Loss of DNAprotein crosslinks from formaldehyde-exposed cells occurs through spontaneous hydrolysis and an active repair process linked to proteosome function
George Quievryn and
Anatoly Zhitkovich1
Department of Pathology and Laboratory Medicine, Brown University, Box G-B511, Providence, RI 02912, USA
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Abstract
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DNAprotein crosslinks (DPC) involving all major histones are the dominant form of DNA damage in formaldehyde-exposed cells. In order to understand the repair mechanisms for these lesions we conducted detailed analysis of the stability of formaldehyde-induced DPC in vitro and in human cells. DNAhistone linkages were found to be hydrolytically unstable, with t
= 18.3 h at 37°C. When histones were allowed to remain bound to DNA after crosslink breakage, the half-life of DPC increased to 26.3 h. This suggests that ~30% of spontaneously broken DPC could be re-established under physiological conditions. The half-lives of DPC in three human cell lines (HF/SV fibroblasts, kidney Ad293 and lung A549 cells) were similar and averaged 12.5 h (range 11.613.0 h). After adjustment for spontaneous loss, an active repair process was calculated to eliminate DPC from these cells with an average t
= 23.3 h. Removal of DPC from peripheral human lymphocytes was slower (t
= 18.1 h), due to inefficient active repair (t
= 66.6 h). This indicates that the major portion of DPC is lost from lymphocytes through spontaneous hydrolysis rather than being actively repaired. Depletion of intracellular glutathione from A549 cells had no significant effect on the initial levels of DPC, the rate of their repair or cell survival. Nucleotide excision repair does not appear to be involved in the removal of DPC, since the kinetics of DPC elimination in XP-A and XP-F fibroblasts were very similar to normal cells. Incubation of normal or XP-A cells with lactacystin, a specific inhibitor of proteosomes, caused inhibition of DPC repair, suggesting that the active removal of DPC in cells may involve proteolytic degradation of crosslinked proteins. XP-F cells showed somewhat higher sensitivity to formaldehyde, possibly signaling participation of XPF protein in the removal of residual peptideDNA adducts.
Abbreviations: BSA, bovine serum albumin; BSO, L-buthionine-R,S-sulfoximine; DPC, DNAprotein crosslinks; DPTA, diethylenetriaminepentaacetic acid; FA, formaldehyde; GSH, glutathione; mBBr, monobromobimane; PBS, phosphate-buffered saline; PMSF, phenylmethylsulfonyl fluoride; XP, Xeroderma pigmentosum.
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Introduction
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Formaldehyde (FA) is a genotoxic chemical found in thousands of household, medicinal and industrial products (1,2). Large amounts of FA-containing resins are used in the production of plywood, particle board, textiles, paints, lubricants and dyes. FA is also extensively used in the furniture industry, upholstery, carpeting, drapery and garments. Offgassing of paints, carpets, fabrics, wall panels and many wood products creates FA-containing fumes, leading to widespread human exposure outside industrial settings (3). Although the major source of human exposure is through direct inhalation, FA can also be metabolically produced in the liver from many ingested or inhaled compounds.
Genotoxic activity of FA has been found in several test systems, including formation of nasal tumors in experimental animals (4,5). It is well established that FA is mutagenic to bacteria and lower eukaryotes (1) and clastogenic in mammalian cells (6,7). FA was also found to be mutagenic in human lymphoblasts at the thymidine kinase (8,9) and HPRT loci (10). Point mutations and deletions of various sizes were the predominant types of mutations induced by FA exposure (1012).
Exposure of cells to FA leads to the formation of DNAprotein crosslinks (DPC) as the major form of DNA damage (13). Reaction of FA with purified DNA can also yield N-hydroxymethyl adducts with dG, dA and dC. However, their presence was not detected in exposed cells (14,15). Histones are commonly found to be crosslinked to DNA by FA exposure (16). All five histones are capable of crosslinking to DNA, with a somewhat higher activity detected for histones H1 and H3. The formation of DNAhistone crosslinks proceeds through an initial rapid reaction of FA with histones followed by conjugation with amino groups of DNA (17). Epsilon amino groups of histone lysines and exocyclic amino groups of adenosine/guanosine bases are involved in crosslink formation. The general structure of FA-induced DPC can be illustrated as follows: histoneNH-CH2-NHDNA. Formation of DNAhistone crosslinks was shown to be strongly correlated with tumor incidence at the target sites, and these lesions are now considered to be largely responsible for the carcinogenicity of FA (13,18). Levels of FA-induced DPC are considered to represent a good molecular dosimeter of FA exposure and are frequently used for risk modeling and prediction of FA carcinogenicity for different species (1820). Despite the importance of DPC in understanding the genotoxic activity of FA and many other crosslinking agents, little is known about the repair mechanisms of these lesions. Previous studies reported a relatively rapid loss of FA-induced DPC from cultured cells (7,21). However, it was not clear whether it was a result of a repair process or whether DPC were lost due to hydrolytic instability. Aldehyde-induced DPC are known to undergo spontaneous hydrolysis, particularly at elevated temperatures (22,23).
In this work we have determined the relative contribution of hydrolytic instability to the overall kinetics of DPC removal from exposed cells. We have found that both spontaneous loss and an active repair process contribute to the rate of disappearance of DPC. The activity of the DPC repair component was unchanged in nucleotide excision repair-deficient Xeroderma pigmentosum (XP) complementation group A and XP-F fibroblasts, but could be blocked by inhibition of the proteolytic function of 26S proteosomes. The results suggest that DPC removal involves a novel repair pathway which appears to act through the proteolytic degradation of crosslinked proteins.
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Materials and methods
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Materials
Histone H1, calf thymus DNA (type I), SDS, phenylmethylsulfonyl fluoride (PMSF), Tris, EDTA, agarose, HPLC grade methanol and monobromobimane (mBBr) were obtained from Sigma (St Louis, MO). Calf thymus DNA was additionally purified by treatment with RNase A (100 µg/ml, 37°C, 60 min) and proteinase K (0.2 mg/ml, 50°C, 16 h) and phenol/chloroform extraction. A methanol-stabilized solution of formaldehyde was purchased from Mallinckrodt. Working solutions of formaldehyde were prepared immediately before use. PicoGreen was from Molecular Probes, lactacystin from CalBioChem and [3H]thymidine from Amersham.
Cells
Human Ad293 kidney and A549 lung cell lines were obtained from the American Tissue Culture Collection. HF/SV cells are human diploid fibroblasts transformed with SV40 virus (Dr H.Ozer, Rutgers University). Nucleotide excision repair-deficient XP-A (GM04312C) and XP-F (GM08207B) fibroblasts were obtained from the Coriell Cell Depository (Camden, NJ). A549 cells were grown in F-12 medium supplemented with 10% serum, HF/SV cells in DMEM plus 10% serum, Ad293 cells in DMEM plus 10% serum and XP-A fibroblasts in DMEM supplemented with 10% serum. Blood from three healthy male volunteers was collected by venipuncture into heparin-containing tubes. Mononuclear cells were isolated from freshly obtained blood by gradient separation using a Ficol/sodium diatrizote reagent (Histopaque-1077; Sigma). Cells were washed twice in phosphate-buffered saline, counted and then seeded at 0.5x106 cells/ml in multiwell plates containing RPMI 1640 medium, 10% serum. After an 1824 h adaptation period, the lymphocytes were used in experiments.
In vitro crosslinking
For all in vitro crosslinking reactions, 25 mM Na phosphate buffer, pH 7.0, 5 µg calf thymus DNA and 1 µg histone H1 or BSA were combined in a 50 µl volume and incubated at 37°C for different periods of time. Reactions were stopped by addition of 500 µl of 1% SDS, 20 mM Tris, pH 7.5, 2 mM EDTA and 1 mM PMSF.
Treatments of cells
Cultured cells (A549, Ad293, HF/SV, XP-F and XP-A) were radiolabeled with 1.0 µCi/ml [3H]thymidine for 48 h, the radioactive medium was removed and the cells incubated in regular medium for 3 h prior to formaldehyde exposure. Freshly prepared formaldehyde solutions were used for all experiments. Experiments for calculation of the half-lives of DPC removal involved 3 h exposure in complete medium. At the end of the exposure period cells were washed with phosphate-buffered saline (PBS), followed by addition of fresh medium. Cells were collected with trypsin/EDTA, washed in PBS and lysed in 1% SDS, 25 mM Na phosphate buffer, pH 7.0, 1 mM PMSF and 1 mM EDTA. Lysed samples could be stored at 70°C for several months without any detectable effect on the yield of crosslinks. Cytotoxicity of FA treatments was determined by colony formation assay. Cells were collected by trypsinization, counted and seeded on 60 mm dishes (2001000 cells/dish). Approximately 2 weeks later colonies were fixed with methanol and stained with Giemsa solution for counting.
DNAprotein crosslinking assay
The procedure for measurement of DPC was based on the earlier published protocol (24) with some modifications. In this method DNA fragments containing covalently attached proteins were selectively precipitated in the presence of KCl/SDS. Cells or DNA/histone mixtures were lysed in 1% SDS, cellular lysates were sheared by passing through 25G needles and DPC were precipitated by addition of 200 mM KCl. All heating steps were performed at 50°C for 10 (initial) or 5 min (washes). Dependence of DNA precipitation on the presence of attached proteins was verified by elimination of KCl/SDS-precipitable DNA by pre-treatment of samples with 0.2 mg/ml proteinase K for 1 h at 37°C prior to crosslink analysis. DNA was measured in the presence of 0.5 µM PicoGreen in a multiwell fluorescence reader (SPECTRAFluorPlus; Tecan). Fluorescent measurements were recorded 15 min after addition of the dye (emission 535 nm, excitation 485 nm). All manipulations with PicoGreen-containing samples were done under reduced illumination. In cultured cells DPC were determined by measuring [3H]thymidine radioactivity of the pellet and supernatant fractions. The number of crosslinks per 108 bp was calculated from the fraction of DNA fragments containing crosslinks and from the size of DNA fragments. It was important to determine the weighted average length of DNA fragments, which was calculated from the computer scanned images of agarose gels. Crosslinks/108 bp = CFx108/L, where CF = ln (1 crosslinked fraction) and represents the number of crosslinks per DNA fragment. The crosslinked fraction represents the ratio of SDS-precipitable DNA to total DNA. L is the weighted average length of DNA fragments produced during shearing in the KCl/SDS assay. L is measured in DNA base pairs. To determine the DNA fragment size, aliquots of sheared DNA lysates were digested with 0.2 mg/ml proteinase K for 1 h at 50°C and then analyzed on 0.51% agarose gels. Digitized profiles of ethidium bromide stained gels acquired on a GelDoc 2000 photodocumentation system (Bio-Rad) were transferred to Microsoft Excel and the weighted average length was calculated. A typical DNA fragment size for monolayer cell lines was in the range 2023 kb. Lymphocyte and lactacystin-treated samples frequently contained DNA fragments <20 kb.
Calculations of t
for DNAprotein crosslinks
Final DPC values were adjusted for background levels of crosslinks. The following equation was used: t
= ln(2)/k, where k (rate of removal) was calculated from exponential fits of background-corrected DPC values determined at different time points after FA removal. The R2 values were never less than 0.95 and typically were better than 0.98. The t
value for an active repair component was calculated as follows: t
= ln(2)/kactive, where kactive = koverall kpassive. Details of determination of the rate of spontaneous loss of DPC (kpassive) are given in Results. Due to high cytotoxicity at later time points, a single point was used to calculate the half-life in XP-A fibroblasts treated with lactacystin. The following formula was used: t
= txln(2)/ln(y), where t is time (h) and y is the DPC fraction remaining at that time point.
Glutathione measurements
Intracellular concentrations of glutathione (GSH) were determined by HPLC analysis of cell lysates derivatized with mBBr. The procedure was adopted from Newton and Fahey (25). Approximately 2x106 cells were collected, washed twice in PBS and suspended in 50 µl of PBS containing 5 mM diethylenetriaminepentaacetic acid (DPTA). The samples were acidified by addition of 200 µl of 50 mM methanesulfonic acid and then twice frozen (70°C) and thawed (37°C). GSH-containing supernatants were obtained after centrifugation at 12 000 g for 10 min at room temperature. The derivatization samples (total volume 250 µl) contained 50 µl cell extracts, 50 mM HEPES, pH 8.0, 5 mM DPTA, 15 mM NaOH and 2 mM mBBr. The reaction was allowed to proceed for 10 min in the dark at room temperature and was then terminated by addition of 25 mM methanesulfonic acid. HPLC measurements used a Shimadzu LC-10ADvp liquid chromatograph, equipped with a SIL-10ADvp autoinjector, an Ultrasphere ODS column (5 mm, 250x4.6 mm) and a RF-10AxL fluorescence detector. A 20 µl injection volume was used, an excitation wavelength of 390 nm, an emission wavelength of 480 nm and a flow rate of 1.2 ml/min. The aqueous solvent was 0.25% acetic acid, pH 3.5, and the organic solvent was 100% methanol. A linear gradient of 1525% methanol was performed over 15 min, followed by 5 min of 100% methanol to wash the column and 3 min of 15% methanol to equilibrate the column for the next run. The GSH peak was eluted at 12.8 min. Depletion of GSH in A549 cells was achieved by adding 0.1 mM L-buthionine-R,S-sulfoximine (BSO) for 24 h. This treatment with BSO has been previously shown to be non-toxic to cells (26).
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Results
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Rate of spontaneous loss of DNAhistone crosslinks
Previous studies have shown that aldehyde-induced DPC were hydrolytically labile and could be completely reversed by incubation at elevated temperatures (22,23). Our first experiments were performed to determine the rate of spontaneous hydrolysis of FA-induced DPC at physiological temperature and pH. The intrinsic instability of DPC was examined in crosslinks produced in cells and with in vitro formed DNAhistone crosslinks. Analysis of DPC stability in cell samples could be hampered by the presence of nucleases and proteases, activation of which could lead to an apparent loss of DPC. DNAhistone H1 crosslinks produced in vitro are free of hydrolytic enzymes, but they need to resemble those formed in cells. Figure 1
shows that the time courses of DPC formation by FA in vitro with histone H1 and in cultured A549 cells were very close, suggesting involvement of similar chemical groups in crosslink formation. DPC formation by FA in vitro was dependent on binding of histone to DNA, since dissociation of histoneDNA complexes in the presence of SDS or 0.8 M NaCl blocked crosslinking reactions (Figure 2
). It has been previously shown that 0.8 M NaCl completely blocks DNAhistone H1 binding (27). Bovine serum albumin (BSA), a protein that does not bind to DNA, was also incapable of forming DPC. Increasing the FA concentration to 1 or 5 mM did not lead to formation of BSADNA crosslinks (data not shown). To determine the hydrolytic stability of DPC, we lysed FA-exposed cells and in vitro formed DNAhistone H1 crosslinks in SDS and then measured levels of DPC at different time points following incubation at 37°C (Figure 3
). The overall kinetics of DPC loss with time were very similar between in vitro DPC and those produced in cells, as evidenced by very close t
values. The half-life was 17.7 h for DPC formed with purified histone H1 and 18.8 h for cellular DPC. The primary reason for the addition of SDS to the crosslink samples was to block nuclease activity in cell lysates. The presence of SDS also prevents reformation of broken crosslinks, since histones will immediately dissociate from DNA after crosslink rupture. Therefore, loss of DPC under these conditions is determined only by the intrinsic hydrolytic instability of FA-induced DPC. When the stability of DPC was assessed under conditions allowing histone H1 to remain bound to DNA after crosslink breakage, the loss of crosslinks was slower, with t
= 26.3 h. This suggests that under physiological conditions a fraction of previously broken DPCs between histones and DNA reform.

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Fig. 1. Time course of DPC formation by formaldehyde in A549 cells and in vitro. Cells or 1:5 mixtures of histone H1 with DNA were treated with 0.2 mM FA for the indicated periods of time. Samples were lysed in 1% SDS, sheared and frozen at 80°C. DPC were measured by the KCl/SDS precipitation assay.
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Fig. 2. Importance of proteinDNA binding for the formation of DPC. Histone H1DNA or BSADNA mixtures were reacted with 0.5 mM FA for 3 h in phosphate buffer, pH 7.0. Two sets of histone H1 samples additionally contained 1% SDS or 0.8 M NaCl. DPC were determined as described in Materials and methods.
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Fig. 3. Hydrolytic stability of DPC formed by FA in vitro and in cells. DPC were formed by exposure of A549 cells or DNAhistone H1 mixtures to 0.2 mM FA for 3 h. Cells and a portion of in vitro samples were lysed in 1% SDS and stability of crosslinks was examined over a 72 h period at 37°C. A fraction of DNAhistone H1 crosslinks was dialyzed overnight against 2x 500 vol phosphate buffer to remove unreacted FA. Stability of dialyzed DPC samples was examined as described above.
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DPC removal from control and GSH-depleted cells
The rate of DPC loss was determined by measuring the number of remaining crosslinks at different time intervals following removal of FA-containing medium. The overall rate of DPC removal from three human cell lines of different histological origin was very close, with t
averaging 12.5 h (Figure 4A
). After adjusting for spontaneous loss of DPC (t
= 26.3 h), we calculated that there was also an active repair process involved, which was removing DPC with an average t
= 23.3 h for the same three cell lines (Figure 4B
). Human peripheral blood lymphocytes lost DPC at a significantly slower rate (t
= 18.1 h) compared with established cell lines (Figure 4A and B
). The active repair component was calculated to remove DPC from lymphocytes with an average t
= 66.6 h (range 59.877.0 h), which is approximately three times longer than the half-life in other cells.

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Fig. 4. Half-life of DPC in FA-exposed cells. Cells were exposed to 0.2 mM FA for 3 h. FA-containing medium was removed and cells were collected for DPC determination at five different time points following addition of fresh medium. (A) Overall half-life of DPC; (B) calculated half-life of DPC attributed to active repair.
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In the next series of experiments we tested a possibility that intracellular levels of GSH could modulate the ability of cells to repair/remove FA-induced DPC. Two observations provided a basis for these experiments. Oleinick et al. (28) found that repair of radiation-induced DPC in A549 cells could be inhibited by depletion of GSH. It has also been reported that human lymphocytes contain lower amounts of GSH than other cells (29,30). We measured GSH as its fluorescent derivative with the thiol-specific dye mBBr following separation by HPLC (Figure 5
). BSO, a potent inhibitor of
-glutamylcysteine synthetase (31), was used to deplete cells of GSH. An actively growing population of control A549 cells had a large GSH peak, while BSO-treated cells did not show detectable levels of GSH. Based on the reported volume of A549 cells (3.6 pl) (32), the intracellular concentration of GSH in unexposed cells was calculated to be 4.6 mM. A loss of GSH did not result in any significant difference in the rate of repair of FA-induced DPC (Figure 6A
). The total number of DPC in BSO-treated cells was marginally lower than in controls. Long-term survival of GSH-depleted and control cells exposed to different concentrations of FA was not statistically different (Figure 6B
).

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Fig. 5. HPLC determination of GSH concentrations in control and BSO-treated A549 cells. BSO (100 µM) was added to cells for 24 h prior to sample collection. Cell lysates were treated with mBBr and a fluorescent derivative of GSH was quantified by HPLC. (A) Control cells; (B) BSO-treated cells. GSH, glutathione; R, reagent peak.
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Fig. 6. Effect of GSH depletion on the rate of removal of DPC and survival of A549 cells. Cells were treated with BSO for 24 h prior to FA exposures. Cell survival was determined by colony forming assay. (A) Time-dependent loss of DPC; (B) survival of FA-treated cells.
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DPC repair in XP cells
XP cells are deficient in nucleotide excision repair, a system which represents a major pathway to remove bulky DNA lesions (33). To test the involvement of this repair process in the elimination of DPC from cells, we compared the rates of DPC repair in normal HF/SV cells and XP-A and XP-F fibroblasts. XP-A cells have a severe defect in nucleotide excision repair and XPA protein is believed to be involved in the recognition of DNA lesions (33,34). XP-F cells are unable to perform the incision step at the 5'-position during nucleotide excision repair due to mutations in the XPF protein, which is part of the XPF/XRCC1 5'-excinuclease (33). Figure 7A
shows that the kinetics of DPC removal were very similar among all three cell lines. These results indicate that the nucleotide excision pathway is unlikely to be involved in repair of DNAhistone crosslinks. However, nucleotide excision repair may play a role in the repair of other forms of FA-induced DNA damage, since survival of FA-treated XP cells was somewhat lower compared with controls (Figure 7B
). We calculated that a 50% reduction in colony forming ability was achieved after exposure to 171 ± 77, 119 ± 55 and 94 ± 9 µM FA for controls, XP-A and XP-F cells, respectively. Only the difference in survival between XP-F and normal cells approached statistical significance (P = 0.08).

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Fig. 7. Kinetics of DPC removal and toxicity of FA treatment in nucleotide excision repair-deficient XP cells and normal HF/SV fibroblasts. In all experiments FA treatment was for 3 h. Toxicity was determined by the colony forming assay. (A) Time course of DPC removal from XP-A, XP-F and normal HF/SV fibroblasts; (B) survival of FA-treated cells.
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Effect of a proteosome inhibitor lactacystin on repair of DPC
Cells eliminate damaged or unnecessary proteins through a specialized proteolytic process (35). Protein targets are first tagged by ubiquitination and then cleaved by a high molecular weight protease complex (proteosomes). We examined the possibility that this normal process of protein degradation is involved in elimination of DPC. Lactacystin is a microbial product that has been shown to be a very potent and specific inhibitor of the proteosome (36). This compound inhibits all three major peptidase activities of the proteosome: caspase-like (reversibly) and chymotrypsin-like and trypsin-like (irreversibly) (37). Addition of micromolar concentrations of lactacystin resulted in a dose-dependent inhibition of DPC repair in both normal and XP-A fibroblasts (Figure 8
). There was a 3-fold decrease in the rate of DPC elimination by the active repair component at 10 µM lactacystin. This concentration of lactacystin has been shown to block the proteolytic function of nuclear proteosomes (38). Prolonged exposure to lactacystin resulted in cytotoxicity, evident as accelerated detachment of cells from the dishes. XP-A cells were particularly sensitive to lactacystin and t
values for these cells were calculated using the earliest post-exposure data points.

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Fig. 8. Suppression of DPC repair by lactacystin. Lactacystin was added to HF/SV and XP-A cells for 3 h prior to FA exposure and was present in all post-exposure incubations. Shown are data for activity of cellular repair expressed as t for DPC. 1, 0 µM; 2, 3 µM; 3, 10 µM lactacystin.
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Discussion
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The main goal of this investigation was to identify major factors that control removal of DPC formed by FA. We initially set out to determine whether spontaneous loss of DPC could represent one of the important contributors to the overall rate of DPC repair in cells. DNAhistone H1 crosslinks were used in all in vitro experiments, largely due to concerns about the presence of hydrolytic enzymes in DPC preparations from exposed cells. We believe that crosslinks with histone H1 represent a typical DPC from cells because: (i) histone H1 is one of the proteins most actively crosslinked by FA in cells (16); (ii) the overall kinetics of DPC formation with histone H1 and in cells were very close, indicating similar formation mechanisms; (iii) the stabilities of histone H1 and cellular DPC were almost identical in the presence of SDS, which suggests identical chemical structures of these adducts. At 37°C and under physiological ionic/pH conditions the half-life of DNAhistone H1 crosslinks was 26.3 h. In a separate set of samples SDS was included in the incubation mixtures to force release of histones from DNA immediately after DPC breakage. The loss of DPC was faster under these conditions, with average t
= 18.3 h. Based on the differences in the rates of DPC disappearance under histoneDNA binding (no SDS) and dissociating (plus SDS) conditions, we calculated that ~30% of spontaneously broken DNAhistone H1 crosslinks are re-established under physiological conditions. The similarity of DNAhistone H1 crosslinks to DPC produced in cells suggests that the percentage of reformed DPC in chromatin is probably in the same range.
The rate of DPC disappearance from cells was almost twice as fast as the estimated hydrolytic instability. This suggests that cells also possess an active repair process for DPC removal. Half-life values were close among three different cell types (fibroblasts and kidney and lung cells). The overall half-life of DPC in peripheral lymphocytes was ~50% longer than that in cultured human cells. This translates into three times less efficient removal of DPC by the active repair component (66.6 versus 23.3 h). Lymphocytes are also known to have inefficient nucleotide excision repair (39). Lymphocytes are terminally differentiated cells and, because there is no danger of converting DNA lesions into mutations, the presence of active DNA repair in these cells may not be biologically important. Damaged lymphocytes can also be easily replaced through production of new cells. The rate of DPC loss from FA-exposed cells found in our experiments appears to be slower than reported by other investigators (7,21,40). The results of time course studies reported previously were presented either as the rate of DNA elution through filters (21,40) or as the length of DNA `tails' in a Comet assay (7), however, the exact relationship between these parameters and the number of DPC is unknown and could be non-linear. All rate calculations in this work were done using numbers of crosslinks per 108 bp. FA is also known to cause extensive proteinprotein crosslinking (41), which can result in non-specific trapping of high molecular weight DNA fragments during alkaline elution or single cell electrophoresis. The overall rate of increase in DNA mobility in these assays will then be a result of both loss of DPC and dissociation of crosslinked protein structures. The sensitivity of DPC measurement by KCl/SDS precipitation is also higher than that for other methods (7,24).
Depletion of GSH from A549 cells did not significantly change the rate of DPC repair or initial levels of DPC. The later contradicts findings by Lam et al. (42) and Casanova and d'A Heck (43), who reported higher DPC formation in the nasal mucosa of animals whose GSH was depleted by acrolein or phorone exposure, respectively. One of the reasons for this discrepancy could be related to the metabolism of FA. Oxidation of FA to formic acid by formaldehyde dehydrogenase requires formation of S-formylglutathione (44). While nasal epithelium is known to oxidize FA at a high rate (43), A549 cells were not tested for this activity. If these cells are poor metabolizers of FA, then changes in GSH levels would not affect intracellular concentrations of FA and levels of DPC. The use of different approaches to cellular GSH depletion in our work and in previous studies (42,43) could also be responsible for the different experimental findings. In our experiments cells were depleted of GSH using BSO, which is a non-toxic, specific inhibitor of a key enzyme in GSH biosynthesis (31). Use of less specific depletors can lead to additional toxicological consequences. For example, acrolein is also known to be toxic (45), it forms DNA and protein adducts (46,47) and affects many GSH-independent physiological functions in exposed cells (48). Depletion of GSH by BSO did not change the cytotoxicity of FA, indicating that GSH plays no significant role in scavenging FA. Lack of a protective role of GSH against FA-induced toxicity is probably related to the very low stability of S-formylglutathione, which is the major adduct of FA with GSH formed under physiologically relevant conditions (49).
In order to identify the repair pathway involved in removal of DPC, we compared the rate of DPC repair in normal and nucleotide excision repair-deficient XP-A and XP-F fibroblasts. Both the overall rates of removal and active repair rates were very similar among these three cell lines, suggesting that nucleotide excision repair is unlikely to be involved in removal of FA-induced DPC. Similar kinetics of DPC loss in XP-A skin fibroblasts and human bronchial epithelial cells were also reported by Graftstrom et al. (21). We have also tested involvement of the proteosome in removal of DPC from FA-exposed cells. Proteosomes are ubiquitous protease complexes of high molecular weight containing a 20S catalytic subunit and some additional components (35). Proteosomes play a major role in proteolytic degradation of proteins that are no longer needed or are damaged as a result of stressful conditions. A possibility of proteosome involvement in DPC removal was studied in experiments in which the proteosome 20S catalytic subunit was inactivated by lactacystin. Addition of lactacystin resulted in a dose-dependent increase in the half-lives of DPC in both normal and XP-A fibroblasts. Desai et al. (38) reported that lactacystin inhibited proteolytic degradation of topoisomerase I crosslinks with DNA induced by stabilization of DNAtopoisomerase I intermediates by camptothecin. `Frozen' topoisomeraseDNA complexes were ubuquitinated prior to proteolytic degradation. Based on the inhibitory effect of lactacystin on the disappearance of FA-induced DPC, we think that removal of crosslinked histones proceeds through a similar mechanism. The presence of proteosomes in the nucleus has been demonstrated (50,51) and putative nuclear localization signals were identified in some proteosome subunits (52). Palmer et al. (51) measured the distribution of proteosomes inside the cells using anti-proteosome antibodies and they found that the density of labeling in the nuclear compartment was almost as high as in the cytoplasm. Although the large size of proteosomes would seem to impose significant restrictions on their ability to function in the dense structure of the nucleus, the rapid destruction of ubiquitinated topoisomeraseDNA complexes (t
= 12 h) (38) indicates that nuclear proteosomes are highly active. Because DNA-crosslinked proteins remain fixed in a specific position in the nucleus, proteosomes are expected to be able to diffuse throughout the nucleoplasm seeking proteins targeted for destruction.
Proteosomal degradation of DNA-linked proteins cannot completely remove DPC and peptideDNA or amino acidDNA crosslinks will be the end-products of this process. The question arises what repair pathway can deal with these residual crosslinks? In the case of DNAtopoisomerase I covalent complexes the residual peptide or tyrosineDNA phosphodiesters can be hydrolyzed by a recently discovered phosphodiesterase (53). An attractive possibility for dealing with residual FA-induced DPC could be the involvement of nucleotide excision repair. We found that the LD50 in XP-F cells was approximately 1.8 times lower compared with normal fibroblasts, which might result from the presence of unrepaired peptideDNA adducts. XP-F cells contain mutated XPF protein which leads to loss of the ability of nucleotide excision complexes to make an incision 5' of the lesion (33). The colony forming ability of XP-A fibroblasts was not very different from controls. XPA protein is involved in the initial steps of nucleotide excision repair (33) and it is possible that recognition of residual DPC proceeds through a different mechanism. However, the actual excision and filling steps could be carried out by regular members of the XP family of proteins and this suggestion is consistent with a lower survival of XP-F cells following exposure to FA.
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Notes
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1 To whom correspondence should be addressed Email: anatoly_zhitkovich{at}brown.edu 
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Acknowledgments
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This work was supported by a start-up grant to A.Z. from Brown University.
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Received January 12, 2000;
revised April 14, 2000;
accepted May 1, 2000.