Imprecision and DNA Break Repair Biased towards Incompatible End Joining in Leukemia

Whole-genome sequencing revealed the presence of numerous clonal and subclonal somatic mutations as well as somatic rearrangements in cancer genomes (1–3). Upon cancer therapy, subclonal refractory cells harboring acquired novel chromosomal aberrations or gene mutations can expand and may thus contribute to therapy resistance and disease relapse (4–6). A major source for chromosomal rearrangements are DNA double-strand breaks (DSB). DSBs occur constantly during the lifetime of any cell and are normally repaired. However, aberrant repair of DSBs can lead to misjoining of distant DNA ends, generating deletions, inversions, or complex rearrangements of chromosomes (7, 8). DSBs are repaired by two major pathways, which are homologous recombination (HR) and nonhomologous end joining (NHEJ). HR utilizes the sister chromatide as template for repairing DSBs and is thus primarily confined to S–G₂–M phase of the cell cycle, whereas NHEJ simply rejoins two DNA ends and, hence, is the main repair pathway for G₁ or G₀ arrested cells (7). For NHEJ, a classical and an alternative pathway are described, termed c-NHEJ and a-NHEJ, respectively. Although c-NHEJ is dependent on recognition of DNA ends by XRCC5, XRCC6, and DNA-PKc and ligation by DNA Ligase 4 (LIG4)/XRCC4, a-NHEJ is carried out independently of LIG4 and can probably be executed by a diverse set of factors, including different DNA polymerases (δ, θ), DNA nucleases (ERCC1-XPF), and ligases (LIG1, LIG3/XRCC1; refs. 9–11). As a-NHEJ is frequently associated with short microhomologies at the site of DNA junctions, a-NHEJ is also termed microhomology-mediated end joining (MMEJ).

Chronic lymphocytic leukemia (CLL) is a B-cell malignancy characterized by the progressive accumulation of clonal B cells in peripheral blood and lymph nodes. Clinically, the mutation status of the B-cell receptor divides patients into subgroups with poor (CLL with unmutated BCR, CLL-UM) and more favorable prognosis (CLL with mutated BCR, CLL-Mut; refs. 12–14). Although many of the hematologic malignancies show typical entity-specific chromosomal abnormalities (15), there is no specific aberration defining CLL. Instead, in addition to more common chromosomal abnormalities like del(17p), del(11q), del(13q), and Tri12, conventional cytogenetics as well as novel whole-genome sequencing approaches revealed a complex set of different chromsomal aberrations in CLL, with CLL-UM samples showing a slight increase in the number of aberrations (3, 5, 6, 16–19).

In this study, we asked whether the frequent occurrence of chromosomal abnormalities is associated with an inherently aberrant DNA DSB repair in CLL. To test this, we analyzed DNA DSB repair in primary CLL cells compared with B cells from healthy controls by assessing in vivo repair of artificial DNA substrates carrying diverse DNA break structures using next-generation sequencing (NGS) of repair junctions. These results were corroborated by analyzing the induction of a large chromosomal deletion using TALEN (transcription activator-like effector nuclease)/Cas9 (clustered regularly interspaced short palindromic repeats associated protein 9) endonucleases (20). Our data show that DNA end joining is less precise and more promiscuous in CLL with a bias toward microhomologies as well as toward joining of incompatible DNA ends, likely due to increased expression levels of LIG1 and XRCC1.

Plasmids were generated by cloning synthesized inserts (MWG eurofins) into pAX plasmids (sequence shown in Supplementary Fig. S1). Plasmids were linearized using the respective restriction enzymes (Fermentas), gel purified (Qiagen), and transfected into 293 cells (genejuice, Novagen) and primary B and CLL cells using nucleofection (Amaxa, Lonza). Seventy-two hours posttransfection, cells were harvested and DNA was isolated (Qiagen). Repaired substrates were PCR amplified using high-fidelity Phusion polymerase (Biozym) and tagged 3AOX/5AOX primers (3AOX 5′-GCA AAT GGC ATT CTG ACA TCC-3′; 5AOX 5′-GAC TGG TTC CAA TTG ACA AGC-3′). PCR products were pooled and sequenced on the MiSeq platform (Illumina).

pmCherry was generated by cloning the PCR-amplified coding region of mCherry (RG1081 5′-CGCGGGCCCGGGATCGCCACCATGGTGAGCAAGGGCGAGG-3′; and RG1082 5′;-TCTAGAGTCGCGGCC TACTTGTACAGCTCGTCC-3′;) into BamHI/NotI digested pEGFP-N1 (Clontech) using In-Fusion HD cloning (Clontech). Plasmids pGFP-1 and pGFP-2 were generated by introducing a BsmBI or an EcoRV site at position nt 137 and nt 141 of the GFP coding region using PCR-based mutagenesis of the pEGFP-N1 plasmid. Plasmids were digested using BsmBI/MfeI (pGFP-1) and EcoRV/PstI (pGFP-2) and gel extracted prior transfection together with pmCherry into 293 cells using genejuice transfection reagent or nucleofection (Amaxa) as indicated. Six hours before transfection, cells were incubated with highly specific inhibitors for LIG1/3 (L67), DNA-PKc (NU7441), and PARP (Olaparib, all from Selleckchem). After transfection, cells were incubated for further 2 days in the presence of inhibitors prior analysis for mCherry/GFP expression using flow cytometry (FC500, Beckman Coulter).

RNA was isolated from purified CLL samples and B cells from healthy controls using High Pure RNA Isolation Kit including DNase digestion (Roche). The same CLL samples as used for plasmid repair assays was used for RNA isolation. First-strand cDNA was generated using iScript (Bio-Rad). Gene expression levels in CLL, Mec1, and B cells were determined using TaqMan assays (Thermo Fisher Scientific) for LIG1 (Hs01553527_m1), XRCC1 (Hs00959834_m1), PARP1 (Hs00242302_m1) with 18S rRNA as controls (Hs99999901_s1).

Cas9 constructs specific for chr17 position 41,621,709 (KRT17 locus) were generated by cloning annealed oligos (RG1087 5′;-ccggGGTGGGTGGTGAGATCAATG-3′; and RG1088 5′;-aaacCATTGATCTCACCACCCACC-3′;) into prelinearized pGuide-it-ZsGreen1 Vector (Clontech). TALENs specific for chr17 position 40,819,524-40,819,583 (KRT10 locus) were kindly provided by Oliver March (Paracelsus Private Medical University Salzburg, Salzburg, Austria). The CLL cell line Mec1 (21) was incubated for 6 hours in RPMI medium supplemented with or without 30 μmol/L LIG1/3 inhibitor L67 followed by Amaxa nucleofection (Lonza) with TALEN/Cas9 constructs. Cells were kept on RPMI medium supplemented with or without L67 for further 2 days prior sorting of GFP-positive cells on a FACS AriaIII (Beckton Dickinson; purity ≥ 94.1% GFP⁺) and isolation of DNA (Qiagen). Chromosomal deletions were PCR amplified using Phusion polymerase and specific primers (RG1142 5′;-TGGCATCTTCTTGGGGTTTA-3′; and RG1143 5′;-CCACATCCCCTTTTTCCATA-3′;). For amplicon sequencing, PCR products from 50 ng template DNA from two independent experiments were pooled and sent for sequencing on the MiSeq platform (Illumina). For quantification of chromosomal deletions, input DNA from cells descending from four independent experiments was titrated from 25 to 1.56 ng and subjected to PCR using primers RG1142/RG1143 and HotStar Taq DNA polymerase (Qiagen).

293 cells, Mec1 cells, and primary cells were cultured in RPMI medium supplemented with 10% FCS (Gibco). Mec1 cells were authenticated by DNA fingerprinting and cytometrical analysis of surface markers (DSMZ). After purchase, stocks of Mec1 cells were frozen and passaged less than 6 months. 293 cells were provided by Stefan Hainzl (EB House Austria, Salzburg, Austria) and periodically authenticated by morphologic inspection. Mec1 and 293 cells were confirmed by fingerprinting at the DMSZ on October 9, 2017. Mec1 and 293 cells were tested negative for mycoplasma in September 2017 using PCR-based mycoplasma detection from cell culture supernatants (PanReac AppliChem). Primary cells were isolated from peripheral blood of CLL patients upon informed consent according to the national guidelines and of the ethics committee and according to the Declaration of Helsinki (Ethics committee approval Salzburg: 415-E/1287/4–2011, 415-E/1287/8–2011). CLL cells and B cells were purified untouched using EasySep system (StemCell Technologies) or MACS (Miltenyi Biotec). Patient characteristics are summarized in Supplementary Table S1. The determination of prognostic markers and FISH analysis for trisomy 12, del11q, del13q, and del17p was performed routinely at our department as described previously (22). In addition, FISH and karyogram analysis of sample 9956 (patient ID660) was performed at the laboratory for molecular biology and tumor cytogenetics at the Ordensklinikum Linz (Linz, Austria).

For cell-cycle analysis, purified B and CLL cells were stained using CD5-FITC (clone: UCHT2; eBioscience) and CD19-PE/Cy7 (clone: SJ25C1; eBioscience) antibodies in PBS at room temperature. Cells were resuspended in prewarmed RPMI without Phenol-Red and stained with vybrant cell cycle dye (Vybrant DyeCycle Violet Stain, V35003; Thermo Fisher Scientific) for 1 hour at 37°C followed by immediate analysis by flow cytometry (Gallios, Beckman Coulter)

Paired end fastq files were acquired by eurofins genomics and GATC-Biotech (Germany). Raw reads were trimmed and selected for paired reads using Trimmomatic v0.33 (PE mode, -phred 33,TRAILING:4; ref. 23), resulting in 8,378,777 paired reads surviving (95.56% of input reads). Forward and reverse reads were merged using FLASh v1.2.11 (-r 300 –f 350 –s 35; ref. 24) and 6,932,326 stiched sequences remained for further analysis. Only merged reads flanked by forward and reverse primers were regarded for further data analyses (only <0.001% of the resulting sequences were forward–forward or reverse–reverse primed; Supplementary Table S1). To avoid missing imprecise repair junctions of resected substrates, we decided to use the unique 6-bp stretch 3′; of the conserved primer-binding sites for mapping. Analyses of read length, frequency, and sequence were performed using custom BASH and PERL scripts, which can be provided upon request. Sequence alignment for detection of deletions was perfomed using MAFFT alignment program (25). Circos plots were created in R using the circlize package (v0.3.8). All other graphs were created in GraphPad Prism 5 and Inkscape v0.91.

To test the efficiency of DSB repair, we set up a plasmid-based repair assay, consisting of a pool of nine plasmids, which harbor individual 350-bp inserts flanked by conserved primer-binding sites (Fig. 1A; Supplementary Fig. S1). Upon digestion with respective restriction enzymes, the individual plasmids yield defined DNA break structures ranging from blunt ends with and without a 6-bp microhomology to cohesive and noncohesive 3′; and 5′; protruding ends with 4 nt overhangs (Fig. 1B). Upon transfection into primary cells, the linear DNA substrates are recognized by the DNA DSB repair machinery and rejoined. The resulting repair junctions can subsequently be PCR amplified using the conserved primer-binding sites and analyzed by NGS. As each of the nine plasmids harbors a unique DNA sequence between the conserved primer-binding sites, all reads obtained by NGS can be exactly mapped to the respective plasmids, allowing examination of efficiency and accuracy of DNA repair in context of DNA end structures and microhomologies.

To detect aberrant DSB repair in CLL cells, we used our pool of linearized plasmid substrates to nucleofect primary CLL cells (CLL-Mut n = 4, CLL-UM n = 3; patient details shown in Supplementary Table S2) and B cells from healthy controls (n = 3). Three days after nucleofection, we isolated DNA and PCR-amplified repair junctions from each sample. Notably, we observed repaired substrates immediately after nucleofection in some CLL samples, which is in line with published data on rapid end joining of DNA ends (Fig. 1C; ref. 26). Upon NGS of pooled amplicons, we obtained a total of 2,676,354 reads, ranging from 96,000 to 587,418 reads for all samples (Fig. 1D).

By calculating the ratio of inter- to intramolecular repair events, we surprisingly obtained a robust bias toward intermolecular repair junctions in CLL samples, irrespective of the BCR mutation status (CLL-Mut and CLL-UM), whereas normal B cells preferred an intramolecular repair (ratio inter/intramolecular repair CLL mean 1.88 ± 0.71 STD; B cells 0.79 ± 0.22, P = 0.017 Mann–Whitney test; Fig. 2A). In particular, we observed significantly increased intermolecular repair of incompatible DNA ends in CLL cells compared with B cells (e.g. joining of plasmid #6 with plasmid #2 or plasmid #8 with plasmid #5), whereas compatible 3′; and 5′; overhangs or blunt ends were repaired at comparable efficiency (Fig. 2B; Supplementary Table S3). None of the repair junctions were significantly overrepresented in B cells compared with CLL cells (Fig. 2B; Supplementary Table S3). Also, by analyzing only intramolecular repair frequencies of the nine different DNA break structures, we observed that noncohesive 5′; ends (substrate #5) as well as incompatible 3′;–5′; overhangs (substrate #7) were repaired with significantly higher efficiency in CLL cells compared with B cells (substrate #5: CLL mean percentage of reads 4.8 ± 2.4 SD, B cells 0.7 ± 0.5; P = 0.004; substrate #7: CLL mean 12.4 ± 5.6 SD, B cells 2.5 ± 2.4; P = 0.005 two-tailed t test assuming unequal variances; Fig. 2C; Supplementary Table S4).

We next aimed at determining differences in the occurrence of deletions at repair junctions between CLL and B cells, resulting from joining of end resected DNA ends. Therefore, we first identified the most frequent repair junction for CLL-Mut, CLL-UM, and B cells for all individual intramolecular repair events and mapped all shorter sequences to the most frequent consensus. We thereby found that the most frequent repair junctions of the nine DNA substrates were the same within each sample group (CLL-Mut, CLL-UM, B cells; Fig. 3A): blunt ends, irrespective of a 6 bp microhomology, and cohesive 3′; and 5′; ends were preferably simply rejoined, whereas 1-bp microhomologies were preferentially used when repairing noncohesive 3′; (substrate #3; Fig. 3A) and 5′; overhangs (substrate #5; Fig. 3A). Incompatible 3′;–5′; overhangs (substrate #7), 3′; overhang/blunt and 5′; overhang/blunt ends (substrate #8 and #9) were preferentially joined and the ssDNA gaps filled up (Fig. 3A).

We next graphically illustrated the occurrence of deletions at repair joints, which we discerned by NGS. In all cases, deletions clustered around the repair junction of the most frequent consensus sequences (Fig. 3B). By comparing the median numbers of resected bases (deletions) at repair junctions, we found that deletions were significantly more frequent in CLL cells compared with B cells (differences in the median number of deleted bases per sequence were significant with P < 0.0001 between CLL-Mut and B cells and CLL-UM and B cells for each substrate; except substrate #4 CLL-UM vs. B cells: P = 0.13; Wilcoxon rank sum test with continuity correction; Fig. 3C).

Upon analyzing the sequences harboring deletions at the repair junction, we observed a high incidence of 1–9 bp microhomologies flanking the deleted region irrespective of the sample group (CLL and B cells; Supplementary Fig. S2). As these microhomologies could at least in part derive from repair by MMEJ and as deletions were occurring at higher incidence in CLL (Fig. 3), this prompted us to test whether the MMEJ pathway is generally more active in CLL compared with B cells. To this end, we more thoroughly analyzed repair junctions of substrate #2, featuring a 6-bp microhomology at both DNA ends. As already shown by Verkaik and colleagues using a similar approach, repair by the MMEJ pathway leads to deletion of one of the two 6-bp microhomologies, whereas repair by c-NHEJ results in direct joining of the DNA ends, yielding a direct repeat of 6 bp within the repair junction (27). Hence, calculating the ratio of the number of junctions deriving from 6 bp microhomology-mediated versus direct joining of substrate #2 reflects MMEJ versus c-NHEJ activity. As shown in Fig. 4A, we found that direct joining of substrate #2 was the preferred repair mechanism in CLL as well as in B cells; however, repair by MMEJ was significantly increased relative to c-NHEJ in CLL cells, irrespective of BCR mutation status (ratio microhomology-mediated/direct joining of substrate #2 for CLL: mean 0.067 ± 0.023 SD, B cells 0.024 ± 0.005 SD; P = 0.017 Mann–Whitney test; Fig. 4A).

Next, we tested whether CLL-specific efficient joining of incompatible DNA ends was solely dependent on microhomology-based repair of end-resected substrates. To test this, we only considered repair junctions mapping to the most frequently occurring sequence of a particular repaired substrate and calculated their relative frequencies, without considering repair of resected substrates. Thereby, we found that again, various incompatible DNA ends were more efficiently repaired in CLL cells compared with B cells, leading to a total of 12 junctions, which were significantly overrepresented in CLL samples (Fig. 4B; Supplementary Table S5). Again, none of the repair events were significantly overrepresented in B cells compared with CLL cells (Fig. 4B; Supplementary Table S5). Although 7 of these 12 junctions revealed short microhomologies (1 to 5 nt, Supplementary Table S5), 5 of these junctions were generated by direct intermolecular joining of 3′;–5′; overhangs or 5′; overhang-blunt ends, implying that incompatible DNA ends were more efficiently joined in CLL cells also independent from terminal microhomologies (Fig. 4C; Supplementary Table S5).

To elucidate the underlying mechanism for biased DSB DNA repair in CLL cells, we first analysed cell-cycle stages in CLL and B cells (Supplementary Fig. S3). However, both CLL samples as well as B cells were mostly in G₁ phase (range 90.2%–95.2% for CLL; 87.5%–97.5% for B cells) with no apparent cell-cycle differences (Supplementary Fig. S3). We next examined expression levels of key factors involved in MMEJ and c-NHEJ in CLL and B cells (9). By extracting data from Haslinger and colleagues (28) using the oncomine database (29) (www.oncomine.org), we found that several genes, including PARP1, XRCC1 (forming a complex with LIG3), and LIG1, which all are involved in MMEJ were significantly upregulated in CLL compared with normal B cells (Fig. 5A). In contrast, within the set of factors involved in c-NHEJ, only DNA-PKcs (PRKDC) and PARP3 showed a slight but significant overexpression in CLL, and none of the factors examined showed downregulation in CLL (Fig. 5A). To validate expression of MMEJ factors in our CLL samples, we performed real-time RT-PCRs. Consistently, LIG1, XRCC1, and PARP1 were all higher expressed in CLL samples compared with B cells from healthy donors (Supplementary Fig. S4). As increased expression levels of MMEJ-specific factors could likely be causative for our observed bias toward MMEJ in CLL cells, we finally wanted to test whether PARP1, LIG1, or LIG3/XRCC1 could also contribute to microhomology-independent joining of incompatible ends, in particular intermolecular joining of 5′; overhangs with blunt ends, which we particularily observed to be increased in CLL samples (Fig. 4B and C). Therefore, we performed a 293-based assay for direct joining of two GFP-based reporter constructs (pGFP-1 and pGFP-2), which encode a functional GFP protein only upon intermolecular direct joining of pGFP-1 (encoding the N-terminal portion of GFP with a 4 nt 5′; overhang) and pGFP-2 (encoding the C-terminal portion of GFP starting with a blunt end; Fig. 5B). GFP expression based on direct joining of pGFP-1/pGFP-2 was analyzed in the presence of specific inhibitors targeting PARP (olaparib), LIG1/3 (L67), and DNA-PKc (NU7441) as control. To normalize for transfection efficiencies, we cotransfected a plasmid encoding mCherry (pmCherry). As expected, direct repair of the GFP plasmids was strongly impeded by inhibition of DNA-PKcs, which are central to c-NHEJ. To a lesser extent also, inhibition of LIG1/3 showed an effect on direct pGFP-1/pGFP-2 joining, while inhibition of PARP family proteins by olaparib had no effect on DNA repair in this assay (Fig. 5C).

Finally, we tested whether LIG1 or XRCC1/LIG3 is involved in the generation of chromosomal deletions between a 5′; overhang DNA DSB and a blunt end DNA DSB in CLL. Therefore, we transiently transfected the CLL cell line Mec1 with constructs expressing TALENs (generating a 5′;overhang) and Cas9 (generating blunt end DSBs), which both cleave at chromosome 17, with or without treatment with 30 μmol/L LIG1/3 inhibitor L67 (Fig. 6A). Of note, real-time RT-PCR revealed that in Mec1 cells, MMEJ-specific factors LIG1, XRCC1, and PARP1 were also more abundantly expressed than in normal B cells (Supplementary Fig. S4). We positively sorted transfected cells based on GFP expression from the Cas9 construct and estimated chromosomal deletion frequencies of a large 0.8 Mb fragment by PCR on serially diluted input DNA isolated from sorted cells (Fig. 6A and B). As estimated from four independent experiments, the chromosomal deletion frequency of L67-treated cells was lower compared with untreated cells, as amplification of breakpoint junctions failed from 1.56 ng input DNA of L67 treated cells in all experiments, whereas in two of four experiments, PCR-amplified breakpoint junctions were detectable in untreated cells at that DNA concentration (Fig. 6B). Amplicon sequencing of the PCR products from two experiments revealed that L67-treated Mec1 cells had an increased frequency of short (5–49 bp) deletions at the breakpoint junction, whereas large (≥50 bp) deletions were reduced (Fig. 6C), again pointing to involvement of LIG1/3 in the occurrence of large deletions at repair joints. Small (0–4 bp) deletions, likely resulting from direct joining or deletions within the 5′; overhang were also reduced in L67-treated cells (Fig. 6C).

Thus, we infer that the described overexpression of LIG1 or XRCC1 in CLL could explain the observed bias toward both, MMEJ as well as increased repair of incompatible DNA ends with increased deletions at repair junctions.

Aberrant repair of DSB can lead to genetic changes, driving carcinogenesis and clonal evolution of cancer. Here, we describe several differences of DSB repair in B cells versus CLL cells by applying a plasmid-based repair assay analyzed by NGS as well as by assessing repair of TALEN/Cas9–induced chromosomal deletions in the Mec1 cell line. Although plasmid-based assays have been used in the past to investigate DSB repair in cell lines and cancer cells (27, 30, 31), our NGS-based approach allowed simultaneous analysis of efficiency and accuracy of DSB repair in regard to diverse DNA break structures and the cellular context at single-nucleotide level. In general, our assay revealed that complementary 5′; protruding ends as well as blunt ends exhibited highest repair efficiency and 3′; cohesive and 3′; noncohesive staggered ends showing weakest joining efficiency. Weak repair of 3′; overhangs was noted in earlier studies (31), probably because 3′; overhangs rather promote HR as they are required for strand exchange reaction in HR (32, 33). However, our repair assay revealed a complex set of substantial differences in efficiency as well as precision of DNA repair in CLL versus B cells. In addition to the frequent occurrence of deletions at the repair junction, which was already described for leukemic cells (30), our assay revealed that noncohesive protruding ends were joined significantly more efficiently in CLL cells compared with B cells.

In our study, efficient joining of incompatible ends was attributed to increased MMEJ activity, reflected by the presence of short microhomologies within repair junctions of end resected incompatible DNA substrates. Concurrently, a 6-bp microhomology was preferentially used in CLL cells for repair of DNA substrates. These results indicate that DNA DSB repair is skewed toward MMEJ in CLL, which is corroborated by overexpression of MMEJ-specific factors in CLL. Notably, this skewing toward MMEJ does not result from cell-cycle differences, as nucleofected CLL cells in our analysis were also G₁ arrested. We further observed CLL-specific increased joining of incompatible DNA ends independent from end resection and usage of microhomologies. Strikingly, this increased joining efficiency was especially pronounced in intermolecular repair events, indicating a somehow increased promiscuity in DNA end joining. Although MMEJ classically involves end resection of DNA overhangs after stabilization of the paired homologous region by several nucleases (34, 35), direct joining of incompatible protruding ends can be mediated by two possibilities: either DNA ends are blunted prior to DNA joining (refilled or resected) or otherwise DNA is resynthesised across a stabilized DNA junction of two staggered DNA ends by polymerase μ or λ, followed by ligation of DNA nicks (36–38). Although ligation of blunted dsDNA ends is normally mediated by LIG4/XRCC4, nick sealing is efficiently performed by LIG1 and LIG3/XRCC1 complexes (39, 40). Our pGFP-1/pGFP-2 repair assay revealed that although most repair events are catalyzed by c-NHEJ involving DNA-PKcs, repair could partly be mediated by polymerase-dependent DNA synthesis across direct pGFP-1/pGFP-2 junctions followed by nick-sealing by LIG1/3. In that way, overexpression of LIG1/3 could not only contribute to increased MMEJ but also to direct joining of incompatible DNA ends. LIG4-deficient cell lines are still able to repair DNA DSBs, albeit the number of direct joints are decreased, and microhomology usage is increased due to activity of LIG1/3 (41, 42). Although sole activity of LIG1/3 in LIG4-deficient cells led to a higher frequency of endonuclease-induced translocations in rodent cell lines, the frequency in human cell lines was decreased, showing that efficiency and kinetics of DNA DSB repair is likely depending on cell type and species specific expression levels of repair factors (41, 43). Our own data suggest that LIG1/3 may significantly contribute to the formation of TALEN/Cas9–induced large chromosomal deletions in the CLL cell line Mec1 as we observed a reduction in their occurrence upon LIG1/3 inhibition, together with a reduction in large deletions at repair joints. Hence, our results would propose that CLL-specific increased expression levels of the MMEJ factors LIG1 and XRCC1 could contribute to increased propensity to acquire chromosomal aberrations during DNA DSB repair. This fits with the observation that chromosomal aberrations are frequently found to be subclonal in CLL, pointing to their successive generation during disease progression (6). In addition, an increased MMEJ activity could also explain the formation of small deletions and microdeletions frequently observed in CLL (44) and also in other cancer entities (45). These deletions often exhibit sequence homologies at breakpoint junctions, suggesting that they likely arise due to MMEJ of DNA DSBs. Alternatively, they could occur by polymerase slippage during replication (46, 47). As we observed several differences in end joining between CLL and B cells, it would be certainly interesting to screen DNA repair quality/quantity in therapy-sensitive versus refractory samples as well as in other clinically relevant CLL subsets (such as samples with complex karyotype). Deriano and colleagues already reported increased error-prone end joining of DNA ends in CLL cells obtained from a therapy-resistant patient (48). Hence, it is conceivable that CLL subset or risk group–specific differences in DNA DSB repair could contribute to diverse clinical outcomes.

Although a portion of CLL patients exhibits mutations at DNA repair genes (6), our data rather suggest an inherently aberrant DNA DSB repair activity in CLL due to different expression levels of key factors for MMEJ, in particular as we excluded del(11q) and del(17p) patients (harboring deleted ATM and TP53 genes) and as all samples included in our analyses showed a very uniform pattern of repair junction formation, irrespective of the BCR mutation status.

Summarizing, we could identify profound differences in efficiency and precision of DSB repair between CLL cells and normal B cells. Considering a reported increased formation of DNA DSBs in precancerous lesions and cancers (49), their imprecise repair due to increased LIG1 and XRCC1 levels could thus continuously enhance the formation of genomic aberrations, contributing to clonal evolution and chemoresistance. Hence, concomitant usage of LIG1/3 inhibitors during CLL treatment would not only potentiate cytotoxicity of DNA-damaging agents (50) but might also impede clonal evolution by acquisition of novel genomic rearrangements.

No potential conflicts of interest were disclosed.

Conception and design: S. Rebhandl, R. Geisberger

Development of methodology: M. Schubert, S. Rebhandl, S. Blaimer, R. Geisberger

Acquisition of data (provided animals, acquired and managed patients, provided facilities, etc.): F.J. Gassner, M. Schubert, S. Rebhandl, N. Zaborsky, S. Blaimer, R. Greil, R. Geisberger

Analysis and interpretation of data (e.g., statistical analysis, biostatistics, computational analysis): F.J. Gassner, M. Schubert, S. Rebhandl, N. Zaborsky, K. Catakovic, D. Hebenstreit, R. Greil, R. Geisberger

Writing, review, and/or revision of the manuscript: F.J. Gassner, M. Schubert, S. Rebhandl, N. Zaborsky, D. Hebenstreit, R. Greil, R. Geisberger

Administrative, technical, or material support (i.e., reporting or organizing data, constructing databases): F.J. Gassner, M. Schubert, S. Rebhandl, K. Spandl, N. Zaborsky, K. Catakovic, R. Geisberger

Study supervision: R. Geisberger

This work was supported by the SCRI-LIMCR, the Province of Salzburg, the City of Salzburg, and grants from the Austrian Science Fund FWF to R. Geisberger (FWF; P24619 and P28201).

The authors thank Oliver March (Salzburg) for providing TALENs and Markus Steiner (Salzburg) for cell sorting.

The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.