for the absence of accumulation of abasic sites
during DNA heating at low hydration. Quintuplicate depurinated or
undepurinated plasmid DNA samples were vacuum-dried, then heated
for 0, 1 or 2h at 118?C, rehydrated and treated or not with Ape to
cleave abasic sites in parallel to control samples kept in solution. SC
content was determined as described in ‘Materials and methods’
section. On the gels, the upper and lower bands are the relaxed (OC)
and supercoiled (SC) plasmid forms.
1538Nucleic Acids Research, 2010,Vol.38, No. 5
[Supplementary Data S4 (DNA water content, gravimetric
analysis)] performed to determine the minimum time and
temperature necessary to obtain complete DNA dehydra-
tion in open air. At 112?C and above, we could no longer
detect DNA-bound water. Figure 6B shows that between
75% and 5%, the chain-breaking rate decreased almost
linearly with RH so there was probably no threshold,
although it could not be formally excluded that such a
threshold existed for a value below 5%. We also noticed
that the water rate dependency was very different for oxi-
dation and chain-breaking. This could be expected since
water is likely to have different actions on these two
phenomena (see discussion below in the ‘Temperature
dependence of chain-breaking rate: effect of residual
In a second series of experiments, in order to optimize
the conservation conditions, we compared chain-breaking
rates at room temperature in the presence or absence of
trehalose, in open or closed tubes, in closed tubes inside a
box containing CaCl2and in vials crimped under a dry
argon atmosphere. Figure 7 shows that in all the condi-
tions tested, DNA degradation at room temperature was
rather fast and that, surprisingly, stability was not
increased in the crimped vials or only marginally in the
presence of CaCl2.
These facts led us to conclude that our different
containers, and especially the crimped vials, were not
conditionsby control gravimetricanalyses
fully airtight. This was confirmed by a series of gravimetric
controls (‘Materials and methods’ section, Figure 2 and
ref. 85). We then concluded that in all the closed con-
tainers we were using, a constant and rather rapid
exchange with the outside atmosphere was taking place
and that, in the box, water absorption by CaCl2was too
slow for this desiccant to have more than a slight impact
on the internal RH. The experiment was then repeated
with glass bottles containing the more potent desiccant
P2O5. Figure 8 clearly shows that stability was improved
to the point that no degradation could be detected after a
2-month period, while at 75% RH or under ordinary
atmosphere, the degradation clearly appeared.
The degradation rates measured at room temperature
varied from 1.4?10?12s?1nt?1to 1.1?10?11s?1nt?1.
They were in good agreement with those determined by
denaturing gel analysis of high molecular weight genomic
DNA stored for 7 years at room temperature with-
out trehalosein crimped
10?12s?1nt?1. Although originally crimped in anoxic
and anhydrous atmosphere, these samples had to be con-
Supplementary Data S5 (Determination of genomic
DNA degradation rate in air and at room temperature)].
These figures were about 10-fold higher than the rate of
(2.3?10?13s?1nt?1). However, since 8-oxodG is consid-
ered to be only 5% of the oxidized bases, hidden oxidative
most ofthe time [see
Figure 7. Plasmid relaxation at room temperature in various conditions. Plasmid DNA samples with or without trehalose were vacuum-dried in
0.2-ml glass inserts and stored in ambient air, in open or closed tubes, in closed tubes inside a box containing CaCl2or in vials sealed (crimped)
under dry argon. After 0–32 weeks, samples were rehydrated and their supercoiled content was determined on Sybrgreen?-stained agarose gel
electrophoresis as described. Controls consisted in untreated plasmid samples kept in solution at 4?C. The histogram represents plasmid SC content.
The gel performed after 32 weeks of storage is a representative example of the different analyses. On the gel, the upper, intermediate and lower bands
are relaxed (open circle), linear and supercoiled forms, respectively. (*): sample with no SC left.
Nucleic Acids Research,2010, Vol.38, No. 51539
lesions at room temperature could appear at roughly the
same rate as chain breaks.
Regarding trehalose action, although this sugar was
clearly useful to prevent DNA losses [see Supplementary
data S3 (aggregation and its prevention by trehalose)], it
did not show any clear improvement regarding its ability
to protect DNA against degradation.
Temperature dependence of chain-breaking rate
and effect of residual water
Degradation kinetics were run at temperatures ranging
from 70?C to 140?C. A first series was run in open air in
the absence of trehalose. A second series (Figure 9) was
run with a different plasmid in the presence of trehalose in
vials crimped under anoxic and anhydrous atmosphere.
However, we belatedly realized that the second series of
samples also had to be considered as having been run in
air because of the lack of air tightness of the vials. As
explained in ‘Materials and methods’ section, it was not
possible to run kinetics in the presence of P2O5.
In Figure 10, log10(k) obtained from all the kinetics
were plotted as a function of 1/T.
First, we determined that the chain-breaking rate
at 100?C was 1.2?10?9s?1nt?1while according to
Figure 4, the 8-oxodG production rate was 10-fold
lower: 1.2?10?10s?1nt?1. This likely meant that in
these conditions, the overall oxidation rate was again
similar to the chain-breaking rate.
Second, considering the temperature dependence of the
chain-breaking rate, several parameters have to be taken
into account. According to the literature and the experi-
ments reported above, two main pathways lead to
DNA degradation in air: oxidation and acid-catalyzed
depurination (or direct elimination). With regard to
chain-breaking, oxidation is probably a minor pathway
because chain-breaking in the solid state is almost insen-
sitive to the presence of metal ions (17). At high temper-
atures, DNA is almost completely dehydrated and
lowering the temperature induces two opposing effects.
On the one hand, there is a tendency for the degradation
rate to decrease due to a decreased molecular mobility and
chemical reactivity according to Arrhenius’ law. On the
other hand, the water content increases. Water tends to
accelerate degradation rates through a ‘plastifying’ effect
that enhances molecular mobility and by acting as a
reactant for hydrolysis or the production of ROS. This
combination should be noticeable even in closed vials if
the preparation containing DNA is not completely dried
and if its volume is much smaller than the volume of the
Figure 8. Plasmid relaxation at room temperature and low hydration.
Triplicate plasmid DNA samples, with or without trehalose, were
vacuum-dried in 0.2-ml glass inserts and placed in 1.5-ml tubes. The
tubes were kept in open air or placed in bottles containing P2O5with
vacuum grease-coated caps, each bottle containing only one tube
to avoid water uptake during removal of tubes. After incubation,
samples were rehydrated, submitted to electrophoresis on agarose gel
and their supercoiled content was determined after Sybrgreen?staining.
The controls were plasmid samples kept in solution at 4?C or vacuum-
dried and kept at –20?C and run on the different agarose gels.
Figure 9. Degradation
example (second series), plasmid DNA samples containing trehalose
were vacuum-dried and sealed under controlled anhydrous and
anoxic argon atmosphere (but due to leakage, these conditions were
maintained only briefly, see text). The crimped vials were heated at tem-
peratures ranging from 70?C to 140?C, DNA samples were rehydrated
and the SC contents were determined on Sybrgreen?-stained agarose
gel after electrophoresis (‘Materials and methods’ section). The first and
second panels show respectively data for 70?C and 96?C and for tem-
peratures ranging from 104?C to 131?C.
kinetics at varioustemperatures. Inthis
1540 Nucleic Acids Research, 2010,Vol.38, No. 5
container. Both situations have been encountered (26,86).
Water can also originate from additives such as trehalose
added to DNA in large amounts.
Above 70?C, the temperature dependence of the
chain-break kinetic constants followed the Arrhenius
model. This means that either the base depurination
pathway was predominant over oxidation-induced chain-
breaking or that both reactions had similar activation
energy, the latter hypothesis being less likely. At lower
temperatures, the data points drawn from our experiments
and from Anchordoquy and Molina’s data (stars on
the graph) are systematically above the straight line.
In addition, as the temperature decreases, the deviation
This situation is to be expected from the above-
mentioned discussion and it can be concluded that
above 70?C and even more above 110?C, the chain-
breaking rates were relatively independent from water
and that the value (4.6?10?15s?1nt?1) obtained by
extrapolation between 70?C and 140?C could be tenta-
tively taken as corresponding to the chain-breaking rate
at room temperature in the absence of water. Further-
more, in the absence of oxygen and consequently
without oxidation, this likely remains true, so this rate
might approximate the actual degradation rate at room
temperature in the absence of both oxygen and water.
A statistical analysis [Supplementary Data S2 (statisti-
cal analysis)] of the data obtained between 70?C
and 140?C gave an extrapolated k25?C ranging from
2.4?10?15s?1nt?1to 8.9?10?15s?1nt?1(equivalent to
be deduced from our data are 36–38kcalmol?1or
39–45kcalmol?1respectively, depending on whether we
take all the data into account or not. These activation
energies were different from those found for depurination
and oxidation (27–31kcal/mol) (11,13,14,16), probably
because of the contribution of molecular mobility.
Quantitative data from ours and from literature are
compiled Figure 11.
Evolution of the secondary structure of dehydrated DNA
temperature conservation is the secondary structure of
solid-state DNA. Previous studies have shown that, at
room temperature, natural DNA undergoes a reversible
denaturation upon dehydration. We confirmed this revers-
ibility since melting curves run on high-molecular-weight
molecules and samples fragmented to an average of 5kb
(Figure 12A) were undistinguishable from the undried
The experiment reported in Figure 12B extended
this observation to smaller fragments. However, the
results in Figure 3 suggested that upon heating at
110?C, some of the small DNA fragments underwent
irreversible denaturation. For this reason, we studied the
double-strand content in DNA fragments as a function of
RH, fragment size, heating time, temperature and
presence or absence of trehalose.
Figure 13A shows that in the absence of trehalose at
110?C, 200–400-bp fragments underwent an almost
complete loss of reversibility of denaturation in 2–4h.
Figure 13B shows that the rate of loss was considerably
lower in the presence of trehalose at 70?C. As expected,
the kinetics at 110?C, 70?C and 37?C showed that this
phenomenon was also strongly temperature dependent.
Figure 13B and C show that the rate of loss was very
similar at 28 and 75% RH. However, more experi-
ments run at other humidity levels should be performed
before concluding that the phenomenon is RH-dependent
To the best of our knowledge, this is the first time such
an evolution of solid-state DNA has been reported. In the
absence of trehalose, this may have some consequences for
short fragments. However, for long fragments, the time
required for the phenomenon to appear would be
extremely long at room temperature.
These findings could be interpreted as follows. By
removing the screening effect of water, dehydration
might allow phosphate repulsion to overcome hydrogen
bonding and stacking interactions in the double helix.
Owing to the mobility restriction imposed by the solid
state, only very slight movements are possible. Local
stretching and compression might occur, leading to loss
of base-pairing and stacking. Bases might be turned
outwards from the distorted double helix. At this stage,
Figure 10. Determination of room temperature degradation rate of
DNA at low hydration according to Arrhenius’ model. Degradation
rates (kT) were determined at temperatures (T?K) ranging from 70?C
to 140?C in different conditions. Rates measured at 70?C and room
temperatures are taken from the experiments shown in Figures 6–8.
These values are compared to some literature data as indicated. The
log10(kT) values were plotted versus 1/T. The upper and lower limits of
the shaded area represent the confidence intervals of log10(kT)
calculated with the delta method, taking into account values ranging
from 70?C to 140?C, as described in Supplementary Data S2 (statistical
analysis). (17,26): data from references 17 and 26; (F7): data from
Figure 7; (S5): data from Supplementary Data S5 (determination of
genomic DNA degradation rate in air and at room temperature).
Nucleic Acids Research,2010, Vol.38, No. 5 1541
the phenomenon is entirely reversible upon rehydration.
However, if the DNA is heated, mobility is sufficiently
increased to lead to shifts of one chain with respect to
its complementary chains, with the result that the bases
become ‘out of phase’. At this new stage, two alternative
events may take place upon rehydration. If there are still
one or several spots where the cognate bases are able to
base-pair, the whole double strand reassociates immedi-
ately. If not, the chains separate completely. An alterna-
tive hypothesis has also been proposed by Sharma and
Klibanov (82) to account for lyophilized DNA aggrega-
tion in the presence of moisture. The probability of
separation obviously increases if the fragment size
decreases. Either independently or not of this event,
bases expelled from the double helix might stack
between neighboring non-cognate chains to form small
clusters. If given enough time or accelerated by heating,
the clusters might then expand and become stronger,
creating noncovalent cross-links that lead to irreversible
The same mechanism (van der Waals interactions
involving the bases) may explain DNA denaturation asso-
ciated with irreversible adsorption on solid surfaces upon
drying and even in solution (87–89). While it is clear that
Figure 11. Compilation of quantitative kinetic data from literature and our work. (Left figure) The straight lines are the Arrhenius representations
corresponding to depurination (dep), single-strand breaks (SSB) and 8-oxodG (oxo) formation obtained from our work or from the literature. Some
of them have been truncated. The corresponding Ea (kcalmol?1) are indicated in brackets. The reactions were conducted on samples which were
vacuum-dried (vdr) or in solution (sol) DNA.acorrected to pH 7.4 according to (13);bcorrected for strandness according to (13);capproximate
values; F6, F7 and F10: respectively Figures 6, 7 and 10;dCaCl2inside a CaCl2- containing dessiccator (ineffective in providing a low RH atmo-
sphere); diamonds: kinetic constants for 8-oxodG formation, numbers refers to Figure 4. Other symbols: single-strand breaking rates. (Right table)
The term nt?1is used to normalize the rates to one nucleotide to make the reaction rates independent of the size of the molecule. lyo: lyophilized;
U: unknown; vac: sample kept under vacuum; vdr: vacuum-dried; crv: crimped vials; optub: open tubes; cltub: closed tubes; RT: room temperature;
S1, S6: respectively Supplementary Data S1 and S6.
1542 Nucleic Acids Research, 2010,Vol.38, No. 5
trehalose inhibits these phenomena, no mechanism for
their inhibition has yet been demonstrated. Trehalose
may act nonexclusively by stabilizing the DNA secondary
structure, diminishing the molecular mobility or prevent-
ing immediate contact either between neighboring double
helices or between DNA chains.
This work shows first that atmospheric water and oxygen
are detrimental to DNA preservation at room tempera-
ture. Second, although the experiments were mostly
done in the presence of atmospheric water and oxygen,
it suggests that in the absence of both water and oxygen,
DNA has a very long life time. Third, the DNA secondary
structure is also likely preserved or restored upon
rehydration, except possibly for DNA fragments smaller
than 500nt. Finally, these findings show that, if protected
from water and oxygen, dehydrated DNA should
maintain its primary and secondary structure for periods
Figure 12. Hyperchromicity
dehydrated and immediately rehydrated DNA. (A) Melting analysis
genomic DNA samples. Controls in solution are native or heat
denatured DNA. HMW: high-molecular- weight DNA. (B) Relative
populations of decreasing fragment sizes before and after vacuum
drying and immediate rehydration (measures done in triplicate).
asa functionof fragmentsize of
denatured–A260,native)/(A260,denatured) of DNA
Figure 13. Evolution of hyperchromicity as a function of size, presence
of trehalose, RH and heating time. (A) evolution of hyperchromicity at
110?C in the absence of trehalose for high-molecular-weight DNA and
0.4-kb fragments. Controls were equilibrated for one night at the tem-
peratures and in the atmospheres used in the experiments. All measures
were done in triplicate. (B) Evolution at 70?C for 0.4-kb fragments in
the presence or absence of trehalose and 28% RH and 75% RH. The
controls were done as in (A). (C) Same as in (B), at 37?C, *average of
only two measures.
Nucleic Acids Research,2010, Vol.38, No. 51543
of time beyond the most demanding needs of conserva-
Supplementary Data are available at NAR Online.
We thank C. Cabane ´ and A. Vekris for help and discus-
sions, G. Campet and J. Etourneau for discussions and
allowing us to use their dry box and P. Pourquier for
the gift of plasmids, E. Creppy, J. Robert, C. Cullin,
I. Lascu, J.J. Toulme ´ , J. Markovits and M. Bonneu for
help and support. JP Benedetto is thanked for gifts of old/
aged DNA samples and R. Cooke and A.-L. Fabre for
editing the English.
The Conseil Re ´ gional d’Aquitaine; the Po ˆ le Aquitaine
Sante ´ ; and Ose ´ o Innovation. Funding for open access
charge: IMAGENE Company.
Conflict of interest statement. Some of the authors have a
Conflict of interest in relation to the submitted work:
Tuffet,S. isCEO andshareholder
Company; Colotte,M. is an employee of IMAGENE
Company; Bonnet,J. is a shareholder and a consultant
for IMAGENE Company; Coudy,D. is an employee of
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