Báo cáo khoa học: Polyamines interact with DNA as molecular aggregates Luciano D’Agostino1 and Aldo Di Luccia doc

Polyamines interact with DNA as molecular aggregates
Luciano D’Agostino
1
and Aldo Di Luccia
2
1
Department of Clinical and Experimental Medicine, ÔFederico IIÕ University, Naples, Italy;
2
Institute of Food Science
and Technology – National Research Council, Avellino, Italy
New compounds, named nuclear aggregates of polyamines,
having a molecular mass of 8000, 4800 and < 1000 Da, were
found in the nuclear extracts of several replicating cells. Their
molecular structure is based on the formation of ionic bonds
between polyamine ammonium and phosphate groups. The
production of the 4800 Da compound, resulting from the
aggregation of five or more < 1000 Da units, was increased
in Caco-2 cells treated with the mitogen gastrin. Dissolving
single polyamines in phosphate buffer resulted in the in vitro
aggregation of polyamines with the formation of com-
pounds with molecular masses identical to those of natural
aggregates. After the interaction of the 4800 Da molecular
aggregate with the genomic DNA at 37 °C, both the
absorbance of DNA in phosphate buffer and the DNA
mobility in agarose gel increased greatly. Furthermore, these
compounds were able to protect the genomic DNA from
digestion by DNase I, a phosphodiesterasic endonuclease.
Our data indicate that the nuclear aggregate of polyamines
interacts with DNA phosphate groups and influence, more
efficaciously than single polyamines, both the conformation
and the protection of the DNA.

Keywords: DNA conformation; DNA protection; apoptosis;
molecular aggregates; polyamines.
An increased intracellular concentration of polyamines is
necessary for the activation of DNA synthesis and cell
replication [1–4]. The intestinal replicating cells are partic-
ularly capable of accumulating polyamines promoting both
their synthesis, through the activation of the enzyme
ornithine decarboxylase, and their uptake from the extra-
cellular space [5–9]. Caco-2 cells, derived from a human
colon carcinoma, after confluence spontaneously differen-
tiate assuming morphological and functional features sim-
ilar to those of the small intestinal enterocytes. This cell line
represents a useful in vitro model for studying the mecha-
nisms involved in polyamine-dependent cell replication [6,7].
Gastrin, a powerful mitogen for gastro-intestinal cells,
stimulates the growth of Caco-2 cells and increases the
intracellular concentration of polyamines promoting both
their endogenous synthesis and their uptake [7].
The interactions of the cationic polyamines with negat-
ively charged phosphate groups of nucleotidic macromole-
cules are considered to be of great biological importance. In
particular, the interaction of polyamines with DNA
induces important conformational modifications in DNA
structure [10].
In a previous study, we aimed to investigate the fate of
putrescine when taken up from the medium of Caco-2 cells
and to analyse its binding to nuclear proteins. We reported
the presence of compounds with molecular masses of about
8000, 4800 and < 1000 Da (actually, named 180 Da) in the
nuclear extracts of replicating cells. In contrast, nuclear

extracts of differentiated Caco-2 cells lacked the 4800 Da
compound. It was shown that these compounds, detected
by gel permeation chromatography (a separation technique
that does not alter the molecular interactions) were able to
establish noncovalent bonds with the exogenous radioactive
polyamines. We hypothesized that these compounds were
oligopeptides [11].
Our aim in the present work was to: (a) better define the
chemical structure of these nuclear compounds, herein
named NAPs (nuclear aggregates of polyamines); (b) study
their fluctuating concentrations during the various phases of
Caco-2 cell replication induced by gastrin treatment; (c)
ascertain their presence in other replicating cell lines; and (d)
investigate the effects of NAP–DNA interaction on DNA
conformation and DNA protection by means of spectro-
photometric and electrophoretic analyses.
MATERIALS AND METHODS
Cells
Pre-confluent (replicating) Caco-2 cells at day 6 of culture
were used for the experiments [7]. In order to favour cell
synchronization, Caco-2 cells were left without changing the
media for 60 h. Cell replication was promoted by adding
10
)10
M
gastrin (ICN) to the dishes. Nuclear extracts of cells
treated with gastrin for 0, 2, 4, 8 and 12 h were fractionated
by gel permeation chromatography (GPC). The GPC peaks
with the molecular masses of 8000, 4800 and < 1000 Da
were collected and analysed for the detection of Fmoc

derivative polyamines using reversed phase-HPLC. The
replication rate of these cells was evaluated by assessing
bromodeoxyuridine (BrdU) incorporation, an S-phase
marker [12].
NAP formation was also investigated in Caco-2 cells
starved for 4 days.
The following replicating cells, generously donated by
other laboratories, were also used for NAP isolation: the
Correspondence to L. D’Agostino, Facolta
`
di Medicina,
Via S. Pansini 5, 80131, Napoli, Italy. Fax/Tel.: +39 81 746 2707,
E-mail:
Abbreviations: GPC, Gel permeation chromatography; NAP, nuclear
aggregate of polyamines; BrdU, bromodeoxyuridine.
(Received 26 March 2002, revised 24 June 2002, accepted 19 July 2002)
Eur. J. Biochem. 269, 4317–4325 (2002) Ó FEBS 2002 doi:10.1046/j.1432-1033.2002.03128.x
primary cultures chicken embryo chondrocyte, chicken
embryo fibroblast and quail embryo chondrocyte, and the
cell lines KB human epidermoid oropharingeal carcinoma,
PCCl3 rat thyroid and NA101 chicken embryo chondrocyte
transformed by RSV. These cells were cultured in the
recommended standard conditions and used when pre-
confluent.
Nuclei and nuclear extract preparations
Cells were solubilized in solution 1 (15 m
M
NaCl, 60 m
M
KCl, 14 m

M
2-mercaptoethanol, 2 m
M
EDTA, 15 m
M
Hepes pH 7.9, 0.3
M
sucrose) containing 1% Triton-X100
and phenylmethanesulfonyl fluoride. The crude nuclear
pellet was prepared by spinning the extracts at 3500 r.p.m.
for 10 min at 4 °Cona1-
M
sucrose cushion in solution 1.
The purity of nuclei preparations was tested by light
microscopy after Crystal violet staining. Nuclear extracts
were prepared as described [13]. The nuclear pellet was
re-suspended in high-salt concentration (NaCl 400 m
M
)
solution 1 and centrifuged at 10 000 g for 10 min.
GPC
The nuclear extracts were analysed by GPC–HPLC using a
Superose 12 prepacked column HR 10/30, which has a
separation range of 1000–300 000 Da (Pharmacia). The
column was equilibrated with 0.05
M
sodium phosphate
buffer (pH 7.2) containing 0.15
M
NaCl and calibrated

using compounds with varying molecular masses, as
indicated by the manufacturer. Fifty lL of the nuclear
extracts were diluted in equal volume of equilibration buffer
and loaded onto the column. The nuclear extracts were
eluted with the same buffer at 0.4 mLÆmin
)1
anddetectedat
280 nm. The single GPC peaks with a molecular mass
< 10 000 Da were collected and stored at )20 °C. The
GPC analysis allowed the study of the nuclear extracts in
native conditions and in the absence of strong interactions
(electric field or denaturing and reducing conditions), which
disrupt noncovalent bonds. Therefore, it was our sole
possible choice.
RP-HPLC Fmoc-polyamine derivatives
The presence of polyamines in the GPC peaks was analysed
by RP-HPLC using a precolumn Fmoc derivatization [14].
The excitation wavelength was set at 265 nm and fluores-
cence emission was monitored at 305 nm to increase
sensibility in the Fmoc derivative analyses.
Amino acid analysis of GPG peaks by RP-HPLC
of Fmoc derivatives
The presence of oligopeptides and/or free amino acids was
excluded by performing the amino acid analyses by
RP-HPLC of Fmoc derivatives before and after the
hydrolysis of GPC peaks. Dried GPC peaks were dissolved
in 500 lL6
M
HCl. Each solution was put into vacuum
hydrolysis tubes (Pierce, Rockford, IL, USA), gassed with

nitrogen and sealed. The tubes were incubated at 110 °Cfor
24 h in a Reacti-Therm for dry block heating apparatus
(Pierce). Derivatization of amino acids with Fmoc and their
RP-HPLC analysis were both performed as described [15].
The wavelength excitation was set at 265 nm and fluores-
cence emission was monitored at 305 nm.
Spectrophotometric scan of NAPs
One ml of each NAP, obtained from GPC collection of
Caco-2 cells nuclear extracts, was scanned at room
temperature from 400 to 190 nm at 10 nmÆs
)1
by a Cary
spectrophotometer 1E series (Varian Inc., Walnut Creek,
CA, USA).
In vitro
aggregation of polyamines
The in vitro aggregation of polyamines was studied dissol-
ving putrescine, spermidine and spermine (Sigma) at equal
molar concentrations in 0.05
M
sodium phosphate buffer
(pH 7.2) containing 0.15
M
NaCl to obtain a mixture with a
final concentration of 25 l
M
. This polyamine solution was
then analysed by GPC.
In order to assess the role of spermine in NAP formation,
the concentration of this 25 l

M
polyamine solution was
brought to 50 l
M
by adding spermine and then performing
a new GPC run. The GPC analyses were carried out as
described above, using as mobile phase phosphate or
Tris/HCl buffers.
NMR analysis
Putrescine, spermidine and spermine were dissolved in D
2
O
or in D
2
O phosphate buffer (0.05
M
pH 7.2, containing
0.15
M
NaCl) at a concentration of 10 mgÆmL
)1
.
All spectra were recorded by a Bruker DRX-600 NMR
spectrometer, operating at 599.19 MHz for
1
H, using the
UXNMR
software package; 1D-TOCSY experiments were
carried out using the conventional pulse sequences, as
described [16].

NAP–DNA spectrophotometry
Spectrophotometric assays were performed by mixing
200 lL of the 8000, 4800 and < 1000 Da (the most
retained) peaks with 100 lL of human genomic DNA in
Tris/EDTA buffer (1.3 lgÆlL
)1
). This solution was brought
to a volume of 800 lL with 0.05
M
sodium phosphate buffer
(pH 7.2) to obtain a concentration of 0.25 ng total
polyamineÆlg
)1
DNA ratio. The absorbance (A)ofeach
NAP–DNA sample was measured with a thermostated
Cary spectrophotometer 1E series (Varian Inc., Walnut
Creek, CA, USA) at 260 nm after 6 min incubation at 15,
37 and 55 °C. Controls were NAP solutions in the absence
of DNA or single polyamines at 1 l
M
concentration in
water.
DNA electrophoresis
Electrophoresis of human genomic DNA or 1 kb DNA
ladder (Sigma-Aldrich) was carried out in a HE 100
supersub (Amersham Pharmacia Biotech) at a constant
temperature of 37 °C applying an electric field strength of
11.1 VÆcm
)1
in Tris/borate/EDTA.

Ten lL of a mixture of genomic DNA and 8000 or 4800
or < 1000 NAP (0.25 ng total polyamineÆlg
)1
DNA) were
loaded on a 1.5% ultrapure DNA grade agarose gel after an
4318 L. D’Agostino and A. Di Luccia (Eur. J. Biochem. 269) Ó FEBS 2002
incubation period of 6 min at 37 °C. The final concentra-
tion of DNA was 0.4 lgÆlL
)1
. Ethidium bromide buffer,
0.1 lgÆmL
)1
, was added to the gel and to the electrophoresis
buffer. The duration of electrophoresis was 3.5 h.
The influence of NAPs on the electrophoretic mobility of
small linear fragments of DNA was evaluated using a
241 base pair PCR product of the BRCA 1 gene. The
electrophoretic conditions were the same as those used for
the genomic DNA.
Two microliters of 1 kb DNA ladder (200 lgÆlL
)1
)were
mixedwith3lL of the 8000, 4800 or < 1000 NAP. These
solutions were incubated at 37 °Cfor6min,5lL
exonuclease III (65 UÆlL
)1
) were added and incubation
was continued for 30 min. The samples were then separated
by electrophoresis for 1 h in the conditions described above.
Genomic DNA (4 lgper2.5lL phosphate buffer) was

incubated for 6 min at 37 °C with 4.5 lL of 8000, 4800
or < 1000 NAPs (mean polyamine concentration:
0.25 ngÆlg
)1
DNA) or aqueous solutions of single polyam-
ines (0.25 ngÆlg
)1
DNA). The degradation of the genomic
DNA was examined by means of DNase I (RQ1RNase-free
DNase, Promega) at concentration of 0.025 UÆlg
)1
DNA.
Briefly, 1 lL of the DNase I solution was added to 1 lLof
the reaction buffer solution (400 m
M
Tris/HCl at pH 8,
100 m
M
MgSO
4
,10 m
M
CaCl
2
)andthenmixedwithNAP–
DNA or polyamine–DNA solutions. Enzyme action was
stopped after 30 min at 37 °C adding 1 lL20m
M
EDTA
pH 8. Samples were then separated by electrophoresis for

1 h using the conditions described above.
Each gel was photographed with a Polaroid MP-4 L
camera and migration distances were measured with a ruler
from photographs.
Statistics
Differences in polyamine concentrations among NAPs were
tested for significance by one-way
ANOVA
with Bonferroni
test for multiple means comparisons using
SPSS
software
package for
WINDOWS
, release 10.0.7. Values were consid-
ered significant at P < 0.05.
RESULTS
GPC of nuclear extracts of replicating cells
A representative profile of GPC analysis of the nuclear
extracts of Caco-2 cells stimulated to replicate by gastrin is
shown in Fig. 1. The chromatograms showed three peaks.
The molecular masses of first two were estimated to be 8000
and 4800 Da. The third peak, the most retained, fell out of
the column separation range and, for this reason was
marked as < 1000 Da. Compelling variations in GPC
peaks were recorded after 2, 4, 8 and 12 h of gastrin
treatment. The chromatograms at time 0 showed two minor
peaks corresponding to 8000 and 4800 Da (19.6 and 16.1%,
respectively) and a major one corresponding to < 1000 Da
(64.3%). Two hours after gastrin treatment, there was a

huge increase in the 4800 Da peak area value (62.2%). This
peak declined at 4 and 8 h and returned to the initial value
after 12 h of gastrin stimulation. The 8000 Da peak
increased at 4 h (34.1%), remained the same at 8 h and
declined at 12 h. The < 1000 Da peak strongly decreased at
2 and 4 h (18.9 and 19.8%, respectively) after which it
progressively increased, reaching the basal value in the final
stage of observation. The S phase entrance values indicated
that gastrin promotes cell replication: before gastrin
treatment, 31% of Caco-cells incorporated BrdU, whereas
after 2 and 4 h 46% and 50% incorporated BrdU,
respectively. An increased BrdU incorporation (40%) was
recorded as long as 8 and 12 h after gastrin treatment.
The GPC analysis performed on the nuclear extracts of
Caco-2 cells starved for 4 days revealed a very low
< 1000 Da peak at the initial conditions (0 h), and the
retarded and scarce formation of 4800 NAP at 4 and 8 h.
The 8000 Da NAP was essentially unaffected by prolonged
starvation (data not shown).
The GPC analyses were also performed on the nuclear
extracts of primary cultures, chicken embryo chondrocyte,
chicken embryo fibroblast and quail embryo chondrocyte,
and the cell lines KB human epidermoid oropharingeal
carcinoma, PC Cl 3 rat thyroid and NA101 chicken embryo
chondrocyte transformed by RSV. In the chromatograms of
these replicating cells, the 8000, 4800 and < 1000 Da peaks
were always distinguishable (data not shown).
Analysis of polyamine and amino acid by RP-HPLC
of purified GPC peaks
The molar concentrations of polyamines forming NAPs are

shown in Table 1: statistical differences were due to the
lower total polyamines, spermine and spermidine concen-
trations of < 1000 NAP with respect to those of 4800 NAP.
Polyamine molar concentrations allowed us to define the
Fig. 1. Gel permeation chromatography of nuclear extracts from Caco-2
cells at 0, 2, 4, 8 and 12 h following treatment with 10
)10
M
gastrin. The
cells used for the experiment were preconfluent and starved for 60 h.
Each chromatographic run was performed using the fourth part of the
entire nuclear extract of 10
6
cells. The modifications in the 4800 and
< 1000 Da peaks showed an inverse trend after gastrin stimulation.
Minor modifications were observed in the 8000 Da peak.
Ó FEBS 2002 Nuclear aggregates of polyamines and DNA (Eur. J. Biochem. 269) 4319
simplest formulae of NAPs. The concentration of phos-
phates was calculated considering that they have, at
physiological pH, two negative charges (pK
a1
¼ 2.12;
pK
a2
¼ 7.21). Thus, we estimated there were two moles of
polyamines per mole of phosphate.
NAPs extracted from the nuclei of the other cell types had
analogous polyamine composition (data not shown).
Acid hydrolysis ensured the absence of amino acids and
peptidic amino acid residues in NAP composition. The

RP-HPLC profiles of Fmoc-derivatives before and after
acid hydrolysis were, in fact, identical and showed only the
typical polyamine peaks, and not any added and/or
increased peaks that could indicate the presence of peptides
(data not shown).
Polyamine aggregation studies
The absence of oligopeptides and free amino acids, partic-
ularly those exhibiting an absorbance at 280 nm, prompted
us to investigate the absorbance range. We therefore
scanned NAPs between 400 and 190 nm. The maximal
absorbance peak of NAPs obtained from Caco-2 cells was
at 200 nm. Moreover, a lower peak, ranging from 240 to
290 nm, with the maximum at 265 nm, was observed. The
height of this peak was different in each NAP, being lowest
in < 1000 NAP and highest in the 4800 NAP. The absence
of a shoulder at 220 nm confirmed the absence of peptide
bonds (data not shown).
Because both results of acid hydrolysis and spectropho-
tometric scansions were inconsistent with a peptidic struc-
ture of NAPs, we supposed that these compounds could be
formed by the interaction and aggregation between phos-
phates and polyamines. This hypothesis was tested by
performing GPC chromatography of a 25 l
M
polyamine
mixture in phosphate buffer pH 7.2, using tris(hydroxy-
methyl)aminomethane/HCl or phosphate buffer as the
mobile phase (Fig. 2A). Whatever the buffer used, the
GPC profiles that resulted after the Ôin vitroÕ aggregation of
polyamines was very similar to those obtained by analysing

the nuclear extracts of replicating cells. Furthermore, a huge
increase in the 4800 Da GPC fraction and the formation of
intermediate compounds with molecular mass ranging from
<1000 to 4800 Da occurred when polyamine solution was
brought to 50 l
M
by the addition of 125 l
M
spermine
(Fig. 2B).
NMR was used for analysis of the effects of polyamine–
phosphate interaction on the molecular arrangement of
polyamines. In Fig. 3, the NMR-spectra of the single
polyamines (putrescine, spermidine and spermine) dissolved
in D
2
O(A)orinD
2
O phosphate buffer (B) are shown. The
signals of polyamines in D
2
O phosphate buffer show
chemical shifts of about 0.05 p.p.m. higher than the signals
of the same polyamines dissolved in D
2
O. The CH
2
proton
resonances of putrescine dissolved both in D
2

Oandin
phosphate buffer were represented by two single peaks at
2.9 p.p.m. (peaks 2) determined by the protons of methylene
adjacent to the NH
2
terminus and at 1.7 p.p.m. (peaks 1)
due to the b CH
2
protons. In addition to these two signals,
spermidine and spermine gave signals that fall in the
resonance field of 2.0–2.1 p.p.m. determined by the methy-
lene protons in position b included between nitrogen
Fig. 2. In vitro aggregation of polyamines. (A) GPC profile of equal
molar concentrations of putrescine, spermidine and spermine (25 l
M
final concentration) dissolved in phosphate buffer (pH 7.2). The GPC
profile was the same when either the phosphate buffer or a tris
(hydroxyl methyl)aminomethane/HCl buffer was used as the mobile
phase. (B) GPC was repeated after the addition of 125 l
M
spermine to
this polyamine solution, resulting in a huge increase in the 4800 Da
peak. This result indicates that spermine concentration is a determin-
ant of the formation of this compound. This experiment demonstrates
that it is possible to realize an in vitro aggregation of polyamines that
gives rise to compounds with molecular masses identical to those of the
natural aggregates extracted from nuclei of replicating cells.
Table 1. Concentration of polyamines (nmolÆmL
)1
) in the nuclear aggregates of polyamines (NAPs) extracted from nuclei of replicating Caco-2 cells.

The results are expressed as mean ± SD of four determinations carried out on the GPC eluted peaks. The means marked by different letters differ
for P < 0.05 by Bonferroni test. Ph, Phosphate group.
8000 NAP 4800 NAP < 1000 NAP P (one-way
ANOVA
)
Putrescine (Put) 0.272 ± 0.141 0.307 ± 0.044 0.153 ± 0.053 N.S.
Spermidine (Spd) 0.256 ± 0.109
a,b
0.448 ± 0.166
a
0.187 ± 0.085
b
0.04
Spermine (Spm) 0.360 ± 0.153
a,b
0.618 ± 0.207
a
0.306 ± 0.082
b
0.04
Total 0.888 ± 0.299
a,b
1.373 ± 0.379
a
0.646 ± 0.182
b
0.02
Simplest formula Put-Ph-Spd-Ph-Spm Put-Ph-Spd-(Ph-Spm)
2
Put-Ph-Spd-(Ph-Spm)

2
4320 L. D’Agostino and A. Di Luccia (Eur. J. Biochem. 269) Ó FEBS 2002
(peaks 3) and of 2.9–3.1 p.p.m. due to methylene protons
adjacent to NH (peaks 4 and 5). Furthermore, spermine
showed a singlet at 1.9 p.p.m. (peaks 6).
Different profiles were recorded in the spectra of
spermine and spermidine, when dissolved in D
2
Oorin
D
2
O phosphate buffer: peaks 3, 4 and 5 showed differences
in multiplicity of signals and in peak wideness.
The observed variations in chemical shift and in the
width, shape and number of signals, due to the different
proton exchange and the spin–spin coupling, are represen-
tative of the interaction between phosphate groups and
polyammonium cations and, in the case of spermidine and
spermine, could be indicative of their different conforma-
tional arrangement in phosphate buffer.
Effects of NAP–DNA interaction
To consider the likely relevance of NAPs and DNA
interaction to DNA conformation and, in particular, to
assess the effects of this interaction on the exposition of the
inner bases, NAP–DNA solutions were evaluated by
measuring A at 260 nm and at different temperatures. The
A values of the different NAP–DNA solutions, monitored
for 6 min at 15, 37 and 55 °C, are shown in Fig. 4A. Each A
value was calculated by subtracting the A value of the DNA
from those of the NAP–DNA solutions. Only the 4800

NAP–DNA solution showed an isolated huge A increase at
37 °C (0.7 absorbance units), while no absorbance variations
were recorded in the 8000 and < 1000 NAP–DNA solutions
at the three temperatures. The highest A value of 4800 NAP–
DNA solution at 37 °C was reached in about 10 s.
To exclude that the variation of absorbance was due to
NAPs and not to DNA conformational changes, NAP
solutions were evaluated in the absence of the genomic
DNA: we did not observe any modification in the O.D.
values at the different temperatures of the experiment (data
not shown). Furthermore, the absorbance of genomic DNA
solutions did not change in presence of 1 l
M
single
polyamines, a concentration similar to that of polyamines
forming the NAPs (data not shown).
These spectrophotometric results motivated us to investi-
gate the electrophoretical behaviour of NAP–genomic
DNA solutions, in view of the fact that a different
electrophoretic mobility of DNA on agarose gel can suggest
a modification of DNA conformation. The electrophoretic
pattern of the NAP–DNA solutions on a 1.5% agarose gel
is shown in Fig. 4B: lane C, corresponding to the 4800
NAP–DNA, illustrates a faster migrating DNA band
compared to lanes B and D corresponding to 8000 and
< 1000 NAP–DNA solutions. A temperature of 37 °Cwas
essential to the visualization of the fastest migration of 4800
NAP–DNA. When this electrophoretic experiment was
repeated using the 241 base pair DNA fragment, no
significant difference in the migration of NAP–DNA

oligomer solutions was found (data not shown).
To identify the sites of interaction of NAPs on the DNA
strands, the 1 kb DNA ladder fragments preincubated with
the single NAPs were exposed to the phosphodiesterasic
activity of exonuclease III, assuming that the sparing of
DNA fragments from degradation was due to the impedi-
ment of the enzyme action on the DNA strand phospho-
diester bridges that were occupied by NAPs (Fig. 5).
The degradation of the 1 kb DNA ladder by exonuc-
lease III was strongly impeded by previous incubation with
NAPs. In fact, in the absence of these compounds, the
enzymatic digestion was complete for the small fragments
Fig. 3. NMR spectra of polyamines.
1
Hspec-
tra of polyamines (putrescine, spermidine and
spermine) dissolved in (A) D
2
Oor(B)D
2
O
phosphate buffer (50 m
M
, pH 7.2, containing
0.15
M
NaCl). The concentration of polyam-
ines was 10 mgÆmL
)1
.Peaks4and5ofsper-

mine and 2, 4 and 5 of spermidine changed in
chemical shift and signal multiplicity when
these polyamines were dissolved in phosphate
buffer. Only differences in chemical shifts were
recorded for putrescine (data not shown).
Ó FEBS 2002 Nuclear aggregates of polyamines and DNA (Eur. J. Biochem. 269) 4321
up to 1018 bp and partial for those of 1636 and 2036 bp
(lane B). In contrast, when exonoclease III incubation was
preceded by the interaction of each NAP with the DNA,
degradation of these bands was strongly reduced (lanes C,
D and E). Single polyamines, used as controls at a
concentration (1 l
M
) similar to that of polyamines forming
the NAPs, did not show any protective effect.
Degradation of genomic DNA by DNase I, a phospho-
diesterasic endonuclease, was strongly reduced by previous
incubation with each NAP (Fig. 6A). In contrast, the
incubation of genomic DNA with 1 l
M
spermine, spermi-
dine or putrescine solutions did not provide any relevant
protective effect (Fig. 6B).
DISCUSSION
In the present study we have demonstrated that in the nuclei
of replicating Caco-2 cells, and of all the other replicating
cells tested ) i.e. epithelial or mesenchimal, mutated or
nonmutated ) polyamines aggregate with phosphate ani-
ons by ionic bonds to form three molecular structures with
estimated molecular masses of 8000, 4800 and < 1000 Da,

uncharacterized so far. Owing to their positive charge, these
NAPs interact with the phosphate groups of the DNA
strands. Compelling modifications in the concentration of
NAPs paralleled the mitogenic effects produced in Caco-2
cells by gastrin. In particular, the 4800 NAP was apparently
closely linked to the process of cell replication, as this NAP
reached maximal concentration in the few hours following
gastrin stimulation. The importance of 4800 NAP in cell
replication is also strongly supported by its absence in the
differentiated Caco-2 cells, which lost their replicating
activity when confluence was reached [11]. The 8000 NAP
was well represented in both replicating and differentiated
Caco-2 cells [11], and its concentration did not vary much
during Caco-2 cell replication.
The biological role of 8000 NAP, which is presumably
not played in the cell replication, is vaguely definable from
our study. However, other important interactive functions
such as single-strand DNA stabilization and/or DNA
repair and protection can be postulated for this NAP.
Fig. 4. Absorbance (A) values and electrophoretic patterns of NAP–
DNA solutions (r, 8000 NAP-DNA; j, 4800 NAP-DNA; m, <1000
NAP-DNA). (A) The 4800 NAP–DNA solution showed the highest A
values, which reached the maximum at 37 °C. Intermediate A values,
unchanged by temperature variations, were recorded for the 8000
NAP–DNA solution. The < 1000 NAP–DNA solution did not show
any absorbance value. A values were monitored for 6 min at 260 nm at
different temperatures and calculated by subtracting the A value of the
DNA from those of NAP–DNA solutions. Human genomic DNA
was used. NAP solutions without DNA did not show any variation in
A at the different temperatures used. Furthermore, single polyamines,

used as control at a concentration equivalent (1 l
M
)tothatofpoly-
amines composing the NAPs, did not cause any variation in A (data
not shown). (B) Lane A corresponds to the migration of human
genomic DNA. Lanes B, C and D show the same DNA preincubated
for 6 min at 37 °C with 8000 and 4800 and < 1000 NAPs, respectively.
A faster migration of the DNA in presence of 4800 NAP was shown.
Electrophoresis was performed in 1.5% agarose gel in Tris/borate/
EDTA and at a constant temperature of 37 °C.
Fig. 5. Effect of exonuclease III on the electrophoretic migration of
NAPs)1 kb DNA ladder. Lane A, 1 kb DNA ladder; lane B, the same
DNA fragments incubated for 6 min at 37 °C with exonuclease III.
Migration of the 8000, 4800 and < 1000 NAP)1kb DNA ladder
solutions incubated with exonuclease III are shown in the lanes C, D
and E, respectively. In the absence of NAPs (lane B), enzymatic
degradation was partial for the DNA fragments of 2036 and 1636 bp
and complete for the smaller ones. NAPs conferred huge protection
against degradation by exonuclease III to the DNA fragments: bands
< 506 bp are faintly visible, while those of 1018 and 506 bp remain
evident. In the same lanes, only a negligible diminution in the intensity
of the bands corresponding to DNA fragments of > 1018 bp can be
appraised. Electrophoresis was for 1 h in a 1.5% agarose gel. The
sparing of DNA fragments from degradation, due to inhibition of the
enzyme action on the DNA strand phosphodiester bridges occupied by
NAPs, indicates that this is the site of NAP–DNA interaction. Single
polyamines, used as control at a concentration equivalent (1 l
M
)to
that of polyamines composing the NAPs, did not show any protective

effect(datanotshown).
4322 L. D’Agostino and A. Di Luccia (Eur. J. Biochem. 269) Ó FEBS 2002
In particular, a protective role is strongly suggested by its
inhibitory effect on the action of both exonuclease III and
DNase I, an effect exerted by the other NAPs also.
Variations in the < 1000 NAP by GPC analyses indicate
that this compound functions as a store of material to be
used in the formation of 4800 NAP: the diminution of
< 1000 NAP occurs simultaneously with an increase in
4800 NAP. Its highly concentrated presence in the nuclei
appears to be determinant of 4800 NAP production. In fact,
prolongation of starvation for up to 4 days ) conditions
that determine a strong reduction in the concentration of
polyamines in the cells [17] ) caused a strong decrease in
< 1000 NAP and a reduced and delayed production of the
4800 NAP in the 8 h following gastrin treatment. These
data indicate that the rapid formation of 4800 NAP occurs
only in conditions of efficient polyamine metabolism
assuring the maintenance of adequate amounts of < 1000
NAP monomers in the nuclei.
The hypothesis that < 1000 NAP is a precursor of 4800
NAP is also supported by in vitro aggregation studies that
clearly demonstrated that the adjunct of spermine to the
polyamines dissolved in phosphate buffer determines
the increase in the 4800 Da GPC peak, paralleled by the
disappearance of the < 1000 Da peak. Therefore, it is
possible to hypothesize that whenever the concentration of
spermine, and probably also that of spermidine, increase in
the cells, they interact with the < 1000 Da compounds to
form new 4800 Da compounds with the consequent

diminution of those of < 1000 Da.
In spite of the wide biological differences of the cell types
we tested, the NAPs were detected in all nuclear extracts,
thus indicating that nuclear polyamine aggregation could be
regarded as a general biological phenomenon.
The NAPs are rich in spermine and spermidine. The
appearance of these two polyamines in the nucleus is crucially
important for the induction of cell mitosis [1–4]. Because of
their high positive charge, spermine and spermidine interact
with the DNA phosphates by means of nonspecific electro-
static binding [18]. The induction of B-Z DNA conforma-
tion, an important structural modification that prepares
DNA for replication, was also accounted for by spermine
and spermidine in various studies [19–21]. Vajayanathan
et al. recently investigated DNA condensation by polyam-
ines using static and dynamic laser light scattering tech-
niques. In particular, the structural modifications determined
by spermine and a series of its homologues on k-DNA were
assessed. These authors found that the variation of the
number of methylene spacings in the bridging region between
the two secondary amino groups of spermine had a profound
effect on the molecule’s ability to provoke structural changes
in DNA and concluded that the regio-chemical distribution
of positive charge along polyamines plays a major role in the
condensation of DNA [22].
Our data are consistent with the findings cited above, but
further suggest that the interaction of polyamines with the
genomic DNA is more complex, as these polycationic
compounds have an extraordinary intrinsic tendency to
form molecular aggregates, generating ionic bonds with the

phosphate anions present both in buffer solutions or
biological media and in DNA strands.
The interaction of 4800 NAP with DNA determined
interesting changes in both absorbance and electrophoretic
migration of whole DNA, probably as a consequence of
induced DNA conformational modifications. The UV data
indicate that the interaction of the whole DNA with this
NAP is able to determine, at physiological temperature, the
increased exposition of the bases, which, in usual experi-
mental conditions, can be obtained only by denaturing the
DNA at high temperatures or exposing it to a very high pH
[23]. Furthermore, the rapid increase in A values of the 4800
NAP–DNA solution at 37 °C is in accordance with a model
of gradual and oriented B-Z transition starting at (CG)
5
motif: as the superhelical stress increases, the Z-structure
propagates along the remaining part of the repeat by
successive transitions until the full-length sequence is
converted [24]. Similarly, the faster electrophoretic mobility
of the 4800 NAP–genomic DNA solution at 37 °C,
depending on the enhanced permeation into the gel showed
Fig. 6. Electrophoresis of human genomic DNA incubated with NAPs
or single polyamines and digested by DNase I. (A) m: 1 kb DNA ladder
DNA; lanes A and B, DNA and DNA + DNase I, respectively; lane
C, DNA + 8000 NAP + Dnase I; lane D, DNA + 4800 NAP +
Dnase I; lane E, DNA + < 1000 NAP + DNase I. (B) Lanes A
and B, DNA and DNA + DNase I, respectively; lane C, DNA +
spermine + Dnase I; lane D, DNA + spermidine + DNase I; lane
E, DNA + putrescine + DNase I. Electrophoresis of the DNA was
carried out for 1 h in a 1.5% agarose gel at 37 °C. The protective effect

of NAPs or single polyamines on genomic DNA was assayed after
their incubation with DNA and the successive exposition of these
solutions to DNase I. NAPs were shown to be more efficient than
1 l
M
single polyamines in protecting genomic DNA from DNase I
degradation. This polyamine concentration was equivalent to that of
polyamines composing NAPs.
Ó FEBS 2002 Nuclear aggregates of polyamines and DNA (Eur. J. Biochem. 269) 4323
by the condensed molecules [25,26] is also indicative of a
strong rearrangement of the DNA double helix, probably
caused by the closing of phosphate groups in GC rich
regions and the DNA condensation [27,28]. In the case of
the 241 base pair DNA fragment, we did not observe
significant differences in its electroforetic mobility in the
presence of NAPs. This result confirms that the ratio
between the molecular dimension and gel matrix sieving is
crucial for the detection of the DNA conformational
changes, as already demonstrated by others [26]. Thus, we
believe that these evidences are not in contradiction with our
conclusion that, namely, the electrophoretic pattern deter-
mined by the interaction of 4800 NAP with the genomic
DNA is due to a remarkable DNA conformational
modification.
Unlike 4800 NAP, 8000 and < 1000 NAPs did not
change either absorbance or electrophoretic migration
properties of DNA. However, as all of the NAPs are able
to interact with the DNA, as inferred by the fact that
incubation of DNA with these compounds hindered its
degradation by exonuclease III and DNase I, other inter-

active functions can be proposed, such as DNA protection
and/or stabilization and packaging.
The role of NAPs in DNA protection seems to be
relevant, as they were able to protect the genomic DNA
from DNase I digestion with an efficacy hugely superior to
that shown by single polyamines. These data indicate that
NAPs are probably crucial in the defence of DNA in the
case of inappropriate activation of the apoptotic process
[29,30].
As an ulterior aim of our study, we intended to shed light
on the NAP molecular structure. The GPC analysis of
standard polyamines dissolved in phosphate buffer revealed
the formation of compounds with a molecular mass
identical to those of natural NAPs. These results suggest
that NAP supramolecular arrangement derives from the
simple aggregation between the negatively charged phos-
phates and the positively charged polyamines. NMR studies
performed on the polyamines dissolved in phosphate buffer
clearly indicate that the ionic interaction between phosphate
groups and polyammonium cations of spermidine and
spermine induce their molecular rearrangement. We sup-
pose that this new molecular conformation can favour
aggregation among the polyamines and the phosphates,
with the consequent generation of NAPs. Our belief is
supported by evidence from several NMR studies that
demonstrated the formation of complexes among polyam-
ines and phosphorylated molecules [31–33]. The formation
of a polymeric aggregation of polyamines and phosphates
can be also inferred by our spectrophotometric analyses
showing that these compounds gave two absorbance peaks

at 200 and 265 nm. These absorbance values can be
attributed to an electron delocalization similar to that of
the pfip* transition appearing in the spectra of molecules
with pfip bond conjugated structure such as polyene
systems or aromatic structures. Therefore, it can be argued
that the NAP monomers (Table 1) tend to assume a sort of
cyclic structure by phosphate bridges, and form a polymeric
aggregation of a reticular appearance in which electrons are
delocalized in a p-like status, determining different e for
each NAP. This phenomenon explains their detection at
280 nm in GPC analyses.
The aggregation of polyamines and phosphates ascribes
to NAPs a positive charge due to a 2 : 1 polyamine : phos-
phate ratio. Thus, these compounds have a considerable
number of free positive charges available for interaction
with the negatively charged molecules. Our results are all in
favour of the interaction of NAPs with the DNA. Theor-
etically, this phenomenon should be consequent to the
exposition of NAP positive charges to the DNA phosphate
groups. The hindrance to the phosphodiesterasic DNA
degradation exerted by these compounds indicates that the
DNA phosphate groups are really the negative ions
involved in this kind of interaction. Furthermore, as DNA
integrity has been shown to be assured by NAPs with an
efficiency much higher than that exerted by single polyam-
ines, it is possible to suppose that, as already suggested for
polyamines [34–36], the size and probably also the shape of
each NAP are important for its appropriate arrangement in
the DNA groove.
In conclusion, we have demonstrated that in the nuclei of

several replicating cell types there exist molecular aggregates
that are able to interact with DNA phosphate groups, i.e.
NAPs, whose quasi-stable structure is based on ionic bonds
between polyamines and phosphate groups.
In our opinion, the identification of these compounds,
which are able to induce DNA conformational changes and
DNA protection, does not contradict the generally accepted
functions of polyamines, but instead provides a better
understanding of several aspects of the interaction of
polyamines with DNA.
ACKNOWLEDGEMENTS
We are grateful to the following for their generous gift of cells: Drs
E. Gionta, G. Pontarelli, M. Santoro and P.S. Tagliaferri. We wish also
to thank Drs A. Menchise, C. Verde, C. Salzano, M.V. Barone, M. Di
Pietro, N. De Tommasi and A. Contegiacomo, and Mr P. Mastranzo
for their help in performing parts of the experiments. Our work was
supported by a research grant from the Campania Region.
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