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Department of Nutritional Science, Fu-Jen University, HsinChuang 242, Tapei, Taiwan, ROC
* To whom correspondence should be addressed. E-mail: 034825{at}mail.fju.edu.tw.
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ABSTRACT |
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 TOP ABSTRACT IntroductionMaterials and MethodsResultsDiscussionLITERATURE CITED |
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Introduction |
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TOPABSTRACTIntroductionMaterials and MethodsResultsDiscussionLITERATURE CITED |
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Although the exact causes of the accumulation of mtDNA large deletions remain elusive, oxidative stress may relate to altered mtDNA genome integrity (14). Sublethal redox stress may result in a dramatic loss of mtDNA molecules or in deletions or loss of function by base modification in the mtDNA (15,16). The smaller size of partially deleted molecules could have a replicative advantage, which may lead to the accumulation of mtDNA large deletions in aging tissues (17,18). This increased mtDNA proliferation requires regulated nuclear gene expression. The mt transcriptional factor A (TFAM) is one of the nuclear-encoded regulators of mtDNA replication and transcription (19). TFAM expression is coordinated and regulated by a specific set of transcription factors such as the nuclear respiratory factor (NRF)-1 and NRF-2 (20). Mt single-strand DNA-binding protein and mt polymerase r are 2 other proteins crucial for mtDNA replication, repair, and recombination (21). Redox stress can activate NRF to induce production of TFAM (22). mtDNA biogenesis is then promoted, as evidenced by increased mtDNA copy number in rat liver (23), rat hippocampus (24), and senescent cells or aging tissues (25–28). Under oxidative stress, both nuclear and mt gene expressions work in concert to regulate mtDNA biogenesis so as to compensate for defective mtDNA.
Our previous study showed that folate deprivation promoted oxidative injuries in liver mitochondria, including elevated protein carbonyl levels, accumulation of mtDNA large deletions, and superoxide overproduction (29). Rats given folic acid supplementation had fewer mtDNA4834 deletions in hepatic tissue upon chemotherapeutic drug treatments (30) or in aging liver tissues (31). As growing evidence supports folate's role as an antioxidant (32,33), the physiological consequences of dietary folate deprivation for oxidative stress (34) suggest that deprivation may also lead to accumulation of mtDNA deletions in other tissues. Little, however, is known about the impact of folate deprivation on genetic modifications of mitochondria in various tissues of young rats. This study aimed to characterize changes in the mtDNA4834 deletion and mtDNA contents in 5 tissues of young rats during dietary folate deprivation. In light of the fact that low folate intake leads to elevated homocysteine (Hcy) concentrations in humans (35) and animals (34), plasma Hcy levels of rats were measured to indicate functional folate deficiency. We tested the hypothesis that folate deprivation-induced mtDNA changes are linked to subcellular folate levels, oxidative stress, and the expression of factors responsive to mt biogenesis.
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Materials and Methods |
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TOPABSTRACTIntroductionMaterials and MethodsResultsDiscussionLITERATURE CITED |
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Preparation of cytosolic and mt fractions from various tissues of rats. Mitochondria were isolated from various tissues of rats by conventional differential centrifugation in buffer containing 250 mmol/L sucrose, 5 mmol/L Tris-HCl (pH 7.4), and 1 mmol/L EGTA (37). Briefly, tissue homogenates were prepared in this buffer by using a Potter-Elvehjem-type homogenizer. The homogenate was centrifuged twice at 5000 x g; 10 min at 4°C and the mitochondria were then pelleted by centrifugation at 20,000 x g; 20 min. The purity was determined by measuring the relative specific activities of glutamate dehydrogenase in the supernatant (cytosolic fraction) and mt pellet, as described previously (37). Yields of mt fractions, as reflected by protein content (38), were similar in each experimental group.
Folate and Hcy assay. Blood and tissue samples for folate analysis were prepared according to Varela-Moreiras and Selhub (39). After incubation of the thawed sample extracts with chicken pancreas conjugase (v:v 4:1) at 37°C for 6 h, a microbiologic assay was performed using cryoprotected Lactobacillus casei in 96-well microtiter plates (40). Absorbance was detected at 600 nm in an MRX model ELISA reader (Dynatech Laboratories). Hcy concentrations in plasma samples were analyzed by fluorescence polarization immunoassay and an Abbott AxSYM system (Becton Dickinson).
Analysis of mtDNA4834 in various rat tissues. The quantity of the mtDNA4834 deletion was determined by coamplifying the mtDNA displacement-loop (D-loop) and mtDNA4834 deletion in a real-time PCR assay. Primers for each were previously described by Branda et al. (30). The degree of mtDNA4834 deletion was quantified with a deletion probe [DYXL-5'-(12952) TCACTTTAATCGCCACATCCATAACTGCTGT (12982)-3' BHQ1] and mtDNA probe [6FAM (15795) 5'-TTGGTTCATCGTCCATACGTTCCCCTTA (15822)-3' BHQ1]. PCR amplification was carried out in a 20-µL reaction volume consisting of TaqMan Universal Master mix (4 µL), 200 nmol/L each mtDNA4834 deletion primer, 50 nmol/L each D-loop primer, and 100 nmol/L each mtDNA4834 deletion and D-loop probe primer. The cycling condition included an initial phase of 2 min at 50°C, 10 min at 95°C, then 40 cycles of 15 s at 95°C and 0.5 min at 72°C. The fluorescence spectra were monitored by the LightCycler Detection system with Sequence Detection software version 4 (LightCycler, Roche Diagnostics).
Qualification of mtDNA4834 deletion by DNA sequencing. After the reaction, we separated the desired amplified DNA fragments by electrophoresis in a 2% agarose gel at 100 V for 40 min. The mtDNA bands were excised from the gel, placed on the filter of a Spin-X Centrifuge Tube filter (Costar), frozen at –20°C, and recovered by centrifugation (10,000 x g; 1 min). DNA sequencing was performed with a Big Dye Terminator Cycle Sequencing kit (Perkin Elmer) and an ABI 377 automated sequencer (Applied Biosystems).
Quantification of mtDNA4834 deletion.
The cycle at which a significant increase in normalized fluorescence was first detected was designated as the threshold cycle number (Ct). The ratio of mtDNA4834 deletion to mtDNA was calculated with 
Ct (= mt Ct del – mt Ct D-loop); a smaller Ct indicates more deletions. The relative expression (RE) indicates the factorial difference in deletions between the 2-wk FD, 4-wk FD, and pooled control groups. RE was calculated as 2–Ct, where Ct = Ct mtDNA deletion in FD group – Ct mtDNA deletion in the control.
Analysis of relative amount of mtDNA content.
The content of mtDNA (abundance) relative to nuclear genomic DNA was determined by coamplifying the mt D-loop and the nuclear-encoded ß-actin gene by real-time PCR assay. Primers for each were previously described by Branda et al. (30). The amount of ß-actin gene was quantified by a fluorescent probe [DYXL 5'-(3347) CGGTCGCCTTCACCG-TTCCAGTT (3325)-3' BHQ1]. PCR amplification and the cycling conditions were as described above. The fluorescence spectra were monitored by LightCycler (Roche Diagnostics). The ratio of mtDNA to genomic DNA content was calculated with Ct (mt Ct D-loop – nuclear Ct ß-actin). A smaller Ct indicates a less relative mtDNA content. RE indicates the factorial difference in mtDNA content between the 2-wk FD, 4-wk FD, and pooled control groups. RE was calculated as 2–Ct, where Ct = Ct mtDNA content in FD group – Ct mtDNA content in the control.
mRNA expression levels. Total RNA was extracted with an RNAzol Bee-RNA isolation kit (Tel-Test). Briefly, 1 µg of each sample was reverse-transcribed using Moloney murine leukemia virus RT in a reaction buffer containing random hexamer primers, deoxyribonucleoside triphosphates, and the ribonuclease inhibitor RNasin (Clontech Laboratories). Gene transcripts were amplified using gene-specific primers (see Supplemental Table 1) and ß-actin was used to control for variation in efficiency of RNA extraction and reverse transcription. The cycling conditions included an initial phase of 2 min at 50°C, 10 min at 95°C, then 40 cycles of 10 s at 95°C, 0.5 min at 60°C, and 10 s at 72°C. Amplified cDNA was quantified using the QuantiTect SYBR Green PCR kit in a LightCycler (Roche Diagnostics).
Analyses of 8-hydroxydeoxyguanosine levels. Total DNA was extracted and the 8-hydroxydeoxyguanosine (8-OHdG) levels were measured according to the method of Shigenaga et al. (41). An aliquot of 25 µg DNA was dissolved in 20 mmol/L sodium acetate/1 mmol/L deferoxamine mesylate, heated to 98°C for 10 min, and then cooled rapidly on ice. The DNA solution was treated with 20 units of DNase I in 25 µL of 0.1 mol/L MgCl2 and incubated at 37°C for 30 min. DNAs were hydrolyzed to nucleotides by further incubation with nuclease P1, sodium acetate (0.25 mol/L, pH 5.4), and 0.1 mol/L zinc sulfate at 37°C for 60 min. The resultant mixture was hydrolyzed to nucleosides by incubation with calf intestine alkaline phosphatase for 30 min at 37°C. A reverse-phase C18 HPLC column (4.6 mm x 150 cm, with a particle size of 5 µm; Waters Associates) was used to separate the nucleosides. The mobile phase of 50 mmol/L KH2PO4 buffer (pH 5.5) and 20% methanol was delivered at a flow rate of 0.8 mL/min. Pure deoxyguanosine (dG; Sigma Chemical) and 8-OHdG (Cayman Chemical) were used as standards. An HP 1100 series UV detector (Shimadzu SPD-M 10A) set at 254 nm was used to measure dG and an ECD detector (Shodex EC-1), with the electrode potential adjusted to 0.594 V, was used to detect 8-OHdG. The results are expressed as the ratio of 8-OHdG:dG.
Statistical methods.
Data are presented as means ± SD. Student's t test was used to compare animal growth and messenger RNA (mRNA) expression between the FD and the control group at 2 and 4 wk, and variables between each control group. Because plasma folate (250.5 ± 14.2 vs. 260.3 ± 20.8 nmol/L), RBC folate (585 ± 121 vs. 646 ± 187 nmol/L), Hcy levels (19.7 ± 3.0 vs. 20.4 ± 4.7 µmol/L), 8-OHdG ratio (0.92 ± 0.42 vs. 1.03 ± 0.29), and liver mtDNA deletions (Ct = 3.67 ± 0.22 vs. 4.34 ± 0.59) in 2-wk and 4-wk control groups did not differ significantly, the 2 control groups were pooled. The effects of dietary folate deprivation with the pooled controls were analyzed by 1-way ANOVA and Duncan's multiple range test using the General Linear Model of SAS Institute. Unequal variances were first transformed using a logarithmic function. Pearson's correlation coefficient was used to measure the association among mtDNA damage, mtDNA biogenesis, and folate depletion variables in the controls and FD groups separately. Differences were considered significant at P < 0.05.
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Results |
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Changes in accumulation of mtDNA4834 deletion among FD tissues. By real-time PCR analysis, we found spontaneous or background mtDNA common deletions in tissues of the controls (Table 2). Compared with the controls, 2-wk folate deprivation significantly increased the frequency of the mtDNA4834 deletion in pancreas, heart, and liver. Further mtDNA deletions in those tissues were accumulated after 4-wk folate deprivation, with RE values of 2.3–5.2 (P < 0.05). mtDNA deletions did not change significantly in brain and kidney until 4 wk. Pancreas accumulated the mtDNA4834 deletion at the greatest rate among the measured tissues.
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Discussion |
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TOPABSTRACTIntroductionMaterials and MethodsResultsDiscussionLITERATURE CITED |
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The exact cause of the increased mtDNA4834 deletion levels in FD tissues remains unknown. Several lines of evidence from this study indicate that mtDNA4834 deletion level in FD tissues is mt folate-dependent. First, the FD tissue with the most depleted mt folate content harbored the highest increase in mtDNA4834 deletions. Also, decreased mt folate levels were strongly correlated with increased frequency of mtDNA deletion among all measured tissues. This relationship was not found in tissues of the control rats. Reduced mt folate content resulted in mt oxidative injuries, including compromised antioxidant enzymatic activities, elevated protein carbonyl contents, and ROS production, as we have previously demonstrated in primary hepatocytes of FD rats after 4 wk (29). The elicited oxidative stress inside FD mitochondria could be attributable to mtDNA large deletions. This speculation is supported by the finding that sublethal redox stress such ROS production could lead to loss of mtDNA molecules or to deletions or a loss of function by base modification in the mtDNA (15,16). Similarly, tissues with high antioxidant capability in mitochondria essentially have a low level of mtDNA deletions (47). It has also been hypothesized that mtDNA large deletions could arise by slip-replication, in which replication is stalled by oxidative damage, allowing slipped mispairing between repeated sequences or by erroneous RNA splicing (48). Indeed, we found that the increased accumulation of the mtDNA4834 deletion in FD rat liver significantly correlated with increased 8-OHdG content, a biomarker of DNA oxidative injuries. Thus, our work and others' suggest that reduced mt folate content and elicited oxidative stress inside mitochondria contribute to the increased frequency of the mtDNA4834 deletion in FD tissues.
Our data demonstrate for the first time, to our knowledge, that mt biogenesis was enhanced during folate deficiency. In FD rat livers after 4 wk, the adaptive response occurred via the genetic regulation of factors for mt biogenesis (Fig. 2). NRF-1 is an oxidative-responsive transcriptional factor that binds to the promoter region of genes for proteins such as TFAM and cyt c (20,21) to trigger mt biogenesis. In 4-wk FD rat liver with elevated oxidative DNA damage, expression of NRF-1 mRNA was significantly induced. The NRF-1 induction coincided with upregulation of NRF-1 target genes, Cyt c, and TFAM (Fig. 2). In parallel, expression of mRNA for nuclear-encoded and mt-encoded respiratory subunits (CcOX IV, V, III, Cyt c) increased. This finding was consistent with reports that showed that oxidative stress caused via the stimulation of LPS (23), reactive oxygen (15), and elevated Hcy (49) could induce NRF and TFAM mRNA expression. It is likely that the enhanced mtDNA biogenesis at the transcriptional level in FD tissues was the consequence of mt oxidative injuries as a compensation to rescue the impaired tissues, a scenario previously reported in LPS-treated rat liver (23) and in senescent cells or aging tissues (24–28). Several studies have shown in cardiac myocytes (15), human fibroblasts (27), leukocytes (28), and mouse brains (24) that the elevated oxidative stress was related to increased mtDNA damage and mtDNA biogenesis. On the other hand, triggered mt biogenesis may reciprocally facilitate propagation of mutated mtDNA already carried within the impaired tissues, particularly in growing cells (16,17). Certain mutant mtDNA molecules exhibit a replicative advantage and mitochondria that undergo replication appear to acquire DNA damage more readily (18). The enhanced mt biogenesis in 4-wk FD liver of young rats that are still growing and undergoing cell division may explain in part, if not all, the accumulation of mtDNA4834 deletion in FD tissues.
Although machinery of mt biogenesis for mtDNA proliferation and transcription was triggered during the 4-wk FD period, mtDNA content of FD tissues did not increase accordingly. This decline of mtDNA content significantly associated with decreased mt folate levels of FD tissues in general and with elevated 8-OHdG levels of FD liver in particular. The data suggested that elicited oxidative stress (29) and DNA oxidative injuries (this study) as a result of mt folate deficit may otherwise interfere with mtDNA replication. The mtDNA molecule contains the noncoding region of a unique D-loop that contains the leading strand origin of replication and the promoters of transcription (50). Mutations in this region might affect the rate of DNA replication by modifying the binding affinity of important trans-acting factors. The D-loop region is a "hot spot" for somatic mutations in human cancers (51). Frequent mutations in the D-loop region and decreased mtDNA contents were found in oxidative stress-associated human liver diseases such as hepatitis and hepatocellular carcinoma (52,53). Whether mt folate deficit and elevated DNA oxidative injuries may result in increased mutations of D-loop leading to reduced mtDNA contents warrants more studies.
In summary, this study demonstrates that the mtDNA4834 deletion is increasingly accumulated in various tissues of young rats during dietary folate deprivation. The corollary findings of elevated oxidative DNA injuries and enhanced mt biogenesis associated with this mtDNA large deletion apparently represent novel phenomena observed in FD tissues of young animals. mt Folate level was a significant predictor of increased accumulation of the mtDNA common deletion and decreased mtDNA content. The folate-related mt biogenesis depicts a molecular mechanism to compensate mtDNA impairment in folate-deficient tissues.
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ACKNOWLEDGMENTS |
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FOOTNOTES |
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2 Author disclosures: Y.-F. Chou, C.-C. Yu, and R.-F. S. Huang, no conflicts of interest.
3 Supplemental Table 1 and Supplemental Figures 1 and 2 are available with the online posting of this paper at jn.nutrition.org.
4 Abbreviations used: CcOX, cytochrome c oxidase; Ct, threshold cycle number; Cyt c, cytochrome c; dG, deoxyguanosine; D-loop, displacement loop; 8-OHdG, 8-hydroxydeoxyguanosine; FD, folate deficient; Hcy, homocysteine; mRNA, messenger RNA; mt, mitochondrial; mtDNA4834 deletion, 4834-bp large deletion in mtDNA; NRF, nuclear respiratory factor; RE, relative expression; ROS, reactive oxygen species; TFAM, mitochondrial transcriptional factor A.
Manuscript received 13 June 2007. Initial review completed 13 June 2007. Revision accepted 3 July 2007.
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