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: luciano@unina.it. 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. 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Biochemistry 32, 14068–14074. Ó FEBS 2002 Nuclear aggregates of polyamines and DNA (Eur. J. Biochem. 269) 4325 . 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;