DNA modification in chick heart and cerebrum

Comparative Biochemistry and Physiology, Part A 138 (2004) 147 – 160 www.elsevier.com/locate/cbpa

DNA modification in chick heart and cerebrum K. Kagawa a,*,1, H. Kagawa b a

Department of Biological Responses, Institute for Virus Research, Kyoto University, Shogoin-kawahara-cho, Sakyo-ku, Kyoto 606-8507, Japan b Department of Physics, Kyoto University, Kitashirakawa, Sakyo-ku, Kyoto 606-8502, Japan Received 30 July 2003; received in revised form 27 January 2004; accepted 28 January 2004

Abstract Heart muscle cells and cerebral neurons are known to lose the ability to proliferate and are called terminally differentiated cells. They are generated in appropriate numbers during embryogenesis and retained throughout adult life without turnover. We are interested in such a longlived DNA. We isolated DNA from chick heart and cerebrum and compared it with DNA from other organs after incubation with DNase I. Single-strand breaks were assessed using a reaction system composed of DNA and Escherichia coli DNA polymerase. The DNA of both organs was relatively resistant to DNase I, and DNA modification occurred during embryogenesis. CIMS (chemical ionization mass spectrometry) indicated that the molecular mass of the deoxynucleoside of both DNAs was larger than that of the corresponding canonical deoxyribonucleoside by m/z 28 (or 30 for the protonated form). The difference between these deoxynucleosides is based on a difference in sugar constituents. Cerebral deoxynucleotides were analyzed by 13C NMR. An extra signal near 173 ppm was observed, which was assigned to the amide carbonyl. We propose a model of the deoxynucleoside where a carbonyl residue exists between the base and the 2-deoxyribose moiety of the canonical deoxyribonucleoside. D 2004 Elsevier Inc. All rights reserved. Keywords: Cardiac DNA; Cerebral DNA; 3-Deoxyglucosone; 3-Deoxyhexonic acid; DNA modification; Modified deoxynucleoside; Modified 2-deoxyribose; Modified DNA

1. Introduction During embryonic development, the cells of the body become differentiated, but not all populations of differentiated cells are subject to turnover and renewal. Muscle cells of the heart and cerebral neurons are known to lose the ability to proliferate and are called terminally differentiated cells. They are generated in appropriate numbers during embryogenesis and retained throughout adult life without turnover (Manasek, 1968; Alberts et al., 1983; Olson and Srivastava, 1996). For example, in the chick embryo, the cell division of cardiomyocytes ceases by day 11 (Manasek, 1968) and that of cerebral neurons by day 9 (Dunn and Bondy, 1974). These cells have an Abbreviations: CIMS, chemical ionization mass spectrometry; MS, mass spectrum; SIM, selective ion monitoring; TIC, total ion current; RI, refractive index. * Corresponding author. Tel.: +81-75-751-4027; fax: +81-75-7615626. E-mail address: [email protected] (K. Kagawa). 1 Both authors contributed equally to this work. 1095-6433/$ - see front matter D 2004 Elsevier Inc. All rights reserved. doi:10.1016/j.cbpb.2004.03.001

extremely long life span, and necessarily live in protected environments. We are interested in such a long-lived DNA. To investigate it, we isolated DNA from chick heart and cerebrum at various stages of differentiation. We compared these DNAs after incubation with a low concentration of DNase I. We found that both DNAs became relatively resistant to DNase I during embryogenesis. We speculated that such a difference was induced by an as yet uncharacterized change in the DNA. To date, only one type of modification, DNA methylation, is known as an epigenetic modification that is capable of altering gene expression. Recent study revealed that methylation-mediated silencing occurs frequently in cancers (DePinho, 2000; Velicescu et al., 2002). The methylation of several genes increases with age in normal tissues (Yuasa, 2002). In this paper, we have identified a new type of DNA modification in heart and cerebrum. We show the change in the molecular weight of the deoxynucleoside in cardiac and cerebral DNA, and propose a model of the deoxynucleoside.

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2. Materials and methods 2.1. DNA isolation DNA in various organs was isolated from chicks (Gallus gallus) at E5 (embryonic day 5) through 1 day of age and 15 months of age. As an example, 30 chick hearts at 1 day of age were suspended in Tris/EDTA buffer (100 ml), which contained 30 mM Tris/HCl (pH 7.5) and 2 mM EDTA, and then homogenized three times with an Ultra-Turrax homogenizer (Janke & Kunkel, Staufen, Germany) at 10 000 rev./min for 1 min. The homogenate was incubated at 37 jC for 3 h in the presence of 0.25% sodium dodecylsulfate and 100 Ag/ml proteinase K. After incubation, DNA was extracted with an equal volume of 80% phenol and 0.15 M NaCl after being neutralized by the addition of a 1:20 volume of 1 M NaOH, and the aqueous phase was pooled after centrifugation. This extraction was repeated with an equal volume of chloroform. Two volumes of cold ethanol and 50 mM sodium acetate buffer (pH 5.1) were added to the pooled aqueous layer. Fibrous precipitates were collected, dissolved in water and dialyzed against 1 mM EDTA. Then, 10 Al of bovine pancreas RNase A solution (10 mg/ml, Wako, Nippon Gene, Japan) was added to 30 A260 units of DNA in Tris/EDTA buffer and incubated at 37 jC for 2 h. After incubation, DNA was extracted with chloroform/ether (2:1). Then, two volumes of cold ethanol were added to the pooled aqueous layer as above. Fibrous precipitates were dissolved in water, and extensively dialyzed against 1 mM EDTA. 2.2. DNA polymerase assay DNA (1 A260 unit/ml) was previously incubated with various concentrations of DNase I (1– 10  10 5 mg/ml, PL Biochemicals) in the presence of 40 mM Tris/HCl, pH 7.5 and 5 mM MgCl2 at 37 jC for 3 min and then the solution was heated at 76 jC for 5 min and immediately cooled in an ice bath (Aposhian and Kornberg, 1962). The standard assay (100 Al) included 0.016 A260 units of DNA in the presence of 66 mM K-phosphate buffer (pH 7.4), 0.33 mM DTT, 6 mM MgCl2, 0.1 mM each of the four dNTPs, and [3H]dTTP (0.2 ACi), and 1 unit of E. coli DNA polymerase I (Roche). Incubation was for 1 h at 37 jC, and acid precipitable [3H]-DNA products were collected on GF/C filters. Total radiolabelled nucleotide incorporation into DNA was measured by liquid scintillation counting (Richardson et al., 1964).

applied immediately to a TSKgel DNA-NPR column (0.46  7.5 cm, Tosoh, Tokyo, Japan). Elution was performed at 35 jC using a linear gradient from 0.5 M NaCl in 20 mM Tris/HCl, pH 7.6 to 1 M NaCl in Tris/HCl, pH 9.0 for 20 min at a flow rate of 0.5 ml/min. 2.4. HPLC analyses deoxynucleosides

of

deoxynucleotides

and

DNA (40 A260 units) was incubated with 20 A of nuclease P1 (400 units/ml, Seikagaku Kogyo Co., Japan), 10 Al 0.5 M Na –acetate buffer, pH 4.5 and 10 Al 0.1 M ZnSO4 in 4 ml of solution at 50 jC for 1 h. After shaking with chloroform, the deoxynucleotides were subjected to HPLC using a Cosmosil 5C18 column (10  250 mm, Nacalai Tesque, Kyoto, Japan) with solvent A [3% acetonitrile and 0.002% trifluroacetic acid (TFA), pH 3.6]. The separated deoxynucleotides were collected and dried in a Speed Vac at 4 jC. Each deoxynucleotide fraction was incubated with acid phosphatase (Type X, Sigma) in 20 mM sodium acetate buffer, pH 4.5, at 37 jC for 1 h. Thereafter, the deoxynucleosides were isolated by HPLC using the same column with solvent B (10% methanol and 0.002% TFA) at a flow rate of 0.7 ml/min. 2.5. HPLC analysis of sugar constituents To obtain sugar constituents, the deoxynucleosides were hydrolyzed in acid as described in the text. After removal of the acid in a rotary evaporator, the sugar constituents derived from the deoxynucleosides were applied to a Shodex 801 column (0.8  50 cm, sulfonated polystyrene gel, Showa Denko, Kawasaki-city, Japan), and monitored by refractometer, using water as mobile phase. 2.6. Chemical ionization mass spectrometry (CIMS) All MS experiments were performed with a Shimadzu GCMS/QP5050 system. CIMS was performed in the positive ion (chemical ionization) mode, using isobutane as the reagent gas. The interface was maintained at 230 jC. The sample was introduced into the mass spectrometer by direct insertion of the probe and mass spectra were obtained continuously within a mass range of 60 –400 m/z, while the probe was heated at a programmed rate of 40 jC/min to 280 jC. The samples (0.05 –0.3 A260 units) were introduced into a sample tube (4 Al), dried in a Speed Vac, and used for CIMS.

2.3. HPLC analysis of DNA 2.7. NMR spectroscopy of cerebral deoxynucleotides DNA (0.3 A260 units/ml) was incubated with various concentrations of DNase I in the presence of 20 mM Tris/ HCl, pH 7.5 and 1 mM MgCl2 at 37 jC for various periods and NaCl was added at 0.12 M. The sample was then

Cerebral DNA isolated from bovine cerebral cortex was digested with nuclease P1 and resulting deoxynucleotides were isolated by HPLC. Cerebral deoxynucleotides

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were subjected to NMR spectroscopy. NMR spectra (1H and 13C) were measured at RT in D2O with a JEOL JNM-GX 400FT NMR spectrometer; at 400 MHz for proton and at 100 MHz for carbon. Values of chemical shifts were recorded in parts per million by taking the signal of tetramethylsilane as the standard. 13C spectra were measured using a broad-band decoupling of proton signals. 2.8. Formation of the sugar-amide Cerebral dG (6 A260 units in 200 Al) was incubated with 10 Al of 1-naphthylamine (100 mg/ml ethanol) at 58 jC for 6 h. The modified 2-deoxyribose (200 Al) was incubated with 10 Al 1-naphthylamine, pH 7.0 at 40 jC for 17 h. After incubation the sample was dried in a rotary evaporator, dissolved in 5% methanol and subjected to HPLC as described in the text. 2.9. Oxidation of sugars with periodic acid Samples (100 Al) were incubated with 25 mM periodic acid (5 Al) under various conditions as mentioned in the text. To terminate the reaction, 2% sodium arsenite in 0.5 M HCl (13 Al) was added. The products were analyzed by colorimetry and HPLC as described in the text. 2.10. Chemical synthesis of 3-deoxyhexonic acid and 3deoxyglucosone 3-Deoxyhexonic acid and 3-deoxyglucosone was chemically synthesized by the alkaline method (Nef, 1910; Kennedy and White, 1983). Glucose (1 g) and potassium hydroxide (0.5 g) in 10 ml water were incubated at 37 jC overnight. Then the solution was neutralized to pH 7 approximately by adding HCl. It was then clarified with activated charcoal powder and concentrated in a rotary evaporator. The concentrated solution was charged to a Bio-Gel P-2 column (1  100 cm) and eluted with distilled water. Two peaks of deoxysugar-acid and deoxy-sugar were detected colorimetrically by periodate oxidation and reaction of the product with 2-thiobarbituric acid (Waravdekar and Saslaw, 1959). The reducing power of the deoxy-sugar was detected colorimetrically (Park and Johnson, 1949). The first peak was predominantly 3-deoxyhexonic acid and the second peak contained 3-deoxyglucosone, which were further purified on a TSK-gel Amide-80 column (0.78  30 cm, Tosoh, Japan) with 80% acetonitrile. Identification of the materials was performed by HPLC (Shodex 801) and paper chromatography on Whatman No.1 using two developing systems: acetone/butanol/water 7:2:1 and water saturated 2-butanone (Kucar et al., 1975). Another procedure was also adopted for the synthesis of 3-deoxyglucosone from glucose by using butylamine (Rowell and Green, 1970).

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3. Results 3.1. Comparison of various DNAs using E. coli DNA polymerase I To investigate the DNA of various organs, DNAs were isolated from chick at 1 day of age. These DNAs were treated with DNase I, and single-strand breaks were assessed by using an E. coli DNA polymerase I reaction system. The level of activity with these DNAs, which was measured as the incorporation of radioactivity of [3H]-TTP into the DNA, was compared. E. coli DNA polymerase I is well known to fill the gaps between fragments of the lagging strand. It preferentially uses DNA with gaps as a template-primer. As shown in Fig. 1, significant incorporation of radioactivity was observed in skeletal muscle DNA and cerebellum DNA at a low concentration of DNase I. Liver DNA was tested, and the same level of activity was obtained (data not shown), whereas, cardiac and cerebral DNA differed from other DNAs. At a low concentration of DNase I, below 2  10 5 mg/ml, little incorporation of radioactivity in cardiac or cerebral DNA was observed, whereas other DNAs were completely activated. At 2.5  10 5 mg/ml DNase I, both DNAs were activated, although at low levels. The decrease in the priming capacity of DNA at high concentration of DNase I, over 2.5  10 5 mg/ml, is due to the partial fragmentation of DNA. Double-strand breaks are not detected in this assay. To study the relation with embryonic development, DNA was isolated at E5. Cardiac DNA at E5 was activated similarly to the DNA from cerebellum and skeletal muscle. We speculate from these findings that cardiac DNA was sensitive to DNase I at E5, but gradually became resistant during embryogenesis.

Fig. 1. Effect of DNase I on priming activity of various DNAs in E. coli DNA polymerase I reaction. Cardiac DNA at E5 and 1 day of age, and DNA from other organs (cerebrum, cerebellum and skeletal muscle) at 1 day of age were preincubated with various concentrations of DNase I at 37 jC for 3 min. The template-primer activity was compared using an E. coli DNA polymerase I reaction system.

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3.2. Sensitivity of cardiac and cerebral DNA to DNase I DNA (whole body DNA at E5, cardiac DNA at 15 months of age and cerebral DNA at 1 day of age) was incubated with various concentrations of DNase I at 37 jC for 3 min and then the sample was applied to a TSKgel DNA-NPR column. Native DNA not treated with DNase I, was identified based on its elution profile, when it was eluted from the column with a linear gradient from 0.5 M NaCl in 20 mM Tris/HCl pH 7.6 to 1 M NaCl in Tris/HCl pH 9.0 (Fig. 2a, upper), whereas, DNA fragments after DNase I treatment flowed through the column (Fig. 2a, lower). At a low concentration of DNase I (2  10 5 mg/ml), all DNAs were adsorbed by the column (Fig. 2b), whereas, at a high concentration (over 5  10 5 mg/ml), E5 DNA was digested and flowed through the column (Fig. 2b). But the cardiac and cerebral DNAs were intact and adsorbed by the column (Fig. 2b). From the E. coli DNA polymerase reaction experiment, we presumed that DNase I introduced a single-strand break in E5 DNA at low concentration, but not in cardiac or cerebral DNA. These

results were consistent with those of Fig. 2b. When incubated with DNase I (7  10 5 mg/ml), for various periods, E5 DNA was broken down in 3 min (Fig. 2c), whereas, cardiac DNA was not digested even after incubation for 10 min (Fig. 2c). These findings suggested that cardiac and cerebral DNA seemed to be relatively resistant to DNase I. When incubated with DNase I (15  10 5 mg/ml) for 3 min, under various pH conditions, E5 DNA was digested at pH 7.5 to 8.0 (Fig. 2d). Under the same conditions, cardiac DNA at 15 months of age was completely resistant to DNase I between pH 7.0 and 9.0 (Fig. 2d), whereas, cardiac DNA at E17 was partially resistant (Fig. 2d). These findings suggested that cardiac DNA, seemed to be modified gradually during embryogenesis, and therefore, was partially resistant to DNase I at E17. 3.3. HPLC analyses of cardiac deoxynucleotides and deoxynucleosides Whole body DNA at E5 and cardiac DNA at 1 day of age were digested with nuclease P1, and the resultant

Fig. 2. Sensitivity of cardiac and cerebral DNA to DNase I. (a) HPLC analysis of DNA with (lower panel) and without (upper panel) DNase I treatment. (b) DNA (whole body DNA at E5, cardiac DNA at 15 months of age and cerebral DNA at 1 day of age) was incubated with various concentrations of DNase I at 37 jC for 3 min and analyzed by HPLC. The amount of DNA adsorbed by the column (%) was quantified from the elution profile. (c) DNA (whole body DNA at E5 and cardiac DNA at 15 months of age) was incubated with DNase I (7  10 5 mg/ml) for various periods, and analyzed by HPLC. (d) DNA (whole body DNA at E5, and cardiac DNA at E17 and 15 months of age) was incubated with DNase I (15  10 5 mg/ml) under various pH conditions and analyzed by HPLC.

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nucleotides were separated by reverse-phase HPLC (Fig. 3a,b). Each of the four deoxynucleotide species of E5 DNA, Cp, Ap, Gp and Tp, eluted similarly to canonical deoxyribonucleotides (dCMP, dAMP, dGMP and dTMP) in a single peak (Fig. 3a). The elution profile of cardiac deoxynucleotides at 1 day of age was different from that of deoxynucleotides at E5, and several new peaks appeared, Cp1, Ap1 and Gp1 (Fig. 3b). In this report, deoxynucleotides Ap, Cp, Tp and Gp were analyzed, but Ap1, Cp1 and Gp1 were not. Ap was incubated with acid phosphatase and the resultant deoxynucleoside was separated by reverse-phase HPLC. Cardiac deoxynucleoside A (dA) at E7 eluted similarly to canonical dA in a single peak (A1) (Fig. 3c), whereas, cardiac dA at E12 eluted in three fractions; A1, A2 and adenine (Fig. 3d). The amount of A2 increased during embryogenesis, and after hatch A2 was predominant (Fig. 3e). Furthermore, the cardiac dA at E12 through E19 was unstable, and adenine was released from the sugar during incubation with acid phosphatase (Fig. 3d). However, cardiac dA at 1 day of age was relatively stable, and little adenine was released (Fig. 3e). Cardiac dC at E7 eluted similarly to canonical dC in almost a single peak, C1 (Fig. 3f), whereas, at 1 day of age it was separated in two fractions, C1 and C2 (Fig. 3g). The amount of C2 increased during embryogenesis like that of A2.

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3.4. Mass spectrometric analyses of cardiac deoxynucleosides To verify the changes to deoxynucleosides, we further analyzed them by CIMS. The analyses were performed in the direct insertion and the chemical ionization (positive ion) modes. Fig. 4A shows the MS (mass spectrum) of A1 from whole body DNA at E5, which coincided with that of canonical dA. The ion at m/z 136 was attributed to adenine (MW, 135.13). The ion at m/z 117 was consistent with 2-deoxyribose. The molecular mass of 252 (MW, 251.24) coincided with the sum of adenine and 2-deoxyribose. Cardiac A2 at 1 day of age was analyzed as above. Cardiac A2 was much more difficult to evaporate than dA at E5. A2 evaporated at approximately 6 min as shown by the monitoring of total ion current (TIC), but dA at E5 evaporated at 3.5 min (arrow in Fig. 4B). SIM (selective ion monitoring) at m/z 252 and 282 (Fig. 4B) and the mass spectrum of A2 (Fig. 4C) indicated that the ion at m/z 252 was scarcely detectable, but a prominent molecular ion at m/z 282 was observed instead, which is greater than the dA at E5 (canonical dA) by m/z 30. A1 at 1 day of age was analyzed as above. On the monitoring of TIC, two peaks appeared to overlap (Fig. 4D). In the first peak, the predominant ion was at m/z 252 and a molecular ion at m/z 280 was observed, which was greater than the

Fig. 3. HPLC analyses of deoxynucleotides (a – b) and deoxynucleosides (c – g). (a) Deoxynucleotides from whole body DNA at E5. Arrows show the positions of canonical deoxyribonucleotides, identified by separate chromatographies. (b) Cardiac deoxynucleotides at 1 day of age. (c) Cardiac dA at E7. An arrow shows the position of canonical dA. (d) Cardiac dA at E12. (e) Cardiac dA at 1 day of age. (f) Cardiac dC at E7. An arrow shows the position of canonical dC. (g) Cardiac dC at 1 day of age.

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3.5. Sugar constituents in cardiac DNA

Fig. 4. CIMS analyses of deoxynucleosides A. (A) MS (mass spectrum) of A1 at E5. (B) TIC (total ion current) and SIM (selective ion monitoring) of cardiac A2 at 1 day of age. (C) MS at 5.9 min in (B). (D) TIC and SIM of cardiac A1 at 1 day of age. (E) MS at 6.1 min in (D).

canonical dA by m/z 28 (Fig. 4E). In the second peak, the ion at m/z 282 was observed (Fig. 4D). It is worth noting that the ion at m/z 252 was detected only in A1 and not in A2. A fragment ion at m/z 136 was also observed and was attributed to adenine (Fig. 4C,E). A fragment ion at m/z 117 was also observed, which was consistent with 2-deoxyribose. Therefore, it is likely that a structure corresponding to 2-deoxyribose is included in A1 and A2. The MS of canonical dC showed a molecular ion at m/z 228, which was attributed to the sum of cytosine (m/ z 111) and 2-deoxyribose (m/z 117) (Fig. 5A). In cardiac C2 at 1 day of age, the ion at m/z 228 was scarcely detectable and a prominent ion was observed at m/z 258 (Fig. 5B,C). This molecular ion is greater than the canonical dC by m/z 30, which paralleled the findings for A2 as described above. In cardiac C1 at 1 day of age, two molecular ions at m/z 256 and 258 were detected (Fig. 5D,E). The ions at m/z 256 and 258 are larger than the canonical dC by m/z 28 and m/z 30, respectively. Furthermore, the ion at m/z 228, which corresponded to a molecular ion of canonical dC (Fig. 5A), was observed (Fig. 5E). It was clear that the molecular ion of canonical dC was detected only in C1 and not in C2, which paralleled the findings for cardiac A1 and A2.

To further identify the sugar, cardiac deoxynucleosides were hydrolyzed in acid and the resultant hydrolysate was analyzed by CIMS. To avoid degradation of the sugar, the sample was hydrolyzed under conditions as mild as possible. As a control, canonical dA was incubated at 55 jC for 10 min in the presence of 10 mM HCl, and analyzed by CIMS after removal of the acid by a rotary evaporator. As shown in Fig. 6A,B, the molecular ion at m/z 252 was lost and only two ions at m/z 136 and 117 were detected, which were attributed to adenine and 2-deoxyribose, respectively. Cardiac A1 at 1 day of age was incubated at 55 jC for 10 min in the presence of 0.1% TFA, and analyzed as above (Fig. 6C,D). The molecular ion at neither m/z 280 nor 282 was detected, therefore, A1 was considered completely hydrolyzed. Fragment ions (Fig. 6D) except the ion at m/z 136 (adenine) were significantly different from those of canonical dA (Fig. 6B). The ion at m/z 117, which is attributed to 2-deoxyribose, was notably less prevalent than that of the acid-hydrolyzed canonical dA (Fig. 6A,B), but other ions at m/z 146 and 186 were detected (Fig. 6D). A2 was relatively heat resistant upon acid hydrolysis, since to remove adenine from A2 boiling was necessary under acidic conditions. Cardiac A2 at 1 day of age was boiled for 3 min in the presence of 5 mM HCl, and it was analyzed as above.

Fig. 5. CIMS Analyses of deoxynucleosides C. (A) MS of canonical dC. (B) TIC and SIM of cardiac C2 at 1 day of age. (C) MS at 5.4 min in (B). (D) TIC and SIM of cardiac C1 at 1 day of age. (E) MS at 5.7 min in (D).

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carbonyl additive 2-deoxyribose as shown in the inset in Fig. 7b, which was later identified as 3-deoxyglucosone. This proposed molecular structure will be verified by 13C NMR spectrometric analysis later. Under the different conditions of hydrolysis, we obtained different sugar constituents from cerebral deoxynucleotide Gp. Cerebral Gp was boiled for 2 min in the presence of 20 mM HCl, and after removal of the acid with a rotary evaporator, it was incubated with acid phosphatase in sodium acetate buffer, pH 4.5, for 1 h at 37 jC. The sample was analyzed on the same column (Fig. 7c). The flowing through peak was salt, but the material eluted at 11 min was the acidic sugar constituent. CIMS analysis showed that the acidic sugar exhibited three major ions at m/z 163, 145 and 127 (Fig. 7d). The structure of the ion at m/z 163 (or 162) likely has an acidic residue of COOH bound to the 1Vcarbon of 2-deoxyribose as shown in the inset of Fig. 7d, which was later identified as 3-deoxyhexonic acid. It is worth noting that the hydroxyl residue was bound to a carbonyl group

Fig. 6. CIMS analyses of hydrolysate of canonical dA and cardiac deoxynucleoside A1. (A) TIC and SIM of hydrolysates of canonical dA. (B) MS of (A). Inset is the chemical structure of hydrolyzed canonical dA. (C) TIC and SIM of hydrolysates of cardiac A1 at 1 day of age. (D) MS at 3.75 min in (C). Inset is the presumed chemical structure of hydrolyzed cardiac A1.

The MS of the hydrolysate of A2 was identical with that of A1 (Fig. 6D) (data not shown). The ion at m/z 146 was detected in cardiac A1 and A2 at 1 day of age (Fig. 4C,E), but not in A1 at E5 (canonical dA) (Fig. 4A). The ion at m/z 186, which was also observed in A1 and A2 (Fig. 4C,E), presumably arose from the degradation of adenine, and the proposed structure is indicated in the inset of Fig. 6D. This structure will be verified later. To isolate the sugar, the hydrolysate of A1 was applied to a Shodex 801 column after removal of the acid (Fig. 7a). Sugars were monitored based on a refractive index (RI) using a refractometer. The large peak, flowing through the column, is salt. Arrows show positions of authentic glucose, ribose and 2-deoxyribose (Fig. 7a). The predominant neutral sugar at the retention time of 20 min was analyzed by CIMS (Fig. 7b). An ion at m/z 145 was observed. An ion at m/z 146, which is a protonated form of that at m/z 145, was observed in the hydrolysate of A1 (Fig. 6D). The same ion at m/z 146 was included in the hydrolysate of A2, therefore, we expected the structure of the sugars in A2 to be the same as that seen in A1. Furthermore, a part of A2 was shifted to A1, when A2, that had been stored in the refrigerator for 1 week, was analyzed again by HPLC (data not shown). We presumed that the ion at m/z 145 corresponds to the

Fig. 7. Analyses of sugars. (a) HPLC analysis of acid-hydrolyzed sugars from cardiac A1 at 1 day of age on a Shodex 801 column. Arrows show the positions of glucose, ribose and 2-deoxyribose identified by separate chromatographies. (b) MS of the neutral sugar at 20 min. Inset is a model of the sugar at m/z 145. (c) HPLC analysis of sugars released from cerebral Gp after boiling in acid and incubation with acid phosphatase. (d) MS of the acidic sugars at 11 min. Inset is a model of the sugar at m/z 163.

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instead of a proton during hydrolysis. The ratio of acidic to neutral sugar was variable. 3.6. NMR spectra of cerebral Ap and Cp in comparison with those of canonical dAMP and dCMP To verify the proposed molecular structure, we further analyzed Ap isolated from DNA of bovine cerebral cortex by 13 C NMR and compared the results with those for canonical dAMP. All signals corresponding to the sugar carbons of canonical dAMP (CV-1 to CV-5; Fig. 8a) were quantitatively observed at the same relative intensities in the cerebral Ap (Fig. 8b). This indicates that the same structure of 2-deoxyribose is present in the cerebral Ap. Furthermore, all signals corresponding to adenine carbons of canonical dAMP (C-2, C-4, C-5, C-6, C-8) were quantitatively observed at the same relative intensities in the cerebral Ap (Fig. 8b). The above observations indicate that the base was not changed, and that the structure of the sugar was similar to that of 2-deoxyribose. However, an extra signal at 173 ppm was clearly observed in the cerebral Ap (Fig. 8b). 13C NMR was performed for canonical dCMP and cerebral Cp (Fig. 8c,d, respectively). All the signals corresponding to cytosine and 2-deoxyribose carbons were also observed in cerebral Cp. However, an extra signal at 174 ppm corresponding to a carbonyl carbon was observed, similar to the cerebral Ap, near position 165 ppm of

C-2 (CMO) (Fig. 8d). The chemical shift value for carbonyl carbon atoms can be used to identify the type of functional group present in the molecule: 160 –180 ppm for amides, 160– 190 ppm for esters, 165 –185 ppm for carboxylic acids, 190– 205 ppm for aldehydes and 200 –220 ppm for ketones. This new signal seems to be attributable to the amide carbonyl between the base and 2-deoxyribose (insertion in Fig. 8b,d). From these findings, we assumed that the constituent sugar in the cardiac and cerebral DNA is 3-deoxyglucosone (3-deoxyhexos-2-ulose) not a 2-deoxyribose. 1 H NMR spectrum was measured for the cerebral Ap, and compared with that of canonical dAMP (Fig. 8e: dAMP; Fig. 8f: cerebral Ap). The cerebral Ap was almost identical to the canonical dAMP. Numbers of the corresponding protons (1V – 5V) were the same and the values of the chemical shifts differed little. Similarity of the spectrum of the cerebral Ap to that of the authentic dAMP implies that all the protons of the cerebral Ap are in almost identical chemical environments as those of the authentic dAMP, which would support our model. 3.7. Detection of carbonyl residues in the modified deoxynucleoside and the sugar If the linkage between the base and the sugar moiety is actually an amide linkage (carbonyl carbon exists between

Fig. 8. NMR spectra of cerebral Ap and Cp. 13C NMR (a – d), 1H NMR (e – f). (a) Canonical dAMP. (b) Cerebral Ap. (c) Canonical dCMP. (d) Cerebral Cp. (e) Canonical dAMP. (f) Cerebral Ap.

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an N of the base and carbon of 2-deoxyribose as shown in Fig. 8b,d), it may be highly probable that the purine base of the deoxynucleotide or deoxynucleoside is nucleophilically substituted with other amines. After the reaction, the base would be released from the deoxynucleoside and the sugaramide would be detected (Fig. 9A). To verify this hypothesis, the cerebral deoxynucleoside was incubated with fluorescent 1-naphthylamine (NH2R in Fig. 9A). If the carbonyl residue is present, 1-naphthylamine would react with it to form a fluorescent sugar-amide, which would be detected by the fluorospectrophotometer. In Fig. 9a, the fluorescent naphthyl residue is shown as red R. Furthermore, the reaction would give a free base, which would be detected by UV (Fig. 9A). To test this possibility, cerebral dG was incubated with 1-naphthylamine at 58 jC for 6 h, and then the sample was applied to a Wakosil 5 C18 column (0.75  25 cm, Wakenyaku Co., Japan) (Fig. 9B). Elution was carried out with 5% methanol at RT. When the sugaramide is formed during the reaction, it is detected by the fluorescence of naphthylamine, which can be monitored at 450 nm with excitation at 350 nm. We observed the fluorescent peak in fraction 33 –40 (Fig. 9B). Under the conditions, free naphthylamine was adsorbed by the column, so the fluorescent peak was not from free naphthylamine. The fluorescent peak was presumed to be the sugaramide, which was confirmed by the next experiments (Fig. 9C,D). The second peak in fractions 65– 80 was identified as the free guanine judging from the UV spectrum. The third

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peak (fraction 130 –160) was the non-reacted cerebral dG. Treatment of canonical dG with naphthylamine caused no such production of fluorescent materials nor any release of guanine whatsoever (data not shown). To exclude the possibility that the carbonyl residue is included in the base, we isolated the acidic sugar from cerebral dG, and incubated it with 1-naphthylamine. If the carbonyl residue is present in the acidic sugar, the fluorescent peak (sugar-amide) should be detected by the same HPLC. The fluorescent peak eluted at nearly the same retention time (Fig. 9C). To confirm that the fluorescent peak is a sugar-amide, the fluorescent fractions in Fig. 9C were collected and hydrolyzed by boiling in 0.1 N HCl for 1 min, and analyzed again on the same column (Fig. 9D). The fluorescence was abolished completely and the acidic sugar appeared instead at fraction 16, which was detected by colorimetry (Waravdekar and Saslaw, 1959). These results showed that the fluorescent peak was the sugar-amide and 1-naphthylamine was reversibly detached from the sugar-amide under acidic conditions (Fig. 9A). These findings suggested that the carbonyl residue exists between the base and the 2-deoxyribose as shown in the inset of Fig. 8b,d. 3.8. Determination of the sugar structure by periodic acid oxidation As mentioned above, the carbonyl residue likely exists in the sugar constituent according to the 13C NMR spectra and

Fig. 9. Reaction of fluorescent 1-naphthylamine (NH2R) and the carbonyl residue. (A) The presumed reaction of fluorescent 1-naphthylamine (NH2R) and the carbonyl residue of the modified deoxynucleoside. Fluorescent naphthyl residue is shown as red R. (B) HPLC analysis of cerebral dG after incubation with 1naphthylamine. (C) HPLC analysis of the acidic sugar after incubation with 1-naphthylamine. (D) The fluorescent peak in (C) was hydrolyzed in 0.1 M HCl at 100 jC for 1 min, and analyzed again as above. Acidic sugar was detected by colorimetry at A532 nm.

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product g was identified as malonaldehyde (Fig. 10c). The product a had a shorter retention time than malonaldehyde, and h was a mixture of both materials. From these observations we speculated that the product a was the X-linked malonaldehyde (formula in Fig. 10a, a). On using a larger amount of periodic acid, cleavage between CV-1 and X occurred to produce malonaldehyde. These results led us to conclude that the carbonyl residue is most likely located next to CV-1. Therefore, we proposed a model of the sugar constituent (m/z 145), which is indicated in the inset of Fig. 7b. The structure of the ion at m/z 163 likely has an acidic residue of COOH bound to the CV-1 of 2-deoxyribose as shown in the inset of Fig. 7d.

4. Discussion 4.1. Structure of the modified deoxynucleoside

Fig. 10. Periodic acid oxidation. (a) Pathway of formation of malonaldehyde and formaldehyde from 2-deoxyribose and the modified 2deoxyribose after periodic acid oxidation. (b) Oxidation of 2-deoxyribose (open circle), and the modified 2-deoxyribose (closed circle) with periodic acid at the indicated concentration for 20 min. Product was detected by colorimetry at A532 nm. (c) After oxidation at 63 jC in (b), the products (a, h, g) were analyzed on a TSK-gel Amide-80 column.

In this report, we showed a new type of DNA modification in chick heart and cerebrum. We proposed a model of the deoxynucleoside in which a carbonyl residue exists between the base and the 2-deoxyribose moiety of the canonical deoxyribonucleoside (Fig. 11a). The carbonyl residue has the chemical properties of an active amide, which was supported by 13C NMR. A significant feature of this model is that the linkage between the base and sugar moiety is an amide linkage rather than an N-glycosyl one in

analysis of the reaction with 1-naphthylamine. The binding site of the carbonyl residue on 2-deoxyribose needs to be clarified. We presumed that the binding site is CV-1 or CV-5, because we observed a fragment ion attributed to 2-deoxyribose at m/z 117 (Fig. 4c,e and Fig. 5c). We adopted the method of periodate oxidation, which is commonly used for the identification of sugars. Periodic acid reacts with vicinal hydroxyl groups to cleave the linkage between them as shown in Fig. 10a. Oxidation of 2-deoxyribose led easily to the formation of malonaldehyde and formaldehyde (Fig. 10a, upper). 2-Deoxyribose was oxidized at 20 jC forming malonaldehyde (Fig. 10b). If the carbonyl residue (X) links to CV-1 in the modified 2-deoxyribose, the product of the oxidation reaction should not be malonaldehyde but another form as shown in Fig. 10a, lower panel. The modified 2deoxyribose (inset of Fig. 7b,d) was oxidized with various amounts of periodic acid (Fig. 10b). The cleaved products were analyzed by colorimetry with 2-thiobarbituric acid (Waravdekar and Saslaw, 1959). When the concentration of periodic acid was lower than 0.3 mM, the product was not malonaldehyde, which displays a faint color (Fig. 10b, a). But if the concentration exceeded 0.4 mM, the product displayed a deep color, indicating it was malonaldehyde (Fig. 10b, g). The oxidation product was separated on a TSK-gel Amide-80 column (0.78  30 cm, Tosoh, Japan) using 80% acetonitrile. Judging from the retention time, the

Fig. 11. Model of the modified deoxynucleoside and sugar constituents. (a) Structure of the modified deoxynucleoside and sugar constituents after hydrolysis. (b) Alkaline degradation of glucose (Kennedy and White, 1983; Knill and Kennedy, 2003).

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the canonical deoxyribonucleosides. Two species of sugar constituents were obtained from the modified deoxynucleoside under different conditions of hydrolysis. The neutral and acidic constituents seem to be 3-deoxyglucosone and 3deoxyhexonic acid, respectively (Fig. 11a). CIMS revealed the m/z of the acidic constituent to be 163 and that of the neutral one to be 145. 4.2. Chemical synthesis of 3-deoxyhexonic acid and 3deoxyglucosone To verify our model, we attempted to synthesize 3deoxyglucosone and 3-deoxyhexonic acid. It is widely recognized that glucose is converted to aldonic acid (3deoxyhexonic acid) via 3-deoxyglucosone (3-deoxyhexos2-ulose) on treatment in an alkaline solution (Fig. 11b) (Nef, 1910; Kennedy and White, 1983; Knill and Kennedy, 2003). Under alkaline conditions, rearrangement of the protons of glucose took place, and glucose was finally converted to 3deoxyhexonic acid having COOH at C-1. We synthesized 3deoxyhexonic acid and 3-deoxyglucosone from glucose by alkaline treatment as described in Section 2. The synthesized 3-deoxyhexonic acid was analyzed by TSK-gel Amide-80 column chromatography (0.78  30 cm, Tosoh, Japan) with 80% acetonitrile (Fig. 12a). It was predominantly eluted in fraction 56. It is known that an aldohexonic acid in aqueous solution is in equilibrium with lactones. The synthesized 3-deoxyhexonic acid gave two stereoisomers, ribo- and arabino-forms. Identification of the materials was

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performed using paper chromatography (Kucar et al., 1975). Fraction 56 (Fig. 12a) was boiled in 0.1 N formic acid for 1 min, and then the sample was analyzed as above (Fig. 12b). With the acid treatment, lactones of 3-deoxy-ribo-hexonic and 3-deoxy-arabino-hexonic acid were formed. The acidic material from the cerebral deoxynucleotide Gp (Fig. 7c) was applied to the same column. The elution profile was nearly identical, however, the acidic material gave only ribolactone. To confirm this finding, fraction 56 from the acidic material was treated with acid as above, and then analyzed again using the same HPLC (Fig. 12c). It was clear that the acidic material gave one kind of lactone, which was consistent with 3-deoxy-ribo-hexonic acid. CIMS analysis revealed that the synthesized 3-deoxyhexonic acid exhibited a molecular ion at m/z 163 (Fig. 12d), which was consistent with the acidic material (Fig. 7d). The formation of 3deoxyhexonolactone with a molecular weight of 163 (MH+) was reported (Heusinger, 1991). The five-membered ring structure was also reported in 3-deoxyhexonic acid (Yaylayan and Kaminsky, 1998; Pedersen, 1999). When it was treated in 0.02 N HCl at 55 jC for 10 min, the ring was opened, and it exhibited an ion at m/z 180 (Fig. 12e). The acidic material exhibited the ion at m/z 180 on the above same treatment (data not shown). Therefore, we concluded that the acidic sugar constituent from the modified deoxynucleotide was 2,5-anhydro-3-deoxy-D-ribo-hexonic acid (Fig. 11a). 3-Deoxyglucosone was synthesized by two procedures: the alkaline method and butylamine method (Rowell and

Fig. 12. Properties of synthesized 3-deoxyhexonic acid and similarity of the acidic material from modified deoxynucleotides. (a) HPLC (TSK gel) analysis of synthesized 3-deoxyhexonic acid. (b) HPLC analysis of fraction 56 in (a) after boiling in 0.1 N formic acid for 1 min. (c) HPLC analysis of the acidic material from cerebral deoxynucleotide Gp after the same treatment as in (b). (d) CIMS analysis of synthesized 3-deoxyhexonic acid. (e) CIMS analysis of synthesized 3-deoxyhexonic acid after incubation in 0.02 N HCl at 55 jC for 10 min.

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Green, 1970). The material was identified based on results of HPLC (Shodex 801) and paper chromatography (Rowell and Green, 1970). The synthesized 3-deoxyglucosone eluted in fraction 44 on Shodex 801 chromatography (Fig. 13a). It was detected by two methods: colorimetry for deoxysugar (at A532) with 2-thiobarbituric acid after periodate oxidation (Waravdekar and Saslaw, 1959) and colorimetry for the reducing power (at A690) (Park and Johnson, 1949). The elution profile was identical to that of the neutral sugar from the modified deoxynucleotide. It is well established that 3-deoxyhexonic acid is formed from 3-deoxyglucosone in the presence of mild alkaline concentrations of Ca(OH)2 or Ba(OH)2 (Rowell and Green, 1970; Kucar et al., 1975). 3-Deoxyglucosone (fraction 44) (Fig. 13a) was incubated in 14 mM Ca(OH)2 at 50 jC for 2 h. The pH was adjusted to slightly acidic and then the sample was analyzed as above. As shown in Fig. 13b, the reducing power decreased, and 3-deoxyglucosone was converted to 3-deoxyhexonic acid. The neutral sugar from cardiac deoxynucleoside (Fig. 7a) was treated with Ca(OH)2 under the same conditions as above for various periods. After incubation, the reducing power decreased (Fig. 13c, closed circles), and the acidic material, which was identified as 3-deoxyhexonic acid, increased (squares). As a control, 2-deoxyribose (open circles) did not change under the conditions. These results led us to conclude that the synthesized 3-deoxyhexonic acid and 3-deoxyglucosone coincided with the acidic material and the neutral sugar from the modified deoxynucleoside. All these results also support our model of the modified deoxynucleoside (Fig. 11a).

4.3. Metabolism of 3-deoxyglucosone 3-Deoxyglucosone and 3-deoxyhexonic acid were not known to be part of the metabolic pathway until recently. However, 3-deoxyglucosone has been found in the serum and erythrocytes of individuals with diabetes and uremia. It is a highly reactive alpha-dicarbonyl sugar and potent protein cross-linker that is important in the formation of advanced glycation end products by the Maillard reaction (non-enzymatic glycation between reducing sugar and protein amino groups), which has been postulated to lead to the development of diabetic complications (Fig. 14) (Yamada et al., 1994; Lal et al., 1997a; Niwa, 1999; Odani et al., 1999; Szwergold et al., 2002; Tsukushi et al., 2002). Quantitation of 3-deoxyglucosone levels in human and rat plasma was reported: a reactive dicarbonyl 3-deoxyglucosone was conjugated with 2,3-diaminonaphthalene to produce a stable adduct which was analyzed by GC/MS (Yamada et al., 1994; Lal et al., 1997a; Odani et al., 1999). The reactivity of the carbonyl of 3-deoxyglucosone was observed in our experiments: the reaction of the modified deoxynucleoside and fluorescent 1-naphthylamine (Fig. 9). Furthermore, 3deoxyglucosone is produced from glucose via sorbitol, fructose and fructose-3-phosphate (polyol pathway) in the lenses and hearts of diabetic rats (Szwergold et al., 1990; Lal et al., 1995, 1997b). 4.4. Biological features of the modified DNA There are marked differences in neurogenesis between the cerebrum and the cerebellum. All neurons in the cerebral cortex are prenatally generated (Rakic, 1974; Luskin and

Fig. 13. Properties of synthesized 3-deoxyglucosone and similarity of the neutral sugar from cardiac deoxynucleosides. (a) HPLC (Shodex 801) analysis of synthesized 3-deoxyglucosone. Deoxysugar was detected by colorimetry at A532 nm (a solid line), and the reducing power was detected by colorimetry at A690 nm (a dotted line). (b) HPLC analysis of synthesized 3-deoxyglucosone after incubation in 14 mM Ca(OH)2 at 50 jC for 2 h. (c) The neutral sugar from cardiac deoxynucleoside and 2-deoxyribose (as a control) was incubated under the same conditions as in (b) for various periods. 3-Deoxyhexonic acid was detected by colorimetry at A532 nm (squares), and 3-deoxyglucosone was detected by colorimetry at A690 nm (closed circles).

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Fig. 14. Metabolism of 3-deoxyglucosone in diabetes and uremia. 3Deoxyglucosone is synthesized via the Maillard reaction, and rapidly reacts with protein amino groups to form advanced glycation end products such as imidazolone (Niwa, 1999).

Shatz, 1985). However, a large proportion of the microneurons in the cerebellum, hippocampus, and olfactory bulb continue to divide during postnatal development (Altman, 1969; Finlay and Darlington, 1995). There is a close parallelism between cerebral neurogenesis and cardiac myogenesis. All cardiomyocytes are prenatally generated, whereas, skeletal muscle cells continue to proliferate during postnatal development and form syncytiums, and some myoblasts persist as satellite cells (Manasek, 1968; Alberts et al., 1983; Olson and Srivastava, 1996). Furthermore, cortial neurons and cardiomyocytes irreversibly lose the ability to proliferate and enter a terminal state of differentiation at almost the same embryonic stage (Finlay and Darlington, 1995). It is interesting that our finding of modified DNA in cardiac and cerebral cells, but not in skeletal muscle or cerebellum, is consistent with the early terminal differentiation of cardiac and cerebral cells. Furthermore, the finding of the sensitivity of cardiac DNA to DNase I (Fig. 2d), supported that the DNA modification occurs during embryogenesis. HPLC analyses of cardiac deoxynucleoside (Fig. 3c – e) also supported that the DNA modification occurs around E12. It is expected that the sugar molecules are modified or converted from 2-deoxyribose to 3-deoxyglucosone during embryogenesis, since the constituent sugar is 2-deoxyribose for DNA and 3deoxyglucosone for cardiac and cerebral DNA (modified DNA). The mechanism behind the modification of DNA needs to be clarified. We have recently isolated enzymes for the modification of DNA from the chick cerebrum. Studies are in progress. Cardiac cells and neurons in the cerebrum are called permanent cells (Alberts et al., 1983). We surmise hypothetically that the DNA modification gives one of the material bases of the nature of these cells, not to divide. The meaning of the presence of the permanent cells, however, is still obscure as discussed in the literature (Alberts et al., 1983). It is noteworthy that the transforma-

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tion of these permanent cells is extremely rare, which might be related to the fact that they are non-dividing cells. For example, intracerebral neuroblastoma has not been reported (Kenny et al., 1995). Even when the chemical carcinogen, nitrosoamides, is applied into the brain, no neuronal tumors were induced (Druckrey et al., 1972). Chemically induced tumors in the brain were found to originate almost exclusively from glial cells, and predominantly located in the subependymal areas of the lateral ventricles. These are the only regions where a significant proliferation of glial cells occurs in the adult mammalian brain (Kleihues et al., 1976). Although several cases of cardiac rhabdomyoma were reported in fetuses and the cardiac and muscular nature of the sarcomas was confirmed by immunohistochemistry, the primary cardiac sarcoma is also uncommon (Bordarier et al., 1994; Medioni et al., 1994).

Acknowledgements We thank Dr Hiroaki Tokimatsu of the Department of Physics at Osaka Medical College for the analysis by NMR spectroscopy.

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