Imaging DNA Damage In Vivo Using γH2AX-Targeted Immunoconjugates

Human cancers are frequently associated with DNA damage response (DDR) defects (1). This helps explain the extensive DNA damage observed in tumors at all stages, from dysplasia to advanced disease (2, 3). Commonly used anticancer treatments, including ionizing radiation (IR) and many chemotherapy drugs, exploit defects in DNA repair by inflicting DNA damage to which the tumor cell is unable to mount a normal response. Small molecule inhibitors of components of the DDR can arrest or reverse tumor growth and many are in preclinical or clinical development (4). It follows that an imaging probe capable of revealing DNA damage in vivo could provide useful prognostic information and be used to predict sensitivity to treatment or monitor response to therapy. The ability to image DNA damage would facilitate evaluation of drugs designed to cause tumoral DNA damage or to inhibit its repair.

DNA damage can be measured in human tissues ex vivo by immunohistochemistry, using fluorophore-labeled antibodies that bind specific DNA repair proteins (5) or using flow cytometry (6). However, direct quantification of DNA damage in vivo is not currently possible. In contrast, investigators have reported efforts to image cell death in vivo using, for example, reporter constructs that are activated by caspase-3 cleavage or radiolabeled ligands such as Annexin-V with affinity for apoptotic cells (7, 8). In some cases, only weak correlation between in vivo detection of apoptosis and outcome for a particular tumor was noted (9). This may be because cancer cell death can result from processes other than apoptosis such as mitotic catastrophe, senescence, and autophagy.

DNA double-strand breaks (DNA dsb) are caused either directly by IR and some radiomimetic drugs or indirectly through replication fork stalling (4). The ability to image DNA dsb in vivo would be particularly informative, as they are extremely deleterious and their number and persistence reflect the likelihood of eventual cell death (10). Also, because it is an early event following genotoxic stress, DNA dsb formation would likely predict cell fate, whatever the nature of the initiating insult. There are, however, 2 potential limitations to DNA dsb as a target for imaging. DNA dsb are generally low in number, limiting the sensitivity achievable with most imaging modalities. Also, because DNA dsb exist within the nucleus, they are separated from circulating imaging probes by the cell and nuclear membranes, rendering them inaccessible, particularly to high-molecular-weight antibody-based agents. Both hurdles are surmountable. Although DNA dsb themselves may not be abundant, they do lead to the accumulation of DNA repair proteins that may provide tractable targets. One is the histone H2A variant H2AX, which is phosphorylated on Ser139, starting immediately after DNA dsb formation. Hundreds of copies of phosphorylated H2AX (γH2AX) accumulate in foci at DNA dsb, measuring up to 40 Mbp (11). Detection of these foci, using an anti-γH2AX antibody, forms the basis of a sensitive in vitro assay for DNA dsb (12).

Generally, basal γH2AX expression is high in cancers and exposure to genotoxic stress results in induction of γH2AX that is more prominent and protracted in cancer compared with normal cells (3). Recent reports suggest that enumeration of γH2AX foci in clinical samples may correlate with outcome (13). These observations support investigation of γH2AX as a biomarker of response to DNA-damaging agents.

The problem of inaccessibility of γH2AX to imaging probes, due to its intranuclear location, can be solved through incorporation of cell-penetrating peptides (CPP) and nuclear localizing signals (NLS) into probe design. Tat peptide is an NLS-containing domain of the HIV-1 transactivator of transcription that has cell-penetrating properties (14). Tat (GRKKRRQRRRPPQGYG) mediates transmembrane movement of various cargoes and the presence of NLS (underlined) enables Tat to bind to importins for nucleocytoplasmic trafficking (15). We have shown previously that Tat-containing radioimmunoconjugates (RIC-Tat) can penetrate cell and nuclear membranes and are retained to an extent that correlates with abundance of the molecular target (16, 17).

As a first step to targeting γH2AX for imaging, we attached a Tat-peptide covalently to a fluorophore- or ¹¹¹In-labeled anti-γH2AX antibody. We report here that 2 agents, Cy3–anti-γH2AX-Tat and ¹¹¹In-DTPA–anti-γH2AX-Tat, specifically target DNA dsb. Cy3–anti-γH2AX-Tat colocalizes with γH2AX in the nuclei of irradiated cells. Both Cy3–anti-γH2AX-Tat and ¹¹¹In-DTPA–anti-γH2AX-Tat accumulate in irradiated cancer cells in vitro and in tumors in vivo following DNA damage. Although further development is required to optimize probe design, the proof-of-principle studies presented here show that noninvasive imaging for spatiotemporal tracking of DNA damage may be feasible using γH2AX-targeted radioimmunoconjugates (RIC) that contain CPPs.

A diagram of the synthesis pathway is shown (Fig. 1A). ¹¹¹In-DTPA–anti-γH2AX-Tat and fluorophore–anti-γH2AX-Tat, probes designed to target DNA dsb, were synthesized. γH2AX antibody (100 μg; Calbiochem) or IgG from mouse serum (mIgG; Sigma-Aldrich) was dissolved in 2-(N-morpholino)ethanesulfonic acid (0.1 mol/L). Tat (GRKKRRQRRRPPQGYG) incorporation was achieved using N-(3-dimethylaminopropyl)-N′-ethylcarbodiimide/N-hydroxysuccinimide (EDC/NHS; Pierce) activation. Tat-peptide was added in 5-fold molar excess for 2 hours at room temperature. Unconjugated Tat was removed using Sephadex G50 gel filtration (SEC) columns, giving Tat/IgG ratio of 5:1. Anti-γH2AX-Tat was incubated with isocyanatobenzyl diethylenetriaminepentaacetic acid (Macrocyclics) or was fluorophore labeled using Cy3- or AF488/AF555-NHS ester. DTPA conjugation using cyclic DTPA anhydride, ¹¹¹In labeling, instant thin layer chromatography (ITLC), and determination of DTPA conjugation efficiency was done as previously described (18). Tat/IgG conjugation ratio was determined by radioiodination of the Tat-peptide as previously described (19). DTPA–anti-γH2AX-Tat conjugate was radiolabeled using ¹¹¹In-chloride and radiolabeling yield, determined by ITLC, was 95% or more. The specific activity range was 0.5 to 6 MBq/μg. Chemical and radiochemical purity determined by size exclusion radio high performance liquid chromatography of the final products was 95% or more. Control probes, ¹¹¹In-DTPA–anti-mouse IgG-Tat (¹¹¹In-DTPA–anti-mIgG-Tat) and Cy3- or AF488/AF555–anti-mIgG-Tat, were also synthesized. Tat-conjugated anti-γH2AX or mIgG were labeled with Cy3, AF555, or AF488, using Cy3-NHS (Amersham), AF555-NHS, or AF488-NHS (Invitrogen). Fluorophore labeling was efficient, with Cy3/IgG ratio of 6:1.

Human breast cancer cell lines MDA-MB-468 (LGC Standards), MDA-MB-231 (LGC Standards), MDA-MB-231 cells stably transfected with HER2 (231-H2N; from Dr Robert Kerbel, Sunnybrook Research Centre, Toronto, Ontario, Canada), and mouse embryonic fibroblasts [(MEF) H2AX^−/− and wild-type (WT); from Dr Andre Nussenzweig, National Cancer Institute, Bethesda, MD] were cultured in 5% CO₂ in Dulbecco's modified Eagle's medium (Sigma-Aldrich) supplemented with 10% fetal calf serum (Invitrogen) and 100 units/mL penicillin/streptomycin (Invitrogen). Cells were tested and authenticated by the providers. The cumulative length of culture was less than 6 months following retrieval. MDA-MB-468 cells (10⁶) were irradiated (0–10 Gy) using an IBL-637 ¹³⁷Cs irradiator (Cisbio International; dose rate 1.0 Gy/min). After incubation for 1 hour, cells were lysed and Western blotting was conducted, probing for γH2AX and β-actin.

231-H2N cells were irradiated (4 Gy) and, after 1 hour, lysed (10 minutes on ice; 50 mmol/L HEPES, 150 mmol/L NaCl, 2.5 mmol/L EGTA, 1 mmol/L EDTA, 1 mmol/L DTT, 0.1% Tween-20, 10% glycerol, with added protease and phosphatase inhibitor). Lysates were used to coat wells of a radioimmunoassay (RIA) plate, incubated overnight at 4°C, washed and blocked [PBS, 2% bovine serum albumin (BSA), 0.5% Tween-20]. ¹¹¹In-DTPA–anti-γH2AX (1 nmol/L) plus increasing concentrations of anti-γH2AX antibody or DTPA–anti-γH2AX-Tat in PBS (100 μL) were added for 2 hours at 4°C. Plates were washed, and radioactivity in each well was determined. IC₅₀ values for competition binding were determined.

Internalization, nuclear localization, and retention of ¹¹¹In-DTPA–anti-γH2AX-Tat and ¹¹¹In-DTPA–mIgG-Tat in MDA-MB-468 cells were determined as previously described (19).

MDA-MB-468 cells or MEFs were grown on coverslips and exposed to fluorophore–anti-γH2AX-Tat or fluorophore–mIgG-Tat (0.125 μg/mL). After incubation at 37°C for 1 hour, cells were irradiated (4 Gy) and 1 or 23 hours later washed, fixed using 4% formaldehyde, mounted with 4′,6-diamidino-2-phenylindole (DAPI) or permeabilized using 1% Triton X-100, blocked (1 hour, 37°C; 0.1% Triton X-100, 2% BSA in PBS), and stained for γH2AX using an AF633-labeled secondary antibody. To investigate the spatial relationship between fluorphore–anti-γH2AX-Tat and γH2AX foci, MDA-MB-231 cells were grown on 0.9-μm mylar film (DuPont Teijin Films UK Ltd.) and incubated with AF555–anti-γH2AX-Tat or AF555–mIgG-Tat (0.125 μg/mL). After 1 hour, cells were exposed to 1.5 keV X-rays through a gold mask with 1-μm slits every 10 μm, resulting in DNA damage in “bands” (20). The mean dose to cells was approximately 5 Gy (local dose was ∼50 Gy). After incubation for 1 hour, cells were stained for γH2AX by using an AF488-conjugated secondary antibody.

Animal procedures were carried out in accordance with the UK Animals (Scientific Procedures) Act 1986 and with local ethical committee approval. MDA-MB-468 xenografts were established in female BALB/c nu/nu mice (Harlan). RICs (10 μg; 5 MBq) were administered i.v. γH2AX was induced by intraperitoneal (i.p.) administration of bleomycin (10 μg) 2 hours prior to injection of RIC or by X-ray irradiation of the tumor (10 Gy) 1 hour after injection of RIC. Radiation was delivered using a Gulmay 320 kV X-irradiator at a rate of 2.0 Gy/min. For optical imaging, mice were anesthetized using isoflurane 24, 48, or 72 hours after RIC injection. Fluorescence imaging was done using an IVIS200 system (Caliper LifeSciences). Autofluorescence correction was done using the built-in correction filter set. For single photon emission computed tomography (SPECT), mice were anesthetized using isoflurane at 24, 48, and 72 h after RIC injection, and SPECT-CT was done using a nanoSPECT/CT scanner (Bioscan). At 72 hours, mice were euthanized and organs were removed, weighed, and counted for radioactivity.

Statistical analysis and curve fitting was done using the GraphPad Prism software package (GraphPad Software). Results were compared using 1-way ANOVA with post hoc Tukey test. Fitted parameters from nonlinear regression curve fits were compared using the F test. A P value of < 0.05 was considered significant.

The cell line MDA-MB-468 was used for evaluation of RIC-Tats. We previously showed induction of γH2AX in MDA-MB-468 cells in response to IR, using an immunofluorescence assay for γH2AX foci (21), and confirmed this by Western blot analysis (Supplementary Fig. S1). To confirm affinity of binding of ¹¹¹In-DTPA–anti-γH2AX-Tat to γH2AX, a competition RIA was conducted (Fig. 1B). 231-H2N cells were irradiated (4 Gy) and incubated for 1 hour to allow the formation of γH2AX. Whole-cell lysates, containing γH2AX, were used to coat wells of an RIA plate. ¹¹¹In-DTPA–anti-γH2AX (1 nmol/L) plus either unlabeled DTPA–anti-γH2AX-Tat or anti-γH2AX (range, 0–1,000 nmol/L) was added. The radioactivity in each well was determined by quantitative autoradiography. The ability of DTPA–anti-γH2AX-Tat and anti-γH2AX to compete with ¹¹¹In-DTPA–anti-γH2AX for binding to γH2AX was similar with log(IC₅₀) of 0.33 ± 0.12 and 0.42 ± 0.14 (mean ± SD), respectively (P = 0.70), indicating that the affinity of the modified antibody, DTPA–anti-γH2AX-Tat, for the target epitope is similar to that of the unaltered antibody, anti-γH2AX.

To investigate the ability of the RIC-Tats to penetrate the cell membrane and accumulate in the nuclei of cells, cell radioactivity internalization and retention experiments were done. To evaluate internalization, MDA-MB-468 cells were irradiated (4 Gy) or sham-irradiated and exposed to ¹¹¹In-DTPA–anti-γH2AX-Tat or ¹¹¹In-DTPA–mIgG-Tat for 0 to 4 hours. At selected times, cells were harvested, fractionated, and the radioactivity in whole cells and in membrane as well as in cytoplasmic and nuclear fractions measured (Fig. 1C and D; Supplementary Fig. S2A and B). The amount of ¹¹¹In that internalized and translocated to nuclei peaked at 15 to 30 minutes in irradiated and nonirradiated samples, reaching approximately 0.13% of the added radioactivity. The nuclear ¹¹¹In content slowly diminished, reaching approximately 0.1% of the total added radioactivity by 4 hours. To investigate retention of ¹¹¹In, MDA-MB-468 cells were loaded with ¹¹¹In-DTPA–anti-γH2AX-Tat or ¹¹¹In-DTPA–mIgG-Tat by exposing them to radiolabeled probe for 1 hour. Cells were then irradiated (4 Gy) or left unirradiated. Radiopharmaceutical-containing medium was replaced with fresh medium, and the amount of ¹¹¹In in the cells was measured for up to 50 hours. In nonirradiated cells, exposed to either ¹¹¹In-DTPA–mIgG-Tat or ¹¹¹In-DTPA–anti-γH2AX-Tat, the amount of retained ¹¹¹In fell rapidly reaching background levels by 10 hours (Fig. 1E). This was also true of irradiated cells, in which γH2AX expression was induced, when exposed to ¹¹¹In-DTPA–mIgG-Tat (half-life 0.23 ± 0.09 hours). However, retention of ¹¹¹In did occur in irradiated compared with nonirradiated cells exposed to ¹¹¹In-DTPA–anti-γH2AX-Tat (half-life 27.71 ± 11.01 hour vs. 1.81 ± 0.87 hour, respectively; P = 0.0076, F test). Thus, the anti-γH2AX-Tat probe was retained only in cells in which γH2AX was induced.

MDA-MB-468 cells were exposed to AF488–anti-γH2AX-Tat or AF488–mIgG-Tat for 1 hour and irradiated (4 Gy) or sham-irradiated. After 1 or 23 hours, cells were fixed and mounted with DAPI and stained for γH2AX (Fig. 2A and B). AF488-anti-γH2AX-Tat was synthesized using an anti-γH2AX antibody raised in rabbit, whereas γH2AX foci were detected using mouse-anti-γH2AX antibody and secondary AF555-labeled mIgG. MDA-MB-468 cells exposed to AF488–mIgG-Tat showed membrane and cytoplasmic staining at 1 hour (attributed to Tat-mediated internalization of probe) that disappeared by 23 hours postirradiation (Fig. 2A and B). At no time was nuclear fluorescence observed in cells exposed to AF488–mIgG-Tat, whether irradiated or not. Fluorescence was not observed in the nuclei of sham-irradiated cells exposed to AF488–anti-γH2AX-Tat. However, cells exposed to AF488–anti-γH2AX-Tat followed by IR exposure showed focal nuclear uptake of AF488 at 1 and 23 hours. Immunostaining for γH2AX showed that many γH2AX foci formed by 1 hour following IR exposure and were still detectable at 23 hours. γH2AX foci colocalized with the fluorescence resulting from nuclear accumulation of AF488–anti-γH2AX-Tat (Fig. 2A and B). This was confirmed by generating a cytofluorogram (CFG) obtained by plotting, for each pixel, AF488 fluorescence (green) on the x-axis and AF555 fluorescence (red), denoting the presence of γH2AX foci, on the y-axis. Coincidence of the 2 signals was observed.

It was shown that the presence of fluorophore–anti-γH2AX-Tat at the time of irradiation does not itself interfere with DNA damage repair, as the number of γH2AX foci at 23 hours after IR exposure is the same whether fluorophore–anti-γH2AX-Tat is present or not (Supplementary Fig. S3A).

To evaluate dependence of fluorophore–anti-γH2AX-Tat signal on the presence of γH2AX foci, a slit-irradiation technique was used (20). Cells were grown on mylar film, exposed to AF555–anti-γH2AX-Tat or AF555–mIgG-Tat, and irradiated through a gold film mask, resulting in cells being irradiated in 1-μm stripes. γH2AX foci formed in stripes and AF555–anti-γH2AX-Tat, but not AF555–mIgG-Tat, colocalized with γH2AX foci (Fig. 3A).

To confirm that retention of anti-γH2AX RIC depends on the presence of γH2AX, WT and H2AX-null MEFs were exposed to Cy3–anti-γH2AX-Tat for 1 hour, irradiated or sham-irradiated, and after 1 or 23 hours, fixed, mounted with DAPI, and stained for γH2AX foci (Fig. 3B; Supplementary Fig. S3B). Colocalization of γH2AX foci with Cy3 was observed in irradiated WT MEFs, but not H2AX^−/− MEFs, exposed to Cy3–anti-γH2AX-Tat. Efficient DNA damage repair in WT MEFs results in rapid resolution of γH2AX foci, accounting for the absence of fluorescence due to Cy3–anti-γH2AX-Tat and AF488 (γH2AX foci) in nuclei at 23 hours (Supplementary Fig. S3B).

These in vitro studies indicate that anti-γH2AX RICs accumulate specifically at γH2AX foci in the nuclei of irradiated cells. Low-magnification microscopy images of irradiated cells exposed to AF55–anti-γH2AX-Tat indicate that colocalization of the probe with γH2AX foci is seen in all nuclei that contain γH2AX foci (Supplementary Fig. S3C).

To test whether anti-γH2AX-Tat RIC can visualize DNA damage in vivo, MDA-MB-468 xenograft–bearing mice were scanned using an IVIS-300 optical camera at 24, 48, and 72 hours following i.v. administration of Cy3–mIgG-Tat or Cy3–anti-γH2AX-Tat and exposure to IR (10 Gy), delivered to the xenograft-bearing hind limb, to induce intratumoral γH2AX. Accumulation of Cy3–anti-γH2AX-Tat was detectable in irradiated but not control tumors. There was no uptake of Cy3–mIgG-Tat in irradiated or control tumors (Fig. 4). Accumulation of Cy3–anti-γH2AX-Tat in irradiated tumors peaked at 48 hours and returned to baseline by 72 hours.

SPECT was conducted 24, 48, and 72 hours following administration of ¹¹¹In-DTPA–anti-γH2AX-Tat or ¹¹¹In-DTPA–mIgG-Tat to mice treated with bleomycin (10 μg) by i.p. injection or IR (10 Gy) delivered to the xenograft. Transverse and coronal images through tumors are shown (Fig. 5A; Supplementary Fig. S4A). Volume-of-interest (VOI) analyses of tumors were conducted (Fig. 5B). In untreated controls, tumor uptake of ¹¹¹In-DTPA–anti-γH2AX-Tat was greater than ¹¹¹In-DTPA–mIgG-Tat at each time point. The uptake of ¹¹¹In-DTPA–anti-γH2AX-Tat expressed as percentage of the injected dose per gram of tumor (% ID/g) at 24 hours p.i. was 3.6 ± 0.1 compared with 1.2 ± 0.1 for ¹¹¹In-DTPA–mIgG-Tat (P < 0.0001). Tumor uptake of ¹¹¹In-DTPA–mIgG-Tat did not increase significantly following treatment with bleomycin or IR compared with controls, with uptake at 24 hours p.i. of 1.7 ± 0.7% and 1.7 ± 0.2 versus 1.2 ± 0.1% ID/g, respectively; P > 0.05. However, both bleomycin and IR led to significantly increased intratumoral accumulation of ¹¹¹In-DTPA–anti-γH2AX-Tat compared with controls, with uptake at 24 hours p.i. of 5.4 ± 1.2 and 5.2 ± 0.6 versus 3.6 ± 0.1% ID/g, respectively; P < 0.0001. At 72 hours, uptake of ¹¹¹In-DTPA–anti-γH2AX-Tat in irradiated tumor had returned to basal levels whereas uptake in bleomycin-treated tumors remained elevated. There was no evidence of accumulation of ¹¹¹In-DTPA–anti-γH2AX-Tat in the normal tissues of the irradiated hind limb compared with the unirradiated hind limb over the time course investigated, including at an early time point (2 hours; Supplementary Fig. S4B). ¹¹¹In-anti-γH2AX-Tat, at the specific activities used for SPECT imaging (<1 MBq/μg), did not reduce cell survival (evaluated in clonogenic assays; Supplementary Fig. S5).

Following acquisition of images at 72 hours, mice were euthanized and selected organs removed and radioactivity counted. This confirmed significant retention of radioactivity in tumors of bleomycin-treated animals at 72 hours (Supplementary Fig. S6; Supplementary Table S1).

To show that tumor uptake of anti-γH2AX RIC correlates with the abundance of γH2AX, xenografts were exposed to IR (0–4 Gy) following administration of ¹¹¹In-anti-γH2AX-Tat. SPECT images acquired 23 hours p.i. show that intratumoral accumulation of the probe increased with increasing IR dose. Transverse images through tumor are shown (Fig. 6A). Tumors and muscle were removed and radioactivity counted for calculation of tumor/muscle (T:M) ratio. A positive correlation between IR dose and intratumoral accumulation of ¹¹¹In was observed (Spearman r = 0.9; P = 0.042; Fig. 6B). Tumors were sectioned and stained for γH2AX, showing that tumors exposed to higher doses of IR contained the greatest number of γH2AX foci (Fig. 6C).

Many common anticancer treatments cause DNA damage. These agents are efficacious because cancer cells are often DDR-impaired and, therefore, are unable to recover from genotoxic stress. IR and several types of chemotherapy, including alkylators, topoisomerase inhibitors, and replication inhibitors, cause DNA dsb, leading to cell death. DNA dsb also result from replication fork stalling, which can be caused by alkylating and antimetabolite drugs (4). Phosphorylation of H2AX by phosphoinositide 3-kinase (PI3K)-like kinases, to form γH2AX, is an early and almost universal feature of the eukaryotic response to DNA dsb. Detection and quantification of γH2AX expression are therefore used as measures of DNA damage.

A recently proposed model of tumorigenesis implicates activated oncogenes as a cause of replication stress and DNA damage (1). This phenomenon, “constitutive DNA damage,” may account for the frequent observation of high γH2AX expression in precancerous lesions and cancers but not in adjacent normal tissue. High γH2AX expression has been described in many types of cancer and in their precursor lesions (2, 22–24). The kinetics of γH2AX induction differs in malignant compared with normal cells (10). The number of γH2AX foci per nucleus increases rapidly following irradiation of normal tissues, returning to baseline levels by approximately 24 hours p.i. (25). In tumors, however, elevation of γH2AX expression is protracted and evident up to 72 hours p.i. (26). These differences in normal versus malignant tissues have stimulated interest in γH2AX as a diagnostic and pharmacodynamic biomarker (27). However, the techniques currently available to study γH2AX expression rely on the analysis of clinical samples ex vivo (28). A method for whole-body imaging of γH2AX would, therefore, be a powerful tool with which to study the spatiotemporal dynamics of DDR.

An average cell contains 30 × 10⁶ nucleosomes (29), with 2 H2A proteins per nucleosome, that is 60 × 10⁶ H2A proteins per cell. Approximately 10% of H2A is present in the H2AX isoform in mammalian cells, although up to 25% has been reported (30). This means that there are about 6 × 10⁶ copies of H2AX per cell. It has been estimated that roughly 2% of H2AX molecules become phosphorylated for every Gy of IR to which cells are exposed (30). Therefore, the number of copies of γH2AX formed following doses of 2 to 10 Gy IR would be 2.4 × 10⁵ to 1.2 × 10⁶. In cells with a higher proportion of H2AX (25%), the numbers would be 6 × 10⁵ and 3 × 10⁶. Each copy of γH2AX could be bound by a Tat-conjugated anti-γH2AX RIC. This number of targets (∼10⁶ per cell) is similar to that of other tumor antigens that have been explored for molecular imaging such as HER2 and epidermal growth factor receptor (EGFR). Also cancers overexpress γH2AX compared with surrounding normal tissues, even in the absence of exposure to a genotoxic treatment (2, 22–24, 31). Therefore, it is possible that the number of copies of γH2AX in some cancers is already high and is then induced further following treatment. These lines of evidence suggest that the abundance of γH2AX expression in cancers would be sufficient for molecular imaging.

We have seen no evidence of accumulation of ¹¹¹In-DTPA–anti-γH2AX-Tat in irradiated normal tissues over the time course investigated. This can be appreciated by comparing the appearance of the right (irradiated) hind limb with the left (nonirradiated) hind limb in the images shown in Figure 4 and Supplementary Figure S4. We speculate that this is because the resolution of γH2AX foci in normal tissue is so rapid and complete that the probe does not accumulate sufficiently to produce a detectable signal. The rapid resolution of γH2AX in the normal cell line WT MEF compared with the protracted expression of γH2AX in a malignant cell line, MDA-MB-468, after irradiation suggests that this explanation may be correct (Supplementary Fig. S3A and B). Bleomycin has been shown to accumulate in tumors. Hou and colleagues, for example, showed that the tumor/blood ratio for [⁵⁷Co]-bleomycin at 48 hours was more than 100 in a murine hepatoma model (32). Intratumoral concentration of bleomycin would cause prolonged induction of γH2AX and this may account for the observation that ¹¹¹In-DTPA–anti-γH2AX-Tat signal is more protracted in bleomycin-treated compared with IR-treated tumors (Supplementary Figs. S3 and S6).

The internalization of anti-γH2AX RIC-Tats into cells is not influenced by γH2AX expression, because Tat peptides are able to penetrate almost any cell (14). However, anti-γH2AX RICs are retained only in the nuclei of cells that express γH2AX, resulting in target/nontarget contrast. We have shown that fluorophore-labeled anti-γH2AX-Tat colocalizes with γH2AX foci that were simultaneously visualized using a standard immunofluorescence method. This indicates that anti-γH2AX RIC-Tats bind their molecular target and accounts for their intranuclear retention in γH2AX-expressing cells. In vivo, fluorophore- and radiolabeled anti-γH2AX RIC-Tats specifically accumulate in tumors in which DNA damage has been induced by IR or chemotherapy.

Cy3–anti-γH2AX-Tat is suitable for imaging γH2AX in vitro and superficial tumors in mice but not for imaging deep-seated tumors because of light attenuation by overlying structures. Therefore, we developed an ¹¹¹In-labeled probe. ¹¹¹In, which has γ-emissions of 245 and 171 kV, is commonly used in nuclear medicine. For example, ¹¹¹In-labeled leukocytes, octreotide, anti-CD20 antibody and capromab pendetide are used routinely to image inflammation, neuroendocrine tumors, lymphoma, and prostate cancer, respectively. Both the γ-photons and the short-range Auger electrons that ¹¹¹In emits are capable of causing DNA damage, although this is unlikely to be significant when small quantities of isotope are used for imaging. To increase their potential applications, it would be possible to label anti-γH2AX RICs with other isotopes such as ^99mTc or ⁸⁹Zr for SPECT and PET imaging.

It has been shown that anti-γH2AX RICs can image DNA damage in vitro and in vivo. Further optimization of the probes is desirable. Currently, cellular internalization of anti-γH2AX RICs is nonspecific. Modest tumor uptake of radioactivity was seen following administration of the control agent ¹¹¹In-DTPA–mIgG-Tat, which may be due to the enhanced permeability and retention (EPR) of tumors (33). The retention of ¹¹¹In-DTPA–anti-γH2AX-Tat but not ¹¹¹In-DTPA–mIgG-Tat in tumor, however, can be ascribed specifically to γH2AX upregulation. It may be possible to improve tumor-specific uptake of anti-γH2AX RICs by incorporation of a tumor-seeking moiety such as a receptor-binding peptide. We have reported success with this strategy previously (17). The kinetics of γH2AX formation and resolution following genotoxic stress are rapid in normal tissues, with removal of foci starting from about 30 minutes. Injected antibodies, on the other hand, have long half-lives and penetrate tissue slowly. Therefore, refinement of the anti-γH2AX RIC by replacement of antibody by Fab' might provide more favorable pharmacokinetics and allow imaging of rapid changes in γH2AX content.

In summary, we report initial evaluation of molecularly targeted contrast agents designed to image DNA dsb. Labeled anti-γH2AX-Tat immunoconjugates hold promise as clinical tools for noninvasive imaging of DNA damage.

No potential conflicts of interest were disclosed.

This research was supported by the CR-UK/EPSRC/MRC/NIHR Oxford Cancer Imaging Centre and the NIHR Oxford Biomedical Research Centre and through a grant from Cancer Research-UK (C14521/A6245).

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.