Intracellular NAD+ modulates DNA repair capacity

NAD+ is a substrate for DNA repair proteins such as PARP1, PARP2 and PARP3 as well as enzymes that can influence DNA repair capacity such as SIRT1 and SIRT615,25,61. As a substrate for these enzymes, NAD+ availability may directly affect DNA repair pathway function as well as modulate chromatin structure to further influence DNA repair capacity. Therefore, we asked if changes in intracellular NAD+ content affects the levels of genotoxin-induced DNA damage as well as DNA repair capacity.

We therefore used the next-generation single cell gel electrophoresis assay, the CometChip62, to evaluate cellular genomic DNA damage and repair in human cells. As we have reported, this newly developed CometChip system resolves many of the limitations of the traditional comet assay such as reproducibility and variability62. The measurement of DNA damage by the CometChip is based on the fact that supercoiled (undamaged) DNA will not migrate in an agarose matrix under an electric field in contrast to fragmented (damaged) DNA which migrates to the positive electrode. The extent and the amount of migrated DNA (% Tail DNA) correlate with the amount of accumulated breaks in the DNA63. In these experiments, we used the alkaline CometChip assay, which results in an increase in % Tail DNA for many different types of DNA damage such as alkaline-sensitive base lesions, abasic sites, single-strand and double-strand DNA breaks64. As shown in Fig. 1A, the methylating agent methylnitronitrosoguanidine (MNNG), an S N 1 type alkylating agent, the methylating agent methyl methanesulfonate (MMS), an S N 2 type alkylating agent, and etoposide, a topoisomerase II inhibitor, induced the highest amount of damage while bleomycin and potassium bromate (KBrO 3 ) treatment resulted in a more modest level of DNA damage. The other three compounds (camptothecin, cisplatin (CDDP) and rapamycin) induced a minimal increase in measurable DNA damage over background (approximately 5% Tail DNA) at the indicated doses and time of exposure (1 hr). For the follow-up repair capacity experiments, we selected two agents that generate different classes of DNA damage: etoposide and MMS.

Figure 1 Effect of NAD+ depletion on DNA damage and repair in MCF-7 cells as determined using the CometChip assay62. (A) Level of DNA damage in MCF-7 cells exposed (1 hr) to different concentrations of various DNA damaging agents. (B) Level of intracellular NAD+ in MCF-7 cells 24 hrs after treatment with FK866 (30 nM) as compared to control. Statistical analysis was conducted via an unpaired t-test (****p < 0.001). (C) Level of DNA damage and repair capacity in MCF-7 cells exposed for one hr to etoposide (10 μM) in the presence or absence of FK866 (FK; treatment with FK866 results in a 70–90% decrease in total cellular NAD+ levels16). (D) Level of DNA damage and DNA repair capacity in MCF-7 cells exposed for one hr to different MMS concentrations (0, 0.125, 0.25, 0.5 and 1 mM) in the presence or absence of FK866 (FK; treatment with FK866 results in a 70–90% decrease in total cellular NAD+ levels16). Data is expressed as % Tail DNA, with standard deviation. Plots are the average of 8–12 wells from two-three independent CometChips; >1000 comets per bar. Statistical analysis was performed using GraphPad Prism 7 and two-way ANOVA followed by post-hoc with Tukey’s multiple comparison test (**p < 0.0029, ***<0.0008, ****<0.0001). Full size image

To assess the effect of alterations in the cellular level of NAD+ on DNA damage accumulation and DNA repair capacity, we used the NAMPT inhibitor FK866 to deplete the intracellular NAD+ pool16. The FK866-treatment protocol (24 hrs; 30 nM or 60 nM) results in an 80–90% reduction in the intracellular level of NAD+, as we reported16 (Fig. 1B). Following FK866 treatment, cells were exposed to damaging agents (1 hr) followed by a repair course whereby the cells were washed to remove the DNA damaging agents and the cells were then incubated in fresh media (at 37 °C) to promote repair of the induced DNA lesions. MCF-7 cells exhibit a very low level of endogenous DNA damage (Fig. 1C, D), which was not significantly affected by NAD+ depletion (FK866 treatment). Treatment of MCF-7 cells with etoposide (10 µM) induced approximately 30% Tail DNA, consistent with our previous CometChip analyses62. A repair course of 15, 30 and 60 minutes was sufficient for greater than 75% repair, as we have shown for TK6 and Jurkat cells62. MCF-7 cells presented with time-dependent repair of the etoposide-induced DNA damage in both the FK866-treated and untreated cells, indicating that NAD+ status did not affect the amount of induced damage nor the DNA repair capacity in response to etoposide treatment (Figs. 1C and S1A). Correspondingly, we find that PARP1-KO cells show no difference, as compared to control cells, in the rate of repair of DNA damage induced by etoposide (J. Li and R.W.Sobol, Manuscript in Prep).

MMS treated MCF-7 cells showed a dose dependent accumulation of DNA damage but in contrast to the etoposide-induced damage, MMS-induced DNA damage required a longer time (3 hrs) for repair of the induced DNA lesions (Figs. 1D and S1B). At low MMS concentrations (0.125 and 0.25 mM), MCF-7 cells demonstrated a greater level of DNA damage in the absence of NAD+ than in the presence of NAD+; however, the level of MMS-induced DNA damage was the same at higher MMS concentrations regardless of the NAD+ content (Fig. 1D). As might be expected by the requirement for PARP1 in BER-mediated repair of MMS-induced DNA damage65, NAD+ depletion significantly suppressed the repair of the DNA damage, with as much as a 40% lower rate of repair when NAD+ was depleted (Figs. 1D and S1B).

Cancer cells express elevated mRNA levels for genes involved in the NAD+ salvage pathway

We next evaluated the genes required for cells to maintain sufficient NAD+ levels that would be critical to support metabolism, as we have shown16, as well as maintain genome stability (Fig. 1). We performed a comparative analysis of mRNA expression of the genes involved in NAD+ metabolism in MCF-7 cells, plus an additional eight cancer cell lines, including LN428, T98G, A375, G09607, HCT116, MDA-MB-231, NCI.H460 and U87MG (Table 1). We divided the genes into three categories; (i) enzymes involved in de novo NAD+ biosynthesis, (ii) enzymes involved in the salvage pathway and (iii) NAD+ consuming enzymes (Table 1). All the cell lines strongly expressed most genes involved in the salvage pathway as compared to those involved in the de novo pathway, most prominently seen for NAMPT and purine nucleoside phosphorylase (PNP). In addition, kynureninase (KYNU) and tryptophan-2,3-dioxygenase (TDO2) of the de novo pathway were also elevated. The elevated expression of NAMPT and PNP is consistent with multiple reports and clinical studies showing elevated levels of these enzymes in several cancers36,37,44,66. NNMT, the nicotinamide N-methyltransferase reported to be elevated in many cancers67 and cancer-associated fibroblasts68, is also shown elevated in most of the cell lines evaluated (Table 1). Further, all three categories of enzymes are shown to be amplified in breast cancer tumors69,70, as shown in Supplementary Fig. S2A–C.

Table 1 –mRNA expression of genes involved in NAD+ metabolism. Full size table

Regarding the group of NAD+ consuming enzymes, the majority of genes that displayed high mRNA expression were PARP-family and sirtuin-family member proteins. However, the gene with the most varied expression was the ectoenzyme CD73 (NT5E)71. Given that the role of PARPs and sirtuins is well documented in a variety of cancers14, we decided to focus on the CD73 ectoenzyme. CD73, together with CD38 (gene = CD38, an NADase/glycohydrolase/ADP-ribosyl cyclase) and CD157 (gene = BST1; an NADase and ADP-ribosyl cyclase), are surface enzymes reported to consume NAD+ as a substrate for their enzymatic activity10,72,73. While the majority of cell lines expressed low levels of CD38 (8 out of 9) and CD157 (7 out of 10), most of the cells analyzed in this study expressed high levels of CD73 mRNA, with the exception of T98G cells. Of the cell lines under study herein, CD38 expression was not detected at the mRNA or protein level (Supplementary Fig. S2D, E). In the present study, we focused primarily on the role of CD73 in NAD+ metabolism, which could impact intracellular NAD+ content and therefore genomic stability of cancer cells. To this end, four cell lines were selected for our investigation: two glioma cell lines (LN428 and T98G) and two breast cancer cell lines (MDA-MB-231 and MCF-7). These cell lines express varying levels of CD73, representing, as such, the variances observed in the cell lines examined (Table 1).

Analysis of NMN, NAD+ and NADH catabolic activity by recombinant, human CD73

The H. influenzae enzyme HiNadN, considered to be the ortholog to the human CD73 enzyme, was previously shown to hydrolyze NAD+ and NMN when evaluated by HPLC and colorimetric assays54,74,75,76, suggesting that the human enzyme (CD73) may also catabolize these extracellular metabolites, processing NAD+ to AMP and NMN and then hydrolyzing both AMP and NMN to adenosine and nicotinamide riboside (NR), respectively (Fig. 2A). Commercially available preparations of human CD73 were all shown to be inactive (Supplementary Fig. S3A–D). NADH, just like NAD+, is likely to be present in the microenvironment of a tumor and potentially be hydrolyzed to the reduced form of NR, NRH, a known precursor to NAD+77. Therefore, to directly evaluate the capacity of the human CD73 enzyme to catabolize NMN, NAD+ or NADH, the soluble form of recombinant CD73 (residues 27–549 including a His-tag at the C-terminus) was produced in Sf9 insect cells and purified as previously described78. The purified rCD73 is highly active against AMP yet inactive against NAD+ or NADH and very poorly active against NMN (Fig. 2B). Similarly, we show that the H. influenzae enzyme HiNadN is able to effectively catabolize NAD+ yet does not process NMN when measured by 31P NMR (Supplementary Fig. S4). Our in vitro biochemical analysis of the recombinant purified enzymes supports a role for the bacterial enzyme HiNadN in the processing of NAD+ with no evidence for NAD+ hydrolysis catalyzed by the recombinant, purified human CD73 enzyme. Further, we find that the recombinant, purified human CD73 enzyme processes NMN extremely poorly (Fig. 2B).

Figure 2 Human, recombinant CD73 activity analysis for the hydrolysis of NMN, NAD+ and NADH. (A) Scheme showing the structure of NAD+ and conversion to AMP (bottom) and then to adenosine with loss of inorganic phosphate and to NMN (top) and then to NR. (B) Recombinant CD73 (rCD73; soluble form, residues 27–549 including a His-tag at the C-terminus) was purified and evaluated for enzymatic activity measured in the presence of the natural substrate, AMP, or in the presence of NMN. There was no detectable activity against NAD+ or NADH (not shown). Full size image

Impact of differential CD73 expression on the intracellular NAD+ content in human cancer cell lines

Given that NAD+ is not hydrolyzed by recombinant human CD73 and is very poorly active against NMN (Fig. 2B), this raised the question as to whether CD73 plays any role at all in metabolizing extracellular NAD+ and therefore in regulating the intracellular level of NAD+ in human cells. We confirmed the expression level of CD73 mRNA and protein in four cancer cell lines by qRT-PCR and immunoblot analysis, respectively. The breast cancer cell line MDA-MB-231 displayed extremely high levels of CD73 at both the mRNA and protein level (Fig. 3A, B). The glioblastoma cell line LN428 also displayed high levels of CD73 expression but significantly lower compared to the MDA-MB-231 cells (Fig. 3A, B). The breast cancer cell line MCF-7 expressed CD73 mRNA and protein yet the glioblastoma cell line T98G exhibited very low to almost undetectable levels of CD73 (Fig. 3A, B). Among the human cancer cell lines we have evaluated, the expression of CD73 is highly varied as compared to other proteins involved in NAD+ metabolism, such as NRK1 (Figs. 3B and S2F).

Figure 3 Comparative analysis of CD73 expression in human cancer cell lines and its effect on NAD+ biosynthesis. (A) Measurement of mRNA expression for the CD73/NT5E gene in cancer cell lines, as determined by qRT-PCR analysis, normalized to the expression of human β-actin mRNA via the ∆∆CT method. (B) Immunoblot analysis of the expression of CD73 in nine breast cancer cell lines, including the MCF-7 and MDA-MB-231 cells used herein (top panel) and two glioma cell lines, including the LN428 and T98G cells used herein (bottom panel). PCNA was used as a loading control for the top panel and actin was used as a loading control for the bottom panel. See Supplement Fig. S2F for the expression of NRK1 in the breast cancer cell lines analyzed from the same cell lysates. (C) Total intracellular NAD levels (NAD+ and NADH) in each of the four cancer cell lines cultured in the presence of NAD+, NMN or NR (100 μM) for 24 hrs. Statistical analysis was performed using GraphPad Prism 7 and two-way ANOVA followed by post-hoc test with Tukey’s correction (ns = not significant, *p = 0.0419, **p = 0.0032). Full size image

Therefore, we first examined whether CD73 is indeed responsible for hydrolyzing NAD+, as shown in the scheme in Fig. 2A, and if the variations in expression impact the intracellular level of NAD+. To this end, we measured the total NAD pool (NAD+ plus NADH) in all four cell lines following addition (100 μM, 24 hrs) of nicotinamide adenine dinucleotide (NAD+), nicotinamide mononucleotide (NMN) or nicotinamide riboside (NR) to the culture media. The basal (control) NAD+ and NADH content (as per the colorimetric analysis) was similar among all the cell lines (Fig. 3C), although there was slight variation. LN428 cells had the lowest levels of NAD+ (0.80 µM) and NADH (0.26 µM) while MCF-7 cells had the highest levels of NAD+ (1.51 µM) and NADH (0.35 µM). While the addition of all of the precursors to the culture media increased NAD+ levels in all cell lines (Fig. 3C), only NR affected NADH levels statistically. Interestingly, the increase in NADH following NR supplementation in the 4 cell lines (LN428, T98G, MDA-MB-231 and MCF-7) does not seem to correlate with the corresponding mRNA levels of PNP (Supplementary Fig. S2G). Importantly, despite highly variable levels of CD73 among the four cell lines, no correlation between CD73 expression level, NAD+ or NMN supplementation and intracellular NAD+ content was observed. Interestingly, MCF-7 cells, which express a lower level of CD73 as compared to the MDA-MB-231 cells (Table 1, Fig. 3A,B) displayed the highest basal intracellular NAD+ level and in the presence of NAD+ and the vitamin B 3 precursors, NMN or NR. This is consistent with the report that MCF-7 is an “NAD keeper” as measured by flux analysis79.

NAMPT inhibition accelerates uptake of NAD+ precursors while CD73 inhibition does not affect NAD+ biosynthesis

The cancer cell lines under study did not show a correlation between the level of CD73 expression and the intracellular NAD+ content when exposed to NAD+ precursors in the absence of stress. We therefore next introduced NAMPT inhibitor treatment to disturb cellular NAD+ homeostasis. Treatment with FK866, a potent NAMPT inhibitor41, is expected to deplete intracellular NAD+, as shown in Fig. 1B, and accelerate an uptake of NAD+ precursors: NAM, NR and NMN.

In the absence of FK866, supplementation of NAM, NR or NMN induced an increase in NAD+ levels in only a few of the cell lines. Supplementation of NAM (24 hrs, 100 µM) to the LN428 cell culture media resulted in a modest increase in the basal intracellular level of NAD+ with no significant change in the other three cell lines (Fig. 4A). Further, supplementation of NR (24 hrs, 100 µM) caused a significant increase in intracellular NAD+ content in LN428 and MCF-7 cell lines with no significant impact on the T98G and MDA-MB-231 cell lines (Fig. 4B), again consistent with a reduced turn-over of the NAD+ pool by NAD+-consuming enzymes in MCF-7 cells compared to MDA-MB-231 cells79. Finally, supplementation of NMN (24 hrs, 100 µM) to the culture media only increased the basal intracellular level of NAD+ in the MCF-7 cell line, with no significant alteration in the other three cell lines (Fig. 4C). As expected, when cells were supplemented with NAM in the presence of FK866, NAM was not able to rescue NAD+ levels since its metabolism entirely depends on NAMPT activity. In contrast, NR supplementation resulted in the greatest increase in NAD+ levels in the presence of FK866. Although there was no significant increase in T98G cells, LN428, MDA-MB-231 and MCF-7 cells all reported a strong increase in NAD+ levels although never higher than the cells supplemented with NR in the absence of FK866 (Fig. 4B). However, NR metabolism does not depend on CD73 activity. The addition of NMN was also able to rescue NAD+ levels but not as efficiently as NR in FK866 treated cells and reached significance only in the MCF-7 cell line. Notably, even when cells are stressed by NAD+ deprivation, the high level of CD73 expression, as seen in the MDA-MB-231 cells, did not affect the uptake of precursors or the increase in NAD+ biosynthesis (Fig. 4C). In summary, there is a lack of any significant difference in NAD+ content between cells with undetectable levels of CD73 (such as T98G), cells with CD73 expression (MCF-7) or cells with high or very high levels of CD73 (such as LN428 and MDA-MB-231, respectively). We also tested if inhibiting CD73, using the competitive inhibitor adenosine 5′-(α,β-methylene) diphosphate (APCP), would impact intracellular NAD+ content. In-line with our cell line analyses (Fig. 4), APCP did not cause major changes in NAD+ levels and inhibition of CD73 exerted no effect on cellular NAD+ content in combination with FK866 treatment (Supplementary Fig. S5A–D).

Figure 4 Effect of NAMPT inhibition on intracellular NAD+ content in cancer cell lines with different basal levels of CD73 protein. (A) Intracellular NAD+ levels in cells treated with FK866 (30 nM) and/or nicotinamide (NAM) (100 μM), as compared to untreated controls. (B) Intracellular NAD+ levels in cells treated with FK866 (30 nM) and/or nicotinamide riboside (NR) (100 μM), as compared to untreated controls. (C) Intracellular NAD+ levels in cells treated with FK866 (30 nM) and/or nicotinamide mononucleotide (NMN) (100 μM), as compared to untreated controls. (D) Top panel : Representative immunoblot analysis of NRK1 expression to correlate the expression of NRK1 in four cancer cell lines (see Supplement Fig. 3E for additional immunoblot figures). Bottom panel : Densitometry analysis of 3 independent immunoblots, performed using Image Lab software. Statistical analysis was performed using GraphPad Prism 7 - one or two-way ANOVA followed by post-hoc test with Tukey’s correction was used (ns = not significant, *p = 0.0176, **p = 0.0018, ***p = 0.0009 or ****p < 0.0001). Full size image

We therefore reasoned it was then critical to determine which enzyme in the NAD+ biosynthesis pathway could be responsible for such a strong effect on NAD+ biosynthesis. Given that NRK1 and NRK2 are the kinases that phosphorylate NR to produce NMN50, they represented the likely candidates that may influence NAD+ biosynthesis in cells supplemented with NR. Not surprisingly, a qRT-PCR assessment revealed no detectable levels of NRK2 mRNA expression in MCF-7 cells (data not shown), since NRK2 is exclusively expressed in muscle tissue80. Similarly, NRK1 protein levels in the four tested cell lines, as assessed by immunoblot analysis, did not correlate with the measured NAD+ levels (Fig. 4D). Overall, these results did not explain the mechanism by which MCF-7 cells were the most effective at utilizing NAD+ precursors.

Kinetic analysis of cell culture supernatants reveals the products of CD73 enzymatic activity

Direct enzymatic analysis of the recombinant human CD73 enzyme (rCD73) revealed that purified rCD73 is highly active against AMP yet inactive against NAD+ or NADH and very poorly active against NMN, if at all (Fig. 2B). Further, we found that NMR analysis of the bacterial orthologue of CD73, HiNadN, revealed that HiNadN can hydrolyze NAD+ to NMN but cannot metabolize NMN to NR. Therefore, to enhance our analysis of human cells expressing CD73, we performed NMR analysis of cell culture supernatants to determine if the supplemented NAD+ precursors were subjected to CD73-mediated enzymatic processing. We focused on the MCF-7 cell line since, in the presence of NAD+ precursors, only MCF-7 cells were able to overcome CD73 and NAMPT inhibition to restore NAD+ levels (Supplementary Fig. S5A–E). Therefore, we treated MCF-7 cells for 24 hrs with NAD+ supplements and then compared the content of the cell culture supernatants to media containing precursors that had never been exposed to cells. To our surprise, incubation in media alone induced hydrolysis of the NAD+ precursors to the same degree as in the presence of MCF-7 cells. This suggested that the media or one of the media components might impact the stability of the NAD+ precursors.

Upon 24 hrs incubation of MCF-7 cells with NAD+, NR or NMN in the presence or absence of FK866 (FK), we only detected NR and NAM in the supernatants from all of the tested samples (Fig. 5A). The ratio between NR and NAM was different when compared to the treatments with NAD+, NR or NMN but very similar when comparing ‘media only’ with ‘media exposed to cells’. Additionally, the presence of FK866 did not impact the composition or distribution of NR and NAM in the tested samples. At the same time, the presence of NR in the cell supernatants could indicate that hydrolysis of NMN to NR took place. The presence of NAM could imply the enzymatic activity of CD157 or CD38. However, MCF-7 cells do not express these consuming enzymes, as shown in Table 1 and we found no detectable levels of CD38 mRNA or protein (Supplementary Fig. S2D,E).

Figure 5 Assessment of the time dependent changes of extracellular NAD+ and NAD+ metabolite composition and intracellular NAD+ levels following metabolite supplementation and/or NAMPT inhibition. (A) NMR analysis of the composition of NAD+/NAD+-metabolites in MCF-7 cell supernatants from cells exposed to NAD+ for 24 hrs. (B) Measurements of intracellular NAD+ content in MCF-7 cells exposed to NAD supplements (NMN and NR) and/or the NAMPT inhibitor, FK866, at different time points after supplement addition. Statistical analysis was performed using GraphPad Prism 7 - one-way ANOVA followed by post-hoc test with Tukey’s correction was used. (ns = not significant, ***p = 0.0004 or ****p < 0.0001) (C) Immunoblot analysis to evaluate changes in CD73 expression impacted by the treatment of MCF-7 cells with FK866 (30 nM), NMN (100 μM) or NAD+ (100 μM) for times ranging from 0.5 to 24 hrs, as compared to the untreated control, as indicated. Full size image

Next, we measured the intracellular NAD+ content in MCF-7 cells at different time points in order to determine how fast the cells would utilize the available precursors for NAD+ biosynthesis. We measured NAD+ levels in MCF-7 cells exposed to NMN and NR as early as 30 min, followed by 1, 6, 16 and 24 hrs. In addition, we also treated cells with FK866 (FK) for 24 hrs, inducing NAD+ deprivation to intensify the uptake of NAD+ precursors. At early time points (30 min and 1 hr), the NAD+ content in cells treated with the supplements was similar to the untreated MCF-7 cells. In-line with our earlier analysis, 24-hr treatment with FK866 depleted ~75% of the intracellular NAD+ and short-term supplementation with NMN or NR did not impact NAD+ levels (Fig. 5B).

At 6 hrs post treatment, we observed an increase in NAD+ content due to NR supplementation. Finally, after 16 and 24 hrs of treatment, we observed a significant increase in NAD+ content upon both NR and NMN supplementation. The presence of FK866 augmented the differences between the conditions, showing a strong effect of NAD+ deprivation on the uptake of NAD+ precursors (Fig. 5B). Importantly, at 16 and 24 hrs, MCF-7 cells showed an increase in NAD+ from NMN uptake. We also tested if NAD+ depletion and NAD+ supplementation affect the status of CD73 at the protein level. To do so, we exposed MCF-7 cells to NMN and NAD+, rather than NR, since their metabolism should depend on CD73 enzymatic activity. We also tested the effect of the FK866 inhibitor on CD73 protein levels in MCF-7 cells. The presence of the NAD+ precursors, as well as FK866 treatment, did not affect the level of CD73 protein at any time point (Fig. 5C). However, it is noted that these experiments excluded the potential regulation of CD73 activity by the level of intracellular NAD+ or the presence of extracellular NAD+ precursors.

MCF-7/CD73 knockout cells are efficient in hydrolyzing NAD+ to NMN and NR

To more effectively probe the requirement for CD73 in NAD+-precursor uptake, we developed an isogenic system by creating a human MCF-7/CD73-KO cell line using the CRISPR/Cas9 editing system81,82. The absence of CD73 protein was confirmed by immunoblot analysis where we compared MCF-7/CD73-KO cells to the isogenic MCF-7/Cas9 cells (Fig. 6A, left panel). This isogenic cell system allows us to exclude the impact of CD73 on NAD+ metabolism and helps to address the data showing hydrolysis of NAD+, NR and NMN in media only as shown in Fig. 5A.

Figure 6 Effect of CD73 knockout on the uptake of NAD+ precursors and on NAD+ biosynthesis. (A) Left panel : Immunoblot analysis of CD73 expression in MCF-7/Cas9 and MCF-7/CD73-KO cells; actin is shown as a loading control. Right panel : Comparative analysis of intracellular NAD+ levels in both cell lines exposed to different NAD+ precursors (100 μM) in the absence or presence of the NAMPT inhibitor, FK866 (30 nM). (B) NMR analysis to define the distribution of NAD+ and NAD+ metabolites (100 μM) in cell culture media 6 hrs after supplementation of NMN or NAD+ to cell-free serum-free media (SFM), cell-free media supplemented with fetal bovine serum (FBS) or when added to SFM or FBS in the presence of MCF-7/Cas9 or MCF-7/CD73-KO cells in the presence or absence of FK866 (30 nM). (C) NMR analysis to define the distribution of NAD+ and NAD+ metabolites in cell culture media 24 hrs after supplementation of NMN or NAD+ to MCF-7/Cas9 cells in the presence or absence of FK866. Cells were exposed to media supplemented with fetal bovine serum (FBS) or heat inactivated fetal bovine serum (FBS-HI) and compared to NMN and NAD+, which have never been exposed to cells. Full size image

We then utilized our control (MCF-7/Cas9) and isogenic CD73 deficient (MCF-7/CD73-KO) human cells to determine if the loss of CD73 impacts the uptake of NAD+ or NAD+-precursors sufficiently to modulate the level of intracellular NAD+. As in the previous analyses, we also exposed cells to the NAMPT inhibitor (FK866; 24 hrs, 30 nM) to increase the uptake of the NAD+ precursors. MCF-7/Cas9 and MCF-7/CD73-KO cells demonstrated a very similar increase in NAD+ content when exposed to all four precursors, with NR supplementation driving the largest increase in NAD+ levels (as compared to control), regardless of FK866 treatment (Fig. 6A). For both cell lines, NAM supplementation in the presence of FK866 could not restore NAD+ (as expected) since NAM metabolism requires NAMPT enzymatic activity. When cells were supplemented with NR, which should not rely on CD73 for uptake, we observed similar levels of NAD+ in both the control (MCF-7/Cas9) and isogenic CD73 deficient (MCF-7/CD73-KO) cells. This may be predictable since there are known nucleoside transporters likely involved in transporting NR or NMN83,84. Finally, when we supplemented both cell lines with NAD+ or NMN, each suggested as substrates for CD7354, there was an equivalent level of intracellular NAD+, for both the MCF-7/CD73-KO and MCF-7/Cas9 cell lines. However, the difference was slightly more pronounced when cells were treated with FK866. Once NAMPT was inhibited (with FK866), NMN supplementation completely complemented the NAD+ levels in the CD73 deficient cells (MCF-7/CD73-KO) whereas NAD+ supplementation resulted in a slight, albeit non-significant, reduction in NAD+ complementation (Fig. 6A). Therefore, these studies would suggest that CD73 does not participate in NAD+ or NMN uptake and metabolism. Importantly, expression of the proposed NMN transporter (Slc12a8) was found to be very low in all the cell lines under study here, with the expression level in MCF-7 cells effectively below detection, as measured by qRT-PCR (Supplementary Fig. S5F).

Next, we performed NMR analysis of cell supernatants (media) collected from MCF-7/Cas9 and MCF-7/CD73-KO cells to evaluate the catabolism of the supplemented NAD+ and NAD+-precursors. The cell supernatants were collected after 24 hrs of incubation with vitamin B 3 precursors as well as the FK866 (FK) inhibitor. Overall, the distribution of precursors between the Cas9 (control) and CD73-KO cell lines showed high similarity. Specifically, supplementation with NR or NR+FK, showed ~50% of the NR being metabolized into NAM, while supplementation with NMN and NMN+FK gave about 40–50% NAM, with NR being the second largest component and a small portion of NMN present in all conditions, with the exception of MCF-7/Cas9 treated with NMN alone. These data suggest robust conversion of NMN to NR and further, NR to NAM, in both CD73 positive and negative cell lines. This confirms that CD73 enzymatic activity is not involved in NMN metabolism in MCF-7 cells since the CD73-KO cells showed the same composition and distribution of the precursors as the MCF-7/Cas9 cells. Finally, supernatants from cells incubated with NAD+ or NAD++FK did not show any non-metabolized NAD+ (in both cell lines). Instead, we observed NR and NAM as the major components, with a small amount of NMN present. Patterns for both cell lines looked comparable, although MCF-7/Cas9 demonstrated stronger enzymatic activity towards NMN and NAD+ when cells where incubated with precursors alone but not when they were co-exposed to the FK866 inhibitor (Supplementary Fig. S5E).

The detection of NAM and NR in the media supplemented with NAD+ but never exposed to the cells (Fig. 5A) as well as the contradictory results of complete metabolism of NAD+ precursors (Supplementary Fig. S5E) suggested that some of the NAD+ precursors were not as stable in media containing fetal bovine serum (FBS), as expected59,84. Therefore, we repeated the same experiment using NAD+ and NMN precursors when comparing cells cultured in full media (FBS) or in serum free media (SFM). In addition, we shortened the time of incubation from 24 to 6 hrs in order to increase the chances to detect non-metabolized precursors by NMR analysis. We discovered that the distribution of metabolites when we compared SFM and FBS media supplemented with NMN and NAD+ was very different. Both NMN and NAD+ were stable in SFM but in FBS supplemented media, we identified NAM, NR, NMN and NAD+, suggesting degradation or breakdown of the metabolites in FBS-supplemented media (Fig. 6B).

Further, when we compared supernatants from MCF-7/Cas9 cells and MCF-7/CD73-KO cells in SFM and in FBS respectively, the profiles of precursor distribution looked similar. On the other hand, the distribution of NAD+ precursor between SFM and FBS among the same cell line looked very different. Specifically, NMN represented 100% of the profiles in SFM while in the presence of FBS supplemented media, the same precursor (NMN) was hydrolyzed to NR and NAM, with about 30–50% non-metabolized NMN detected. Similarly, the profiles of NAD+ supplementation varied between SFM and FBS supplemented media to the point that we were not able to detect NAD+ in the supernatants from cells cultured in FBS supplemented media. However, we identified that NAD+ was hydrolyzed to NMN, NR and NAM. Only a very small proportion of NAD+ was detected in Cas9 cells when incubated with NAD+ in the presence of FK866 (Fig. 6B). In the last set of experiments, we asked if heat inactivation of FBS would improve the stability of these small molecules. Testing heat inactivated serum (FBS-HI) and non-heat inactivated serum (FBS) revealed the presence of degraded metabolites of NAD+ and NMN was much more pronounced in FBS compared to FBS-HI supplemented media. Specifically, FBS-HI did not impact NMN while in FBS, NMN was hydrolyzed to NR and NAM. For NAD+ supplementation, although we did not detect un-metabolized NAD+ in the supernatants from FBS-HI containing media, we were able to show NMN and NR as major components. For cells cultured in FBS, NAD+ was degraded to NAM and a smaller proportion of NR. In all the conditions tested, the presence of FK866 (FK) did not provide additional changes (Fig. 6C).

Impact of intracellular NAD+ content on base excision repair complex formation

As detailed above, NAD+ availability has a profound impact on genomic DNA damage levels and on DNA repair capacity in MCF-7 cells (Figs. 1 and S1). The most prominent DNA damage types that appear to be NAD+-dependent are those induced by alkylating (Fig. 1) and oxidizing agents, likely repaired by the base excision repair (BER) pathway65. Given the significant role that PARP1 plays in BER15,16, it is possible that suppressed NAD+ levels may negatively regulate the PARP1-dependent recruitment of the scaffold protein XRCC185 to sites of DNA damage to form the BER complex. Therefore, we next asked if changes in intracellular NAD+ would affect the recruitment of DNA repair proteins to the site of damage and if the expression level of CD73 correlates with the outcome. To test this possibility, we expressed XRCC1 as a fusion with the fluorescent protein mCherry (XRCC1-mCherry) in the MCF-7/Cas9 and MCF-7/CD73-KO cells (Supplementary Fig. S7A) and utilized laser micro-irradiation confocal microscopy to induce DNA damage and quantify recruitment of the fluorescently labeled XRCC1 protein to sites of DNA damage. To induce DNA damage, we subjected cells to micro-irradiation using a laser with a 355 nm wavelength, which has been used to induce base lesions as well as DNA single-strand breaks (SSBs)86, as described in the methods section. We then measured BER/SSB repair complex assembly and disassembly (XRCC1 recruitment and resolution) at sites of DNA damage by measuring the mean intensity of fluorescence within a DNA damage region of the nucleus as compared to the mean fluorescent signal quantified for the entire nucleus, as described by Holton at al87. Before inducing damage, cells were depleted of NAD+ by treatment with FK866 (30 nM) and/or exposed to NAD+ precursors (NAD+, NMN and NR; 100 μM) for 6 or 24 hrs. We then evaluated the mean intensity of fluorescence of MCF-7/Cas9 and MCF-7/CD73-KO cells (Supplementary Fig. 6) as well as compared the maximum recruitment of XRCC1-mCherry between the two cell lines (Fig. 7).

Figure 7 Impact of NAD+ depletion and NAD+ precursor supplementation on DNA repair complex formation. (A, B) Comparative measurements of maximal recruitment of XRCC1-mCherry protein to the site of laser-induced DNA damage: MCF-7/Cas9 vs MCF-7/CD73-KO cells exposed to NAD+ in the presence of serum-free media (SFM) or media supplemented with fetal bovine serum (FBS) or heat inactivated fetal bovine serum (FBS-HI) for 6 hrs (A) or 24 hrs (B). (C) Representative images of maximal recruitment of XRCC1-mCherry in MCF-7/Cas9 cells cultured in media+FBS, SFM or media+FBS-HI; respectively, with differing NAD+ status. Full size image

We observed the same trend for XRCC1 recruitment in both the MCF-7/Cas9 and MCF-7/CD73-KO cell lines: rapid recruitment with a peak time of approximately 60 seconds and 50% resolution at approximately 200 seconds. As expected, recruitment was blocked when PARP1 was inhibited (Supplementary Fig. S7B,C). Further, NAD+ depletion (FK866 treatment) had a strong negative impact on XRCC1-mCherry recruitment to the site of DNA damage. This is consistent with the requirement for PARP1 activation for XRCC1 recruitment but may also imply other NAD+-dependent factors which may be at play, such as deacetylation by one of the SIRT family proteins88,89,90. In the absence of FK866 treatment, NAD+ or NAD+-precursor supplementation of the cells had little if any impact on DNA repair complex assembly and disassembly (Fig. 7A,B). This is in-line with the minor increase in cellular NAD+ levels seen when cells are supplemented with NAD+ or the precursors (Fig. 3C).

Further, we find that the FK866-effect can be reversed by supplementing cells with NAD+ (Figs. 7 and S6) as well as the NAD+ precursors NMN and NR (data not shown). However, NAD+ supplementation only reversed the recruitment defect in the FBS-containing media but not in SFM or FBS-HI-containing media (Figs. 7 and S6). In the FBS supplemented media, control cells, NAD+ or NAD+/FK866 supplemented cells exhibited robust XRCC1 recruitment with a complete lack of XRCC1 recruitment when NAD+ was depleted from the cells (Fig. 7). Conversely, in SFM and FBS-HI-containing media, NAD+ depletion inhibited XRCC1 recruitment but this could not be rescued by NAD+ supplementation (Fig. 7). Representative images of XRCC1 recruitment in FBS, SFM and FBS-HI, respectively at the maximum peak of the recruitment, are shown in Fig. 7C. Interestingly, in-line with our previous data (Fig. 5B), we observed a difference between 6 and 24 hr incubation with the precursors. In particular, for FBS and FBS-HI, an additional 18 hrs in the presence of NAD+ improved the recruitment of XRCC1-mCherry to the site of laser damage in cells treated with both NAD+ and FK866 (Fig. 7A,B), indicative of the role of the media in generating precursors of intracellular NAD+.

Collectively, our findings suggest that CD73 does not play a role in NAD+ precursor uptake and biosynthesis and importantly demonstrated the significance of intracellular levels of NAD+ on DNA repair complex formation, as measured by XRCC1 recruitment to sites of genomic DNA damage and the impact on the repair of alkylation-induced DNA damage.