Mutagenesis, Vol. 16, No. 6, 455-459,
November 2001
© 2001 UK Environmental Mutagen Society/Oxford University Press
Lysis of whole blood in vitro causes DNA strand breaks in human lymphocytes
2 Rowett Research Institute, Greenburn Road, Bucksburn, Aberdeen AB21 9SB and 1 Safety Assessment, AstraZeneca R&D, Bakewell Road, Loughborough LE11 5RH, UK
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Abstract |
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DNA damage in lymphocytes, as measured by alkaline single cell gel electrophoresis (pH 12.7), is greatly increased by the concurrent lysis of whole blood in both freshly isolated samples and in PHA-stimulated cultures over a period of 7 days. Further, there is a marked progressive increase in DNA damage with time in PHA-stimulated lymphocytes cultured in whole blood even when the lymphocytes are separated before analysis; no such increase is seen in lymphocytes cultured alone. This indicates that there are components in whole blood that can cause DNA damage in lymphocytes, with granulocytes and lysis of red blood cells likely candidates. The DNA damage is greatly reduced in granulocyte-depleted whole blood cultures, but even in these significant increases are seen at later sampling times. Consequently, careful sample preparation is of paramount importance if the Comet assay is to be successfully used to assess DNA damage in human peripheral blood lymphocytes. Further, the progressive increase in DNA damage in whole blood cultures may influence other methods using lymphocytes for population biomonitoring and may be significant for in vitro genotoxicity testing.
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Introduction |
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Human peripheral blood lymphocytes (PBL) have been used to monitor environmentally induced genetic damage by a variety of methods, including cytogenetic end-points such as micronuclei, chromosome aberrations, sister chromatid exchanges (Perera and Whyatt, 1994) and somatic mutation (Cole and Skopek, 1994
). These involve monitoring lymphocytes after various times in culture, either as a separated mononuclear cell fraction or as whole blood, with or without lectin stimulation, depending on the particular end-point. Most commonly, T cells stimulated to divide by phytohaemagglutinin (PHA) have been used. Despite extensive use of lymphocytes in these systems, to our knowledge little work has addressed how time in culture or the system employed (whole blood or separated lymphocytes) influences DNA damage. Time in culture and cell division may influence pre-existing levels of damage or response to genotoxic insult, as quiescent lymphocytes are functionally excision repair-deficient due to low endogenous deoxyribonucleoside levels (Green et al., 1994). Findings such as this and anecdotal evidence that contamination of mononuclear cell fractions with red blood cells increases DNA damage estimated by the Comet assay led to the present study. Single cell gel electrophoresis (SCGE), known as the Comet assay, is a rapid, sensitive, reliable and inexpensive technique for measuring DNA strand breaks in individual cells (McKelvey-Martin et al., 1993). Cells are embedded in low melting point agarose on a microscope slide, lysed with detergent and treated with high salt. This results in the formation of nucleoids containing non-nucleosomal but still supercoiled DNA. The slides are then exposed to alkaline electrophoresis. Breaks in the DNA cause local relaxation of the supercoiled genetic material, enabling loops of DNA to be pulled towards the anode during electrophoresis. Comets are visualized by fluorescence microscopy after staining with the fluorescent DNA-binding dye 4',6-diamidine-2-phenylindole dihydrochloride (DAPI). The relative intensity of fluorescence in the tail is proportional to the frequency of DNA strand breaks (Olive et al., 1990). The Comet assay is frequently used in the field of genetic toxicology for the evaluation of in vitro and/or in vivo genotoxicity. A wide range of cells (normal and transformed) have been used, including human, animal and plant cells. In addition, the Comet assay has been shown to provide useful information in environmental biomonitoring, assessing genetic hazards of pollutants. The range, sensitivity and specificity of the alkaline Comet assay have been increased by including bacterial DNA repair enzymes which detect altered bases (Collins et al., 1993; Duthie and McMillan, 1997). |
Materials and methods |
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Histopaque-1077, L-glutamine, penicillin G, sodium pyruvate and streptomycin sulphate were obtained from Sigma (Poole, UK). Dutch modified RPMI 1640 medium was obtained from ICN Flow Laboratories (Irvine, UK). Fetal calf serum (heat-inactivated) was obtained from Globepharm Ltd (Surrey, UK). Low melting point (LMP) and high melting point (HMP) agarose, together with Nunc sterile tissue culture plastics, were supplied by Gibco Life Technologies Inc. (Paisley, UK). LymphoPrep was supplied by Nycomed UK (Birmingham, UK). Murex Diagnostics Ltd (Dartford, UK) provided HA15 PHA. Recombinant interleukin (Aldesleukin, 18x106 IU) was purchased from Euro-Cetus UK Ltd (Harefield, UK). DAPI was obtained from Boehringer Mannheim (Lewes, UK). Frosted microscope slides were from Richardson Supply Co. (London, UK). Dynabeads M-450 CD 15 were from Dynal Ltd (Wirral, UK).
The effect of storage and red blood cell contamination on DNA strand breakage in human lymphocytes
DNA damage: isolated lymphocytes versus lymphocytes in whole blood
Isolated human lymphocytes were prepared from a `finger prick' sample from eight volunteers as follows. Whole blood (30 µl) was mixed with 1 ml of RPMI 1640 medium supplemented with 10% fetal calf serum (FCS), underlayed with 100 µl Histopaque and centrifuged at 200 g for 3 min at 4°C. The lymphocyte-containing `buffy coat' (100 µl) was washed once in phosphate-buffered saline (PBS), pH 7.4, centrifuged as before and resuspended in LMP agarose (85 µl) for Comet analysis. Lymphocytes in whole blood were obtained by mixing 30 µl of whole blood with 1 ml of RPMI 1640 medium supplemented with 10% FCS and centrifuging the sample for 3 min at 200 g at 4°C. The pellet was washed once in PBS, pH 7.4, centrifuged as before and resuspended in LMP agarose for Comet analysis (Duthie and McMillan, 1997).
The effect of storage on DNA strand breakage in human lymphocytes
Venous blood samples (2x10 ml) were collected from the same eight volunteers. Fresh blood was immediately taken for Comet analysis as described above. One vacutainer from each volunteer was stored in a polystyrene box at either 4°C or room temperature. Comet analysis was carried out on blood after 24 and 48 h storage.
DNA damage in PHA-stimulated lymphocytes grown as a separated mononuclear fraction, in granulocyte-depleted blood culture and in whole blood cultures
Lymphocytes grown as a separated mononuclear fraction
Venous blood (1x10 ml) was collected from the same eight subjects. Lymphocytes were immediately isolated from 5 ml of whole blood. The blood was centrifuged at 1500 g for 15 min at 4°C. The `buffy coat' was removed and diluted 1:1 with RPMI medium before layering onto an equal volume of LymphoPrep lymphocyte separation medium (specific gravity 1.077 ± 0.001 g/ml) and centrifugation at 700 g for 30 min at 20°C. The lymphocytes were removed to a fresh centrifuge tube, washed using medium and spun for a further 15 min under the same conditions. The supernatant was decanted and the cells resuspended in RPMI medium containing 10% FCS before being counted using a Neubauer Improved Haemocytometer. Isolated lymphocytes were resuspended at 1x105 cells/ml and stimulated to divide in medium containing 100 µg/ml pyruvic acid, 2 mM L-glutamine, 100 µg/ml streptomycin, 100 U/ml penicillin, 20% (v/v) FCS, 100 U/ml interleukin and 1% PHA. All cell cultures were maintained at 37°C in a humidified atmosphere of 95% air/5% CO2. Comet analysis was performed on days 0, 1, 2, 3, 4 and 7. Samples (1 ml) were spun at 200 g for 3 min at 4°C. The supernatant was discarded and the pellet washed in 1 ml of PBS, centrifuged as before and resuspended in LMP (85 µl) agarose for Comet analysis as described.
Lymphocytes grown in granulocyte-depleted whole blood cultures
Depleted whole blood cultures were prepared by removing granulocytes from 2.5 ml of venous blood sample using Dynabeads M-450 with a covalently bound mouse anti-CD15 monoclonal antibody. Dynabeads were washed in washing buffer (PBS with 2% FCS), separated on a Dynal magnetic particle concentrator and resuspended in buffer. Dynabeads (12x107) were added to 2.5 ml of whole blood, previously diluted 1:1 with washing buffer, mixed gently and incubated for 20 min at 4°C on a rolling mixer. After incubation, the sample was placed in a Dynal magnetic particle concentrator to remove Dynabead-bound granulocytes and the supernatant transferred to a fresh tube. These samples were centrifuged at 700 g for 20 min, resuspended in 5 ml of RPMI 1640 medium with 20% (v/v) FCS, 100 µg/ml pyruvic acid, 2 mM L-glutamine, 100 µg/ml streptomycin, 100 U/ml penicillin, 100 U/ml interleukin and 1% PHA. The cell suspension was added to 20 ml of medium and grown at 37°C in a humidified atmosphere of 95% air/5% CO2. Comet analysis was carried out on days 0, 1, 2, 3, 4 and 7 following lymphocyte isolation from 1 ml of culture, i.e. underlaying the sample with 100 µl Histopaque, centrifuging at 200 g for 3 min at 4°C, washing the pellet in 1 ml of PBS and resuspending in LMP.
Lymphocytes grown in whole blood cultures
Cultures were established by mixing 2.5 ml of whole blood with 22.5 ml of RPMI 1640 medium containing 20% (v/v) FCS, 100 µg/ml pyruvic acid, 2 mM L-glutamine, 100 µg/ml streptomycin and 100 U/ml penicillin. The cells were stimulated to divide by adding interleukin (100 U/ml) and PHA (1%) to the medium. Whole blood cultures from each volunteer were maintained at 37°C in a humidified atmosphere of 95% air/5% CO2. Comet analysis was carried out on days 0, 1, 2, 3, 4 and 7. Lymphocytes were sampled in two different ways. (i) Blood culture samples (1 ml) were removed from the flasks and spun at 200 g for 3 min at 4°C. The supernatant was discarded, the pellet washed in 1 ml of PBS, centrifuged and resuspended in LMP agarose for Comet analysis. (ii) Lymphocytes were isolated before Comet analysis from 1 ml of whole blood culture by underlaying with 100 µl of Histopaque and centrifuging at 200 g for 3 min at 4°C. The supernatant was discarded, the pellet washed in 1 ml of PBS and centrifuged before resuspension in LMP agarose for Comet analysis.
DNA strand breakage measured using the Comet assay
Cells were suspended in 85 µl of 1% (w/v) LMP agarose in PBS, pH 7.4, at 37°C and immediately pipetted onto a frosted glass microscope slide precoated with a layer of 1% (w/v) HMP agarose similarly prepared in PBS. The agarose was allowed to set for 5–10 min at 4°C and the slide incubated in lysis solution (2.5 M NaCl, 10 mM Tris, 100 mM Na2EDTA, NaOH to pH 10.0 and 1% v/v Triton X-100) at 4°C for 1 h to remove cellular proteins. After lysis, the slides were aligned in a 260 mm wide horizontal electrophoresis tank containing buffer (1 mM Na2EDTA and 0.3 M NaOH, pH 12.7) for 40 min before electrophoresis at 25 V for 30 min (at an ambient temperature of 4°C with the temperature of the running buffer not exceeding 15°C). The slides were washed three times at 4°C for 5 min each with neutralizing buffer (0.4 M Tris–HCl, pH 7.5) before staining with 20 µl of DAPI (5 µg/ml) (Duthie and McMillan, 1997).
Quantitation of the Comet assay
Nucleoids were scored visually using a Zeiss Axioskop fluorescence microscope. One hundred comets from each slide (scored at random) were classified into one of five classes according to the relative intensity of fluorescence in the tail and given a value of 0–4 (from undamaged, 0, to maximally damaged, 4). The total score for 100 comets can range from 0 (all undamaged) to 400 (all maximally damaged). This method of visual classification has been extensively validated by comparison with comets selected using computerized image analysis. Representative images of comet classes were analysed (Komet 3.0; Kinetic Imaging Ltd, Liverpool, UK) and the percentage of fluorescence in the comet tail (representing the fraction of DNA in the tail) plotted against the total score for 100 comets in each class. There is a clear linear relationship (r = 0.987) between visual classification and the percentage of DNA measured in the tail (Duthie and McMillan, 1997).
Flow cytometer analysis
The granulocyte-depleted whole blood cultures were analysed on a FACScan (Becton-Dickinson, Oxford, UK) flow cytometer, enabling quantitation of independent cell groups. Briefly, 1 ml of sample was added to 1.4 ml of 1x lysing solution (FACS Lysing Solution; Becton Dickinson), mixed and incubated for 10 min at room temperature in the dark. After incubation, the sample was centrifuged (300 g for 5 min), the supernatant discarded, the pellet resuspended in 1 ml of PBS and centrifuged (200 g for 5 min). The supernatant was discarded and the pellet resuspended in 0.5% formaldehyde in Sheath fluid (Becton Dickinson) for flow cytometer analysis. The lymphocyte, monocyte and granulocyte populations in each sample were identified and gated manually by visual inspection of the size (forward scatter) versus complexity/granularity (side scatter) analyses.
Statistical analysis
Student's t-test and ANOVA, together with Tukey's honestly significant difference test, were carried out as appropriate using SPSS 8.0 for Windows.
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Results |
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The presence of whole blood induced a significant increase in DNA damage (single-strand DNA breaks and/or alkali-labile sites measured using the Comet assay) in lymphocytes obtained from a finger prick sample. DNA damage was elevated 10-fold in lymphocytes analysed in the presence of red blood cells compared with isolated lymphocytes (Figure 1
).
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A significant increase in DNA damage was observed after 24 h storage at both 4°C and room temperature. A further increase in DNA damage after 48 h was only observed after storage at room temperature (Figure 2
).
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This increase in DNA damage might have been due to release of components from other cell types present in whole blood during the lysis step of the Comet assay. To test this hypothesis, PHA-stimulated lymphocytes were grown either as a separate mononuclear fraction or as whole blood cultures. Samples were obtained on days 0, 1, 2, 3, 4 and 7 for Comet analysis. There was a significant increase (P < 0.0005) in endogenous DNA strand breakage from day 0 between lymphocytes grown in whole blood cultures, compared with those grown as a mononuclear fraction or with those isolated from a whole blood culture. There was no significant difference between lymphocytes grown as a mononuclear fraction and lymphocytes isolated from whole blood culture on days 0 and 1. However, there was a linear increase in DNA damage in lymphocytes isolated from whole blood cultures after 2 days (Figure 3
).
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4. **P < 0.005, differences in DNA breakage in lymphocytes grown in mononuclear cell culture (black bars) or lymphocytes isolated from whole blood culture (hatched bars). *P < 0.005, differences in DNA breakage in lymphocytes isolated from whole blood cultures (hatched bars) compared with lymphocytes sampled from whole blood cultures (white bars).Two cell types are potentially responsible for induction of DNA damage in whole blood cultures; granulocytes and erythrocytes. To determine which caused induction of DNA damage, whole blood cultures were compared with whole blood cultures depleted of granulocytes. To ensure that Dynabead treatment in fact depleted granulocytes, the samples were analysed on a FACScan flow cytometer. The granulocytes comprised ~60% of total white cells in whole blood cultures, as expected, compared with 1.4% in Dynabead-depleted cultures (Figure 4
). Samples were obtained on days 0, 1, 2, 3, 4 and 7 for Comet assay analyses. Lymphocyte DNA strand breakage was significantly higher after culture for 1 day in whole blood compared with lymphocytes grown in granulocyte-depleted blood (Figure 5). Similarly, lymphocytes grown for 2 days in granulocyte-depleted blood showed higher DNA damage compared with those grown as a mononuclear fraction (Figure 5). There was no increase in DNA strand breakage in lymphocytes grown for 7 days in isolated culture. Conversely, DNA damage increased linearly in lymphocytes from both whole blood and granulocyte-depleted blood. Strand breakage was significantly higher in lymphocytes from whole blood (Figure 5).
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Discussion |
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Our results clearly show that endogenous DNA strand breakage (measured using the alkaline Comet assay) is significantly increased in lymphocytes upon storage at either 4°C or room temperature when compared with freshly isolated cells. This is in contrast to a previous study using the Comet assay to detect both background and chemically induced DNA damage in lymphocytes that indicated that human blood samples could be stored under these conditions for up to 4 days (Anderson et al., 1997
). Subtle differences in assay conditions may partly explain this discrepancy. In the present study nucleoids were allowed to unwind for 40 min, compared with 20 min in the earlier study, which may allow greater relaxation of the DNA. Similarly, slides were electrophoresed for 30 min, compared with 20 min in the experiment by Anderson et al., presumably resulting in different rates of migration through the agarose. Finally, four times as many images per gel were analysed in the work presented here, when compared with the previous study reported by Anderson and co-workers (Anderson et al., 1997). Our results suggest that human blood samples cannot be stored for more than 24 h, even at 4°C. Rather, lymphocytes should be used fresh or cryopreserved and stored at –196°C for future analysis. DNA damage in lymphocytes is greatly increased by the concurrent lysis of whole blood and this level of damage is very similar in freshly isolated samples and PHA-stimulated cultures over a period of 7 days. Further, although the level of damage declines slightly with time in PHA-stimulated separated lymphocyte cultures, there is a marked progressive increase with time in whole blood cultures even when the lymphocytes are separated before analysis.
Although it would seem most likely that we are observing DNA damage in lymphocytes caused by some factor(s) present in whole blood, it is also possible that the differences may be due to different cell populations being compared in the various preparations used in these experiments. Whole blood contains ~5x109 red blood cells (RBC) and 4–10x106 white blood cells (WBC) per ml; of the WBC, 25–40% are lymphocytes, with neutrophils (the major granulocyte type) comprising the majority of the rest. Mononuclear cell preparations separated on Ficoll gradients should contain at least 90% lymphocytes (Gurtoo et al., 1975), so it is possible that the differences in response between separated lymphocytes and whole blood seen on day 0 represent differences in DNA damage in lymphocytes and the other WBC present. However, background damage assessed by the Comet assay appears to be very similar in freshly isolated lymphocytes, phagocytes and monocytes from normal individuals (Vijayalaxmi et al., 1993; Hannon-Fletcher et al., 2000). The available evidence, therefore, indicates that different responses of the various WBC types do not account for the findings in freshly isolated cells.
The WBC populations in PHA-stimulated whole blood cultures are complex (O'Donovan et al., 1995b), with the monocytes and granulocytes disappearing within the first 2 days; by ~4 days the majority of the living cells are probably dividing T cells, as would be expected. The behaviour of T cells with respect to DNA damage is also complex. Resting T cells are effectively excision repair deficient through having low intracellular deoxyribonucleotide pools, but this is up-regulated after mitogen stimulation (Green et al., 1994). Although complex, there is no evidence to suggest that background levels of DNA damage increase 4–5 days after PHA stimulation in comparison with the levels in unstimulated cells (Green et al., 1992, 1994). This agrees with our findings with separated lymphocytes and would appear to reflect DNA damage in lymphocytes alone; immediately after preparation the population comprises a mixture of T and B cells, but 7 days later >99% are T cells (O'Donovan et al., 1995a). The most obvious conclusion is that the increases in damage seen with time of culture in whole blood reflects DNA damage in lymphocytes caused by some factor(s) in whole blood, not differences in the cell populations being examined.
There are two obvious candidates for the components in whole blood that may be capable of causing the observed DNA damage: neutrophils and lysis of RBCs. Both appear to increase lymphocyte DNA strand breakage in this study. Neutrophils, the most numerous WBC type in the peripheral blood of healthy individuals, undergo an oxidative burst when activated, releasing various reactive oxygen species, including the superoxide anion and H2O2 into the extracellular environment. In addition, they have myeloperoxidase that can also generate reactive nitrogen species (Byun et al., 1999). Activated mouse neutrophils produce DNA damage in adjacent cells in vitro which is longer lasting than that produced by H2O2 alone (Schacter et al., 1988). Moreover, granulocytes incubated at a 1:1 ratio with plasmocytoma cells produced DNA single-strand breaks equivalent to a 1 min incubation with 20 µmol/l H2O2 (Schacter et al., 1988). Similar results were obtained when activated mouse neutrophils were co-cultivated with B cells using unscheduled DNA synthesis as the end-point (Janz and Schachter, 1993). Therefore, lysis of neutrophils during comet preparation from whole blood or prolonged incubation in whole blood could contribute to DNA damage seen in lymphocytes, although their activation status in these culture conditions has not been determined.
The overwhelming majority of blood cells are RBC, so even relatively small amounts of DNA-damaging material released from them could have marked effects on the WBC population. Conversion of oxyhaemoglobin to methaemoglobin generates superoxide and H2O2 and, consequently, RBCs have a range of protective mechanisms, including superoxide dismutase, catalase and glutathione peroxidase (Halliwell and Gutteridge, 1985). During lysis, either suddenly during preparation of samples for Comet analysis or with time in culture, large amounts of haemoglobin will be released into the medium. Perhaps surprisingly, we could find no published work on possible oxidative damage generated by haemolysis in vitro.
Overall, it appears that both granulocytes, particularly neutrophils, and RBC lysis contribute to the increased DNA damage seen in lymphocytes in the present study and that the effects could be additive. Since granulocyte numbers are known to decline rapidly with time in whole blood culture, it is possible that they contribute more to the DNA damage seen during the first few days, with progressive red blood cell lysis responsible for the increases at later sampling times. Platelets, the third major cellular component of blood, are not likely to contribute to the DNA damage, since it has been reported that they can act as scavengers of neutrophil-derived oxidants (Dallegri et al., 1989) and that this may represent a natural defence mechanism against neutrophil-mediated oxidative stress.
In conclusion, the findings in the present study confirm the anecdotal evidence that contamination of lymphocyte preparations with whole blood significantly affects the levels of DNA damage as measured by the Comet assay. In practice, this means that lymphocytes must be prepared extremely carefully if the Comet assay is to be used as an accurate tool for human population biomonitoring. Further, methods for measuring DNA damage in human peripheral blood lymphocytes such as cytogenetics and assays for point mutations may be significantly influenced by the culture conditions employed before the end-points are measured. In this context, it has been reported that both chromosome aberrations (Ivanov et al., 1973) and micronucleus frequency (Lee et al., 1999) are increased in lymphocytes in heparinized blood samples after 3 and 5 days storage, respectively. Finally, for in vitro tests for genotoxicity the possibility must exist that agents causing cell lysis may cause DNA damage through indirect mechanisms.
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Acknowledgments |
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This work was funded by SERAD.
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Notes |
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2 To whom correspondence should be addressed. Tel: +44 1224 712751; Fax: +44 1224 716629; Email: sd{at}rri.sari.ac.uk
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References |
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Received on April 10, 2001; accepted on June 14, 2001.
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