Auranofin Protects Intestine against Radiation Injury by Modulating p53/p21 Pathway and Radiosensitizes Human Colon Tumor Free

Radiation-induced intestinal injury is a limiting factor for definitive chemoradiotherapy for abdominal malignancies such as gastric, pancreatic, and colorectal cancer. Thus, tumoricidal radiation doses cannot be administered in the treatment of abdominal tumors, resulting in poor survival and early metastatic spread. The radioresistance of abdominal tumors further reduces the efficacy of radiotherapy. Even with advanced intensity-modulated radiotherapy (IMRT), significant toxicity in the small bowel was noted, which also reduced quality of life in patients. Therefore, a selective radioprotector for normal tissue and/or a radiosensitizer for tumor tissue is needed to improve efficacy of radiotherapy and achieve a complete cure. In this study, we have shown for the first time that a repurposed antirheumatoid agent, auranofin (AUR), sensitizes colon tumors while protecting nonmalignant gastrointestinal epithelium.

Intestinal epithelium is a radiosensitive organ due to its high cell-renewal rate. Radiation injury to intestine induces a rapid loss of mucosal epithelium, resulting in malabsorption, diarrhea, bacterial influx, and sepsis (collectively, the gastrointestinal or GI syndrome), leading to death. The epithelial response against radiation exposure to the small intestine is regulated by the tumor suppressor protein p53. In some cell types, ionizing radiation–induced activation of p53 promotes cell-cycle arrest by inducing the cyclin-dependent kinase (cdk) inhibitor p21 (1–7), which may allow time for repair of DNA damage. p53^−/− mice are more susceptible to the GI syndrome when compared with mice containing two copies of the wild-type (WT) p53 gene (7–9). Transgenic mice, overexpressing p53 in all tissues, were protected from the GI syndrome after irradiation. However, pharmacologic induction of p53 signaling and its effect on radioprotection of the GI epithelium have not been studied.

Auranofin, a gold-containing compound, has been used clinically to treat rheumatic arthritis since 1985. It has also been reported that auranofin has anticancer effects (10–13). Auranofin was recently tested in a phase II clinical trial of chronic lymphocytic leukemia (http://clinicaltrials.gov/ct2/show/NCT01419691). To date, auranofin has been administered orally with only minor GI adverse events reported. To minimize any possibility of GI toxicity and to have maximum control of in situ drug levels, we used an intraperitoneal route throughout this study. We have demonstrated that auranofin, a deubiquitinase (DUB) inhibitor, stabilizes p53 by inhibiting DUB HAUSP7 involved in regulation of p53 stabilization (14). Pretreatment with auranofin induces p53/p21–mediated reversible cell-cycle arrest to reduce radiation toxicity in nonmalignant GI epithelium and thereby ameliorates the radiation-induced gastrointestinal syndrome (RIGS).

According to previous reports, auranofin targets thioredoxin reductase, thus inducing the generation of reactive oxygen species (ROS; refs. 15–17). However, a separate set of studies confirms that auranofin-mediated apoptosis is associated with inhibition of proteosomal degradation rather than ROS generation. Auranofin inhibits proteasome-associated DUBs to impair proteosomal degradation and thereby induces cytotoxicity in malignant cells (18). The anticancer effect of auranofin has been tested in many preclinical models. It has been shown that auranofin promotes cytotoxicity in human chronic leukemia and gastric cancer cells with the induction of lethal endoplasmic reticulum (ER) stress. The hypoxic tumor microenvironment generates oxidative ER stress, resulting in protein misfolding and unfolded protein response (UPR). Inhibition of proteosomal degradation increases the misfolded or unfolded protein burden. The accumulation of misfolded proteins in the ER produces a stress response, which induces HSPs to initiate cytoprotective measures to correct protein misfolding and restore normal ER function (19–22). These responses are collectively known as UPR. If the correction machinery fails to control erroneous protein burden, resulting in prolonged activation of the UPR, the cytoprotective response switches to initiate programmed cell death or apoptosis. Thus, the signaling cascade of UPR involves both cytoprotective and apoptotic/cell death pathways. In this manuscript, we have demonstrated that combination treatment of auranofin and radiotherapy increases the misfolded protein load beyond the tolerance level and thereby induces cell death with the activation of a UPR downstream apoptotic pathway. We have also demonstrated that in organoids derived from human colonic surgical specimens, auranofin protects the nonmalignant tissue but sensitizes malignant organoids to radiation. Therefore, this is the first report demonstrating the dual role of a small molecular agent to promote efficacy of radiotherapy both in mice and human tissue.

Five- to 6-week-old male C57BL/6 and BALB/c (Jackson Laboratory) mice were maintained ad libitum. All studies were performed under the protocols approved by the Institutional Animal Care and Use Committee of the University of Kansas Medical Center (ACUP Number: 2016-2316).

The murine colorectal carcinoma cell lines CT26 (ATCC CL-2638) and IEC6 (ATCC CRL-1592) were obtained from the ATCC (with authentication by short tandem repeat) and maintained in complete medium as described previously (23). Cell lines were routinely characterized in the laboratory based on morphology and gene expression patterns. Cells were confirmed to be free of Mycoplasma contamination using a Mycoplasma Detection Kit (Thermo Fisher Scientific). Cells were maintained with RPMI1640 medium (Corning Cellgro) supplemented with 10% heat-inactivated FBS (Corning Cellgro) in a 5% CO₂ atmosphere at 37°C.

Surgically resected tumor and normal colon from patients were obtained through the University of Kansas Medical Center Biospecimen Repository.

Human colon tissues were received from the University of Kansas Medical Center Biospecimen Repository (HUS#5929) to perform organoid culture as described previously (24, 25). The tissue was scraped and chopped into approximately 5-mm pieces. The tissue was then washed with cold PBS, and incubated in dissociation buffer for 20 minutes with agitation at room temperature. The tissue fragments were suspended vigorously with a 10-mL pipette in cold 0.1% BSA in PBS, yielding supernatants enriched in epithelial cells. Samples were passed through 100-μm filters (BD Biosciences) to obtain fraction 1. The same step was repeated three times to get fractions 2, 3, and 4. Fractions 3 and 4, enriched in crypt stem cells, were centrifuged at 300 × g for 5 minutes at 4°C and diluted with advanced DMEM/F12 (Invitrogen) containing B27, N2, 1 μmol/L n-Acetylcysteine, 10 mmol/L HEPES, penicillin/streptomycin, and Glutamax (all Invitrogen). Crypts were passed through 100-μm filters (BD Biosciences), and centrifuged at 275 × g for 5 minutes at 4°C and single cells were discarded. Crypts were embedded in extracellular matrix (BD Biosciences) and plated in a prewarmed 24-well plate. After the matrix/Matrigel solidified, Intesticult Medium (Stemcell Technologies) with supplements and gentamycin (50 μg/mL) were overlaid. Passage was performed once per week. The number of organoids per well was counted by Evos Microscope (Thermo Fisher Scientific). Images of organoids were acquired using fluorescent (Nikon, 80i) and confocal (Nikon, A1RMP) microscopy. The total number of crypt structures and number of budding crypts were quantified and expressed as a ratio of budding crypts/total crypt structures.

The single-cell suspension from human normal and tumor colon–derived organoids was prepared by dissociating the organoid with Cell Dissociation Media (Corning Cellgro) as per the manufacturer-provided protocol. Human p53 and p21 siRNA were purchased from Santa Cruz Biotechnology. The transfection of p53 and p21 siRNA and control-scrambled RNAs in the normal colon–derived organoid cells was carried out using Lipofectamin 2000 (Thermo Fisher Scientific) as per the manufacturer-provided protocol. Western blot analysis was done to confirm the silencing of p53 and p21 protein expression.

MTT assay was performed as reported earlier (26). A total of 5,000 cells in 100-μL medium per well were plated in 96-well plates and allowed to adhere for 24 hours. Auranofin (Enzo Life Sciences) was added at indicated concentrations for 2 hours, and cultures were rinsed with fresh medium and reincubated for an additional 24 hours. Next, 10 μL of the MTT Reagent (5 mg/mL, Promega) was added for 2 hours, and 150 μL of DMSO was admixed to dissolve the formazan crystals. Absorbance was measured at a wavelength of 540 nm using a spectrophotometer.

Clonogenic assay was performed as reported earlier (27). Cells were seeded in a 6-well plate and treated with 2 μmol/L auranofin for 24 hours; then X-irradiated at a dose rate of 2.26 Gy/minutes (XENX, XStrahl). After 8 days, cultures were fixed with crystal violet and colonies (>50 cells) were counted. Surviving fractions were fitted to the linear quadratic model using GraphPad Prism 5 Software (GraphPad Prism Software Inc.). The auranofin concentration used is similar to the peak concentration of gold that is achieved in humans following chronic treatment with the standard dose of 6 mg daily (28, 29).

Intestinal epithelial cells were prepared from the jejunum of adult male C57Bl6 mice as described previously (25, 30). A catheter was inserted into the intestine of anaesthetized mice through an incision in the most proximal part of duodenum. Another incision was made just proximal to the caecum to perfuse the entire small intestine with ice-cold PBS followed by flushing twice with ice-cold PBS plus 1 mmol/L dithiothreitol (DTT). The entire jejunum was tied at the distal end and filled with isolation citrate buffer (0.9% w/v NaCl, 1.5 mmol/L KCl, 27.0 mmol/L NaCitrate, 8.0 mmol/L KH₂PO_4, and 5.6 mmol/L Na₂HPO₄, pH 7.3) followed by incubation at 37°C for 15 minutes. After incubation, the fluid was emptied from the jejunum and it was further filled with 5-mL ethylene diamine tetra acetic acid (EDTA) buffer [0.9% w/v NaCl, 8 mmol/L KH₂PO₄, 5.6 mmol/L Na₂HPO₄, 1.5 mmol/L Na₂ EDTA, pH 7.6, plus 0.5 mmol/L DTT, and 0.23 mmol/L phenylmethylsulfonyl fluoride (PMSF); Sigma-Aldrich]. The jejunum was then physically manipulated and tapped allowing the cells to separate from the interior surface. The jejunum was rinsed twice with 5 mL of EDTA buffer and all the fluid containing epithelial cells was collected, centrifuged at 300 × g for 5 minutes, washed twice with 20 mL of balanced salt solution containing 135 mmol/L NaCl, 4.5 mmol/L KCl, 5.6 mmol/L glucose, 0.5 mmol/L MgCl₂, 10 mmol/L HEPES, and 1.0 mmol/L CaCl₂, pH 7.4, and suspended in 2 mL of this solution. The number of viable cells was quantified using an automated Cell Counter T20 (Bio-Rad) with the Trypan blue exclusion method.

Abdominal irradiation (AIR), or whole-body irradiation (WBI) was performed on mice anaesthetized via a continuous flow 1.5 mL/min of 1.5% isoflurane in pure oxygen, using a small animal radiation research platform (XENX, XStrahl) as described previously (25). For AIR, a 3-cm² area of the mouse abdomen encompassing the GI tract was irradiated (Fig. 1), but shielding the upper thorax, head, and neck as well as the lower and upper extremities. This results in significant protection of the bone marrow, while predominantly inducing RIGS. Radiation was delivered to the midline of the abdomen. To ensure homogeneous dose delivery, half of the dose was delivered from the anterior–posterior direction and the other half from the posterior–anterior direction. The total irradiation time to deliver the intended dose was calculated with respect to dose rate, field size for radiation, and fractional depth dose to ensure accurate radiation dosimetry.

As described previously (25), the intestine was dissected, washed in PBS to remove intestinal contents, and the jejunum was fixed in 10% neutral-buffered formalin prior to paraffin embedding. Tissue was routinely processed and 5-μm sections were prepared for hematoxylin and eosin (H&E) and IHC staining. All H&E (Thermo Fisher Scientific) staining was performed at the Pathology Core Facility in the University of Kansas Cancer Center (Kansas City, KS).

Crypt depth was objectively analyzed and quantified in a blinded manner from coded digital photographs of crypts from H&E-stained slides using ImageJ 1.37 software to measure the height in pixels from the bottom of the crypt to the crypt villus junction. The length from the crypt villus junction to villous tip was measured to determine the villous length. This measurement was recorded in pixels and was converted to length (in μm) using the following conversion factor (1.46 pixels/μm).

Apoptotic cells were detected in situ by performing TUNEL (terminal deoxynucleotidyl transferase–mediated dUTP nick end labeling) staining. Briefly, paraffin-embedded sections were deparaffinized, rehydrated through graded alcohol steps, and stained using ApopTag Peroxidase in Situ Apoptosis Detection Kit (Millipore) according to the manufacturer's instructions. A total of 60 crypts were examined per animal.

Mice exposed to AIR and/or pretreated with auranofin were gavaged on day 5 after irradiation with 0.6 mg/kg body weight of a FITC-dextran solution (4,000 kDa size, Sigma-Aldrich). Mice were euthanized and serum was obtained with cardiac puncture 4 hours after gavage (31). Samples were measured in a 96-well plate using a multiwell fluorometer. A standard curve was constructed using mouse serum having increasing amounts of FITC-dextran to determine the serum levels of FITC-dextran in different treatment groups.

A total of 2 × 10⁶ CT26 cells in 200-μL PBS were inoculated subcutaneously in the flanks of 6- to 8-week-old male BALB/c mice. Upon observation of palpable tumors, animals were randomly distributed into treatment groups and treated with either 10% DMSO, auranofin (i.p., 10 mg/kg/day in 10% DMSO), 12.5 Gy as single-fraction AIR or 4 × 4 Gy of fractionated radiation, or a combination of auranofin and radiation as mentioned in the schematic diagrams (Fig. 4A and E). The auranofin dose of 10 mg/kg has been reported as nontoxic when used alone in preclinical studies (11, 32). Auranofin was given three times a week for 3 weeks and radiation was given twice a week for 2 weeks. Radiation was started 1 week after the first auranofin injection. Tumors were measured every other day using Vernier calipers. Tumor volumes were determined by the formula: volume (in cubic millimeters) = W × (L)²/2 (W, width; L, length). When tumor volumes reached 2,000 mm³ or the tumors became necrotic, animals were euthanized (33).

A single-cell suspension from solid tumor was prepared according to a previously described protocol with little modification (34). Briefly, each tumor was excised from the mouse abdomen without tearing the abdominal cavity. Tumors were transferred to a petri plate containing serum-free media and chopped with a razor. Then, the suspension was incubated at 37°C with gentle shaking in 40 μL of Liberase DL stock solution 28 U/mL (Roche), 80 μL of Liberase TL stock solution 14 U/mL (Roche), and 40 μL of DNase I (15 mg/mL stock solution 100×, Sigma-Aldrich) for 30 minutes. The enzymatic reaction was stopped by adding 2% BSA solution. The suspension was then passed through a 100-μm Cell Strainer (BD Falcon) and lysed with RBC Lysis Buffer (BioLegend) so as to prepare a single-cell suspension. These cells were used for DUB activity assay, Western blot analysis, qPCR, and apoptosis assay.

DUB activity was measured according to a protocol reported earlier (35). Briefly, tumor cell lysate and tumor colon organoid cell lysate (5 μg) was dissolved in ice-cold DUB buffer containing 50 mmol/L/L Tris-HCl (pH 7.5), 250 mmol/L sucrose, 5 mmol/L MgCl₂, and 1 mmol/L PMSF, then incubated with Ub-AMC Substrate (Enzo Life Sciences) in a 100-μL reaction volume at 25°C. AMC release generated from the cleaved substrate was temporally recorded with the microplate reader of a fluorescent spectrophotometer.

USP7 (HAUSP) inhibitor activity of auranofin was determined using the USP7 Inhibitor Screening Assay Kit (BPS Biosciences) as per the manufacturer's protocol. The USP7 Inhibitor Screening Assay Kit is designed to measure USP7 activity for screening and profiling of HAUSP7 inhibitor. The USP7 Inhibitor Screening Assay Kit contains fluorogenic, ubiquitinated substrate Ub-AMC, assay buffer, and purified USP7 enzyme. USP7 is incubated with a sample containing assay buffer and Ub-AMC Substrate in presence/absence of HAUSP7 inhibitor and activity was measured by fluorimetric assay on a fluorimeter.

Whole-cell lysates were prepared in RIPA buffer supplemented with 10 mmol/L β-glycerophosphate, 1 mmol/L sodium orthovanadate, 10 mmol/L NaF, 1 mmol/L PMSF, and 1 × Roche Complete Mini Protease Inhibitor Cocktail (Roche) from crypt cells of BALB/c and C57BL/6 mice, solid tumor of BALB/c mice, and normal and tumor colon organoid with different treatments. Cell lysates were subjected to SDS-PAGE, transferred to polyvinylidene difluoride membrane, and immunoblotted with monoclonal primary antibodies and appropriate secondary antibodies (1:10,000) Infrared-tagged 680RD Goat anti-Mouse IgG and 800CW Goat anti-Rabbit IgG (Li-COR). To detect the level of mono and polyubiquitinated proteins or peptides, we used mono- and polyubiquitinylated conjugated mAb (1:1,000; Enzo Life Sciences). To determine the p53 and p21 level and UPR signaling molecules in cell lysates, mouse mAb to p53 (1:500; catalog No. sc6243), p53 (human; 1:200; catalog No. sc126), p21(1:1,000; catalog No. sc397; Santa Cruz Biotechnology); p53 (acetyl K120; catalog No. ab78316, Abcam), Tip60 (phospho S86; catalog No. ab73207, Abcam); PERK (1:500; catalog No. C33E10), p-PERK (1:500; catalog No. T980), CHOP (1:1,000; catalog No. D46F1), PUMA (1:500; mouse catalog No. ab9643; Abcam; human catalog No. 4976; Cell Signaling Technology), BAK (mouse; 1:1,000; catalog No. 12105; Upstate Biotechnology, Inc.), BAX (mouse; 1:500; catalog No. 2772; Santa Cruz Biotechnology), and β-Actin (13E5) Rabbit mAb (1:1,000; catalog No. 4970; Cell Signaling Technology) was used.

Human colonic organoids grown in 24-well plates were used for immunoblot analysis. Each 24-well plate (number of organoids 15 ± 3/well) was used for an individual treatment group. For each experiment 30-μg protein was loaded for each sample.

Densitometric analysis of immunoreactive bands was performed using LI-COR Image Studio Lite, Version 5.2. Mean band density was plotted as a histogram after normalization of band density of each protein with band density of β-actin.

Full images of all immunoblots are shown in Supplementary Figs. S5 and S6.

Apoptosis in cells from solid tumors and cells from organoids following different treatments was determined by flow cytometry using Annexin V-fluoroisothiocyanate (FITC)/propidium iodide (PI) double staining (Bio-Vision; ref. 36). Briefly, the cells were collected and washed with binding buffer, then incubated in working solution (100-μL binding buffer with 1.0-μL Annexin V-FITC) for 15 minutes in the dark. PI was added just before flow cytometric analysis.

RNA was isolated from mouse intestinal epithelial cells, cells from mouse solid tumor, and human malignant colon organoid cells using RNeasy Mini Kit (Qiagen). Isolated RNA was subjected to cDNA synthesis using the SuperScriptTM First-Strand Synthesis System (Invitrogen). RT-PCR was performed in a QuantStudio 7 Flex Real-Time PCR Machine (Thermo Fisher Scientific) using the ABsolute QPCR SYBER Green Mix (ABgene) according to the standard ABgene protocol. To check for primer amplification specificity, a melting curve was generated at the end of the PCR and different samples containing the same primer pair showed matching amplicon melting temperatures. Primers used for RT-PCR were as follows: XBP1: forward, 5TGG CCG GGT CTG CTG AGT CCG′ 3′; reverse, 5′ ATC CAT GGG GAG ATG TTC TGG 3′; spliced form of XBP1: forward, 5′CTG AGT CCG AAT CAG GTG CAG3′; reverse, 5′ATC CAT GGG GAG ATG TTC TGG3′; CHOP: forward, 5′ ACC AAG GGA GAA CCA GGA AAC G 3′; reverse, 5′ TCA CCA TTC GGT CAA TCA GAG C 3′; GRP78: forward, 5′TTC CAA GGA ACA CTG TGG TG 3′; reverse, CAA AAG TTC CCA GGA GGT GA 3′; ATF4: forward, 5′GTT CTC CAG CGA CAA GGC TA 3′; reverse, 5′ATC CTG CTT GCT GTT GTT GG 3′; PMAIP1 (NOXA): forward, 5′GAG TGC ACC GGA CAT AAC TG3′; reverse, 5′CTCGTCCTTCAAGTCTGCTG3′; BBC3 (PUMA): forward, 5′GTGTGGAGGAGGAGGAGTG3; reverse, 5′ TCGATGCTGCTCTTCTTGTC3′; BIM: forward, 5′GGCCCCTACCTCCCTACA3′; reverse, 5′ GGGGTTTGTGTTGATTTGTCA3′; BAK forward, 5′ACGCTATGACTCAGAGTTCC3′; reverse, 5′ CTTCGTACCACA-AACTGGCC3′; BAX forward, 5′ATGGACGGGTCCGGGGAGCA3′; reverse, 5′CCCAGTTGAAGTTGCCGTCA3′; GAPDH: forward, 5′ TCA GTT GTA GGC AAG CTG CGA CGT 3′; reverse, 5′ AAG CCA GAG GCT GGT ACC TAG AAC 3′. The internal control gene GAPDH was amplified simultaneously in a separate reaction. Threshold cycle number (CT) of triplicate reactions was determined using the ABI-SDS software and the mean CT of triplicate reactions was determined. The levels of expression of the genes of interest were normalized to GAPDH using the formula 2^−ΔDCT, where −ΔDCT = ΔCT (sample) − ΔCT (control) and ΔCT is the CT of the target gene subtracted from the CT of the housekeeping gene GAPDH.

Data from the mouse survival/mortality studies were analyzed by Kaplan-Meier statistics as a function of radiation dose using Graphpad Prism 6.0 software for Mac (25). After genotyping, mice were sorted randomly to each experimental and control group. For mouse intestinal histopathologic analysis, jejunal sampling regions were chosen at random for digital acquisition for quantitation. Evaluation of digital image data was performed blinded as to treatment. A two-sided Student t test was used to determine significant differences between experimental cohorts (P < 0.05) with representative SEM.

Sensitivity of the intestinal epithelium to radiation is always a concern when using any antineoplastic agent as a radiosensitizer for abdominal radiotherapy. Therefore, we first examined whether auranofin alone or in combination with radiation induced GI toxicity. In vitro results in IEC6 intestinal epithelial cells demonstrated that auranofin alone does not have any effect on cell survival (Supplementary Fig. S1A). Moreover, IEC6 cells became more resistant to radiation with auranofin pretreatment (Supplementary Fig. S1C and S1D). Auranofin treatment, however, reduced the viability of CT26 colon cancer cells (Supplementary Fig. S1A). Clonogenic assay demonstrated that auranofin (2 μmol/L) enhances the radiosensitivity of CT26 cells (Supplementary Fig. S1B). Therefore, the in vitro data indicate that auranofin may be used as a radiosensitizer without promoting toxicity in nonmalignant tissue.

Treatment with auranofin in WT C57BL/6 male mice did not show any changes in animal health and overall survival. We examined the protective role of auranofin against radiation-induced intestinal injury in mice. Mice exposed to 14.25 Gy of AIR died within 10–15 days of radiation with characteristic signs and symptoms of RIGS, including diarrhea, black stools, and weight loss. In contrast, animals that received auranofin treatment (three times, alternate days, 10 mg/kg i.p.) prior to AIR (Fig. 1A and B) had well-formed stools, maintained body weight (Fig. 1D), and demonstrated 100% survival beyond 20 days post AIR (P < 0.003, log-rank (Mantel–Cox) test; n = 10/group; Fig. 1C). This experiment was also repeated in C57BL/6 female mice. Female mice receiving auranofin treatment (3 times, alternate days, 10 mg/kg i.p.) prior to AIR demonstrated 90% survival beyond 20 days post AIR (P < 0.0003, log-rank (Mantel–Cox) test; n = 10/group; Supplementary Fig. S2A). Mice rescued from RIGS by auranofin treatment were observed beyond 30 days postradiation. No behavioral changes were noticed, with normal food and water intake.

Histopathologic analysis of jejunal sections collected 96 hours post AIR showed crypt depletion and villous denudation in untreated WT mice exposed to AIR (Fig. 1E; ref. 25). Treatment with auranofin improved overall crypt villus architecture with an increase in crypt depth (P < 0.001; Fig. 1F), number of crypts (P < 0.005; Fig. 1G), and preserved villous length (P < 0.001; Fig. 1H) following irradiation. Because dextran is unable to cross the GI epithelium unless it is compromised, detection of dextran in the blood is a good indicator of epithelial damage. Blood FITC-dextran levels were measured at 4 hours after gavage (25). Exposure to AIR significantly increased the serum dextran level compared with unirradiated control, suggesting increase in epithelial permeability. Treatment with auranofin in irradiated mice significantly reduced the FITC-dextran uptake into the blood (P < 0.006; Fig. 1I). These data indicate restitution of intestinal epithelial integrity by auranofin treatment.

The intestinal epithelium in mice deficient in p53 was more sensitive to higher radiation doses, causing lethal GI syndrome. The presence of p53 in crypt epithelial cells promotes reversible cell-cycle arrest after irradiation. In p53-deficient epithelium, however, dysregulated cell proliferation following irradiation results in accelerated death of damaged cells followed by rapid destruction of villi and accelerated lethality. Super p53 mice that contain one extra copy of the WT p53 allele, for a total of 3 copies of the p53 genes, demonstrated resistance to GI syndrome compared with WT mice (37). Super p53; p21^−/− and WT p53; p21^−/− did not show any protection from GI syndrome (7). p21 is a cell survival protein, which arrests the cell cycle for DNA repair by a p53-dependent pathway. It has been reported that auranofin induces p21 expression with downregulation of Cdk2 and Cdk4 (38). We examined the role of p53 in the radioprotective effect of auranofin. Kaplan–Meier survival analysis demonstrated that auranofin could not rescue p53 knockout (KO) mice from radiation-induced toxicity (Fig. 1C and D). Histopathologic analysis demonstrated villi denudation (Fig. 1E and H), loss of crypt depth (Fig. 1E and F), and number of crypts (Fig. 1E and G) resulting in an increase in mucosal permeability in these irradiated mice, with/without auranofin treatment (Fig. 1I).

The radioprotective effect of auranofin was also examined in a fractionated AIR model (39). Mice receiving auranofin (10 mg/kg i.p.) prior to fractionated AIR exposure (6 Gy × 5 fractions; Fig. 1J and K) demonstrated significant improvement in survival compared with AIR controls (P < 0.005, log-rank (Mantel–Cox) test; n = 10/group).

To understand the effect of auranofin on crypt cell proliferation, we performed Ki67 staining in sections of jejunum from C57Bl6 mice. Exposure to WBI demonstrated a strong presence of Ki67⁺ crypt epithelial cells within the first 4 hours. A rapid decrease in the number of Ki67⁺ cells, however, was observed within 72 hours (Fig. 2A and B). Auranofin pretreatment (10 mg/kg i.p.) reduced cell proliferation prior to radiation, which gradually recovers to a higher proliferation level within 3.5 to 4 days post irradiation (Fig. 2A and B) and thereby serves to repair the crypt villus structure in intestinal epithelium (Fig. 2C–E). Auranofin treatment alone also reduced proliferation of unirradiated crypt cells, which then gradually recover following the auranofin treatment (Fig. 2B). γH2AX staining demonstrated significant reduction in DNA damage accumulation at 4–24 hours post irradiation with auranofin treatment compared with irradiated control (Fig. 2F and G). Moreover, TUNEL staining in jejunal sections at 4–24 hours post irradiation demonstrated significantly less apoptosis in the auranofin-treated group compared with irradiated controls (Fig. 2H and I). Auranofin also reduces radiation-induced mitotic catastrophe (multinucleated structures with multipolar spindles; ref. 40) in mouse crypt cells (Fig. 2J and K). These results suggest that auranofin-mediated blocking of cell proliferation prior to irradiation inhibits DNA damage and reduce apoptosis within hours after irradiation, which then promotes epithelial regeneration within 3–4 days post irradiation.

We have examined the involvement of p53 and p21 in auranofin-mediated regeneration of crypt epithelium in mice exposed to WBI. Histopathologic analysis at 96 hours postirradiation clearly demonstrated that auranofin could not inhibit radiation-induced intestinal epithelial damage (Supplementary Fig. S2B) in p53 and p21 KO mice. Unlike WT mice, auranofin pretreatment failed to halt radiation-induced crypt cell proliferation in p53 and p21 KO mice at 4 hours post irradiation (Supplementary Fig. S2B and S2C) resulting in mitotic catastrophe and significant damage to the intestinal epithelium (Supplementary Fig. S2B).

Cell fate after injury/stress depends on a balance between p53 downstream cell-cycle arrest and proapoptotic signaling (41–43). p53 activates downstream effector genes, such as PUMA and BAX for apoptosis, or a gene expressing cyclin/cdk inhibitor p21 for cell-cycle arrest. In response to radiation-induced DNA damage, acetylation of p53 at k120 switches on the proapoptotic pathway in intestinal epithelium (44). A previous report has shown that pharmacologic inhibition of p53k120 acetylation blocked radiation-induced apoptosis and PUMA induction without affecting induction of p21 expression. In the mouse model of abdominal irradiation, we have observed upregulation of p53k120 acetylation (Fig. 2L and M) resulting in induction of PUMA, BAX, and BAK expression (Fig. 2L and M). Pretreatment with auranofin did not alter the stabilization of p53 but significantly reduced radiation-induced acetylation of p53k120 and expression of PUMA, BAX, and BAK (Fig. 2L and M). Auranofin, a potent DUB inhibitor, stabilized p53 by inhibition of HAUSP DUB (Supplementary Fig. S3A; ref. 14). Absence of HAUSP promoted the ubiquitinated form and thereby proteosomal degradation of MDM2, a negative regulator of p53 levels (14).

Acetylation of p53k120 is mediated by Tip60 acetylatransferase (43). Activity of Tip60 acetyltransferase is modulated by ubiquitination (45). This study demonstrated that pretreatment with auranofin promoted the ubiquitinated inactive form Tip60 (Fig. 2N and O) and thereby reduced the acetylated form of p53k120. The deacetylated form of p53 shifted the balance from a proapoptotic pathway and induced p21 expression (Fig. 2L and M). Involvement of Tip60 in acetylation of p53 at k120 and thereby activation of the intrinsic apoptotic pathway was further confirmed by siRNA-mediated depletion of Tip60 in IEC6 intestinal epithelial cells. In irradiated IEC6 cells, absence of Tip60 inhibited the acetylation of p53k120 and activation of the apoptotic pathway without affecting p21 expression (Supplementary Fig. S3B and S3C). Treatment of IEC6 cells with auranofin prior to irradiation also inhibited acetylation of p53k120 and the downstream intrinsic apoptotic pathway and induced p21 expression (Supplementary Fig. S3B and S3C). Previous studies by other groups have also shown that pharmacologic inhibition of Tip60 inhibits acetylation of p53k120 and rescues intestinal stem cells from radiation-induced apoptosis (44). Therefore, these observations, including our current study, clearly suggest that modulation of p53k120 acetylation shifts the balance from a proapoptotic pathway and induces p21 expression to promote survival of intestinal epithelial cells following irradiation.

To determine the role of auranofin in radiation-induced normal tissue toxicity in human colonic epithelium, we turned to an ex vivo colonic organoid culture system developed from surgical specimens from human colon (24). The effect of auranofin has been examined in six different malignant and nonmalignant paired organoids. We have observed interpersonal radiosensitivity differences in both normal and malignant tissue–derived organoids (Supplementary Fig. S4A and S4B). Following radiation exposure, organoids lose their budding crypt structure in a dose-dependent manner (Fig. 3A and B). The radioprotective role of auranofin was examined in nonmalignant colon–derived organoids. Treatment with auranofin (2 μmol/L) prior to radiation (6 Gy) rescued these organoids from radiation toxicity (Fig. 3A) and restored the number of budding crypts to the unirradiated level as represented by budding crypt/total crypt ratio (Fig. 3A and B). No significant difference was noted in organoid morphology or growth between unirradiated untreated organoids and auranofin-treated unirradiated organoids (Fig. 3A and B). Auranofin also promoted cell proliferation of irradiated organoid cells (Fig. 3A–D), ATP level (Fig. 3C), and inhibited apoptosis (Fig. 3A–E) as demonstrated by Ki67 staining, ATP level analysis, and cleaved caspase staining, respectively. To analyze the mechanism of action of radioprotective role of auranofin, three most radiosensitive organoids (n = 3) were further examined.

To determine the involvement of the p53/p21 pathway in the radioprotective role of auranofin, human colonic organoids were first treated with siRNAs targeted to inhibit p53 expression and then treated with auranofin and radiation. A pooled siRNA approach was used (including two to three siRNAs) to inhibit the expression of the gene of interest. Irradiated organoids pretreated with p53 siRNA did not respond to auranofin and demonstrated complete loss of budding crypt–like structure upon irradiation (Fig. 3F and G) with significant reduction in ATP level (Fig. 3H). These results indicate the involvement of p53 in auranofin-mediated radioprotection of human colonic organoids. siRNA-mediated inhibition of p21 also inhibited the auranofin-mediated protection of organoids against radiation (Fig. 3F–H) suggesting involvement of p21 in p53-mediated radioprotective effect of auranofin. To further validate these results, expression of both p53 and p21 was inhibited in the same organoids, which also showed inhibition of auranofin-mediated radioprotection (Fig. 3F–H). We performed immunoblot analysis to determine the transfection efficiency of p53 and p21 siRNA in organoids. Treatment with p53 or p21 siRNAs completely suppressed these target proteins, respectively. Suppression of p21 expression did not show any effect on p53 expression (Fig. 3K). However, p53 suppression partially reduced the p21 expression suggesting that p21 is downstream of p53 and therefore regulating the p53-mediated cell survival.

Immunoblot analysis showed significant upregulation of p53 levels in irradiated organoids with/without auranofin treatment (Fig. 3I and J). Irradiated organoids in the absence of auranofin treatment demonstrated significant upregulation of the acetylated form of p53 (Fig. 3I and J) suggesting activation of a p53 downstream apoptosis pathway. However, in response to auranofin treatment, level of acetylated p53 was significantly downregulated (Fig. 3I and J) with the upregulation of p21 expression (Fig. 3I and J). Involvement of an intrinsic apoptosis pathway in radiation-induced apoptosis in organoids was examined by siRNA-mediated depletion of PUMA. Depletion of PUMA made those organoids more radioresistant. The radioprotective role of auranofin was also maintained in these organoids (Supplementary Fig. S3D–S3F).

Therefore, auranofin inhibits a radiation-induced p53 downstream apoptosis pathway and promotes p53/p21 pathway–mediated cell survival.

To determine the role of auranofin on the radiation response of mouse tumors, we developed CT26 colon tumors on the abdominal flank region of BALB/c mice by subcutaneous injection of 2 × 10⁶ CT26 cells (22, 30). Mice with palpable subcutaneous abdominal tumors were treated with auranofin (10 mg/kg i.p.) followed by exposure to fractionated abdominal irradiation (AIR; 4-Gy/fraction; two fractions/week for 2 weeks; Fig. 4A). Significant tumor growth delay was observed with combination treatment of AUR+AIR compared with controls (P < 0.0006). The median time to achieve tumor size 2,000 mm³ in control, AIR, auranofin, and AUR+AIR was 22 days, 29 days, 50 days, and 68 days, respectively.

CT26 colon tumor cells do not have the Cdnk2a gene (46) responsible for expressing p19ARF. p19ARF functions as a tumor suppressor by inducing p53 stabilization (47). We have observed that expression of p53 and p21 was not altered by radiation and/or auranofin treatment (Fig. 4D). These results suggest that the absence of Cdnk2a made p53 nonresponsive to radiation/auranofin treatment and therefore, it may not play a significant role in the tumoricidal effect of auranofin.

Radiotherapy with a single large dose is sometimes preferred over fractionated radiation because of better treatment outcome and low risk of tumor recurrence. The high radiosensitivity of intestinal epithelium restricts the use of a tumoricidal large single-radiation dose against abdominal malignancies. In this study, we have examined whether auranofin could be applied with a large single dose of AIR (12.5 Gy) against colon tumors. Mice with palpable subcutaneous abdominal CT26 colon tumor were treated with/without auranofin (10 mg/kg, i.p.), followed by exposure to lethal dose abdominal irradiation (12.5-Gy single fraction; Fig. 4E). Mice exposed to AIR without auranofin pretreatment all died within 15 days of exposure with symptoms of RIGS including loss of body weight (Fig. 4G), diarrhea etc. However, auranofin treatment prior to radiation exposure rescued these mice from radiation toxicity along with significant tumor growth retardation compared with unirradiated controls (Fig. 4F and H). The tumor growth curve of untreated AIR mice was not complete due to mortality in all mice within 15 days postexposure. Mice rescued by auranofin continued to survive beyond 30 days post AIR without any sign or symptoms of RIGS.

Therefore, these results demonstrate that administration of auranofin prior to irradiation enhances the radiation response of the colon tumors while providing radioprotection to the normal tissues, including small intestine, thus increasing the therapeutic ratio of abdominal irradiation.

Auranofin inhibits 19S proteasome–associated DUBs to inhibit proteosomal degradation and thereby increases cytotoxicity in malignant tissue (18). To examine the involvement of auranofin-mediated inhibition of DUBs in radiosensitization of tumor tissue, DUB activity was measured in the CT26 tumor tissue lysate by ELISA. A significantly lower DUB activity was found in the auranofin-treated tumor exposed to radiation compared with irradiated control tumor or untreated tumor (P < 0.007 and P < 0.002; Fig. 5A). Treatment with auranofin without radiation also significantly reduced the DUB activity in tumor tissue compared with irradiated or untreated control (P < 0.005 and P < 0.002; Fig. 5A). Next, we determined the ubiquitinated protein level in tumor tissue (18). Combination treatment of auranofin and radiation significantly increased the ubiquitinated protein level in tumor tissue compared with irradiated or untreated controls (P < 0.005 and P < 0.001, respectively; Fig. 5B and C). Treatment with auranofin alone also increased the ubiquitinated protein level compared with untreated or irradiated control (P < 0.002 and P < 0.007; Fig. 5B and C). Inhibition of proteosomal degradation induces unfolded and misfolded protein burden within cells resulting in initiation of UPR with the induction of ER stress (48). Three major UPR signaling pathways, inositol-requiring enzyme 1α (IRE1α), activating transcription factor 4 (ATF4), and protein kinase R (PKR)-like endoplasmic reticulum kinase (PERK; refs. 22, 48), were examined in auranofin-treated tumor tissue. IRE1α, upon activation, splices XBP1 into the active form, producing a potent transcription factor that translocates to the nucleus and activates several UPR target gene expressions, including HSPs. ATF4 is a downstream target of PERK that can activate an ER stress downstream proapoptotic pathway with induction of the proapoptotic protein C/EBP homologous protein (CHOP; ref. 49). Auranofin treatment of tumor prior to irradiation upregulated sXBP1, GRP78, ATF4, and CHOP mRNA levels compared with irradiated or untreated controls (Fig. 5D). Immunoblot analysis also demonstrated auranofin-mediated upregulation of p-PERK (activated for PERK) and CHOP protein expression, which further increased in response to combination treatment (Fig. 5E and F). Because PERK activation attenuates protein synthesis in response to ER stress through the phosphorylation of translation initiation factor eIF2α at serine 51, we determined the levels of phosphorylated eIF2α. IHC of peIF2α demonstrated higher peIF2α-positive tumor cells in auranofin and the combination treatment groups (Fig. 5I). Consistent with these findings, combination treatment of AUR+AIR also increased mRNA levels of UPR downstream proapoptotic genes PUMA, PMAIP1, BAK, BAX, and BIM (Supplementary Table S1) and promoted apoptotic cell death in CT26 tumor cells compared with irradiated controls (Fig. 5G–I). The apoptotic cell death was determined in the tumor tissue by detecting the Annexin V/PI–positive cells using flow cytometry (Fig. 5G and H) and by using TUNEL staining in tumor sections (Fig. 5I). Percent of Annexin V–positive CT26 tumor cells was higher in the AUR+AIR group compared with irradiated (P < 0.003) or auranofin-treated (P < 0.007) or untreated controls (P < 0.001; Fig. 5G and H). TUNEL staining also demonstrated the presence of more apoptotic tumor cells in the combination treatment group (AUR+AIR) compared with irradiated or auranofin-treated groups (Fig. 5I). These results indicated that auranofin treatment radiosensitizes the tumor with an increase in apoptotic cell death through activation of a proapoptotic ER stress signaling pathway.

To determine the effect of auranofin treatment in human colonic malignant tissue, we developed primary ex vivo organoid cultures from surgical specimens of malignant colon. For these experiments, we used surgical specimens from the same patients considered for the collection of nonmalignant tissue. Irradiation of these organoids resulted in loss of budding crypt structures in a dose-dependent manner (Fig. 6A and B). Similar to their nonmalignant counterparts, malignant tissue–derived organoids also demonstrated interpersonal differences in radiosensitivity (Supplementary Fig. S4C). However, radiation-induced damage in these organoids was relatively less compared with nonmalignant organoids (Supplementary Fig. S4A) indicating the radioresistant nature of colonic malignant tissue. Pretreatment with auranofin radiosensitizes this malignant tissue–derived organoids resulting in significant loss of budding crypt–like structures with the reduction in budding crypt ratio compared with irradiated controls or untreated controls (Fig. 6A and B; Supplementary Fig. S4C and S4D). Ki67⁺ proliferative cells (Fig. 6A and D) and ATP level (Fig. 6A and C) were also significantly decreased in auranofin pretreated, irradiated organoids compared with irradiated controls.

The three most radioresistant organoids were further examined for mechanism of action of auranofin-mediated radioprotection. A significantly lower DUB activity was found in the auranofin-treated human tumor colon organoids exposed to irradiation compared with irradiated or untreated control tumor colon organoids (P < 0.008 and P < 0.002, respectively; Fig. 6F). Treatment with auranofin without irradiation also significantly reduced DUB activity in human tumor colon organoid compared with irradiated or untreated control (P < 0.005 and P < 0.002, respectively; Fig. 6F).

Next, we determined the effect of auranofin on UPR in these malignant colon–derived organoids. Pretreatment of auranofin in irradiated organoids upregulated sXBP1, GRP78, ATF4, and CHOP mRNA levels compared with irradiated organoids (Fig. 6G). Organoids pretreated with siRNAs targeted to inhibit PERK expression did not show radiosensitization with auranofin treatment (Fig. 6H–J). Absence of PERK in siRNA-treated organoids was confirmed by immunoblot (Fig. 6K). However, PERK siRNA treatment in nonmalignant colonic organoid did not reverse the radioprotective role of auranofin (Supplementary Fig. S3G and S3H) suggesting that involvement of PERK is restricted to auranofin-mediated radiosensitization of malignant organoids. Consistent with these findings, auranofin pretreatment also increased the percent of Annexin-V FITC apoptotic cells in these irradiated organoids compared with irradiated (P < 0.002) or auranofin only treatment (P < 0.005; Fig. 6L and M). Similar effect was also observed in organoids stained with cleaved caspase (Fig. 6A and E). These findings clearly indicate that auranofin pretreatment radiosensitizes human malignant colonic organoids by inducing a UPR downstream proapoptotic signaling pathway.

This study demonstrates that repurposing of the antirheumatoid drug, auranofin, radioprotects the normal intestinal epithelium but radiosensitizes malignant tissue. In a model of murine CT26 colorectal tumors treated with whole-abdominal irradiation, pretreatment with auranofin protected the mice from lethal RIGS while improving tumor growth suppression. Moreover, colonic organoids developed from surgical specimens of human colon demonstrated similar effects in response to auranofin treatment. The radioprotective ability of auranofin is restricted to organoids developed from nonmalignant specimens of human colon. Organoids developed from malignant tissue from the same patient were sensitized by auranofin treatment. This is the first use of paired normal and malignant tissue from the same patient to examine the efficacy of a pharmacologic agent for improvement of radiotherapy. This opposite effect of auranofin in irradiated normal and malignant tissue from the same patient is promising for its further use in clinical settings in combination with radiotherapy.

p53-mediated cell-cycle arrest and DNA double-strand break repair is critical for radioprotection in the intestinal epithelium. Previous reports have shown that selective protection of normal tissue from radiation toxicity could be achieved by pharmacologic modulation of cell-cycle arrest (50) using inhibitors of CDKs. But a CDK inhibitor could not prevent radiation injury in p53 KO mice. In this study, auranofin treatment activated the p53/p21 pathway for DNA check point inhibition, resulting in a reduction of radiation-induced damage in the normal intestinal epithelium. However, combination treatment of auranofin with radiation induced synthetic lethality in colon tumor through activation of ER stress and ER stress–associated proapoptotic pathway to induce apoptosis in cancer cells.

It has been reported that p21/p53–deficient mice are more sensitive to RIGS (9). p53-deficient crypt epithelial cells continue to proliferate post irradiation causing mitotic catastrophe within 72–96 hours after irradiation resulting in denudation of villi within 3–5 days post irradiation. However, the presence of p53 induced reversible growth arrest in crypt epithelial cells within 72 hours post irradiation and thereby prevented mitotic catastrophe. After 72 hours post irradiation, these WT p53 crypt epithelial cells are proliferating to repair the damage caused by irradiation. p21 plays a critical role in allowing p53 to protect mice from the GI syndrome (7). In response to radiation, the GI epithelium of p21 null mice also undergoes aberrant cell-cycle progression with persistent DNA damage that leads to cell death via mitotic catastrophe. Super p53 mice, which are very resistant to RIGS, become very sensitive with the deletion of the p21 gene (7). In this study, auranofin pretreatment of WT mice demonstrated initial arrest of crypt cell proliferation, which gradually recovered to a higher level resulting in epithelial regeneration within 96 hours post irradiation. However, p53 or p21 KO mice as well as siRNA-mediated inhibition of p53 or p21 in human nonmalignant colonic organoids completely abrogated the radioprotective role of auranofin. P53 can also induce apoptosis. In response to radiation injury in the intestine, p53-mediated apoptosis depends on Tip60 acetylatransferase–mediated acetylation of p53k120 (44). Our study in irradiated mouse intestinal epithelium and human nonmalignant colon organoids demonstrates clear evidence that auranofin pretreatment inhibits acetylation of p53k120, which reduces radiation-induced apoptosis and activates p21 downstream cell-cycle arrest. Therefore, auranofin reduces radiation-induced mitotic catastrophe and promote intestinal epithelial regeneration by modulation of a p53/p21 pathway.

Auranofin inhibits 19S proteasome–associated DUB to block proteosomal degradation (18). In this study, we have shown that auranofin treatment reduced DUB activity and thereby inhibited deubiquitination in irradiated tumor tissue resulting in a significant increase in ubiquitinated protein level. Deubiquitination is a critical step for proteosomal degradation of unfolded and misfolded proteins in cancer cells. Therefore, DUB inhibition increases the unfolded or misfolded protein burden resulting in a UPR. Increase in unfolded or misfolded protein burden beyond tolerance level switches the cytoprotective effect of UPR to the cytotoxic effect by inducing UPR-associated apoptosis. Our study demonstrated that the combination of auranofin and radiation induced synthetic lethality in cancer cells by tipping the balance from the cytoprotective arm of UPR to the PERK-CHOP–associated apoptotic arm, resulting in significant reduction of colon tumor growth in mice as well as growth inhibition in human malignant colonic organoids. However, we did not observe any changes in p53/p21 expression in malignant tissue in response to auranofin or radiation suggesting that, unlike normal tissue, the p53/p21 pathway is not playing a central role in the mechanism of action of auranofin.

Cancer cells are more sensitive to proteasome inhibition than are normal cells. In our studies, auranofin-mediated proteosomal inhibition increased the radiosensitivity of cancer cells without altering the radiosensitivity of nonmalignant cells. Because of cancer cell–specific cytotoxicity, proteosomal inhibition is being developed as a major therapeutic target to improve the efficacy of radiotherapy. Moreover, our current data demonstrate that auranofin confers radioprotection to the normal GI epithelium via activation of the p53/p21 pathway while radiosensitizing tumor cells through proteosomal inhibition. Considering the high radiosensitivity of the gastrointestinal epithelium, a pharmacologic agent like auranofin should be of great importance in future clinical trials using it in combination therapy with abdominal radiotherapy.

No potential conflicts of interest were disclosed.

Conception and design: D. Nag, P. Bhanja, R. Riha, C. Lominska, S. Saha

Development of methodology: D. Nag, R. Riha, G. Sanchez-Guerrero, S. Saha

Acquisition of data (provided animals, provided facilities, etc.): P. Bhanja, R. Riha, D. Nag

Analysis and interpretation of data (e.g., statistical analysis, biostatistics, computational analysis): D. Nag, P. Bhanja, R. Riha, B.F. Kimler

Writing, review, and/or revision of the manuscript: D. Nag, P. Bhanja, R. Riha, B.F. Kimler, T.T. Tsue, C. Lominska, S. Saha

Administrative, technical, or material support (i.e., reporting or organizing data, constructing databases): P. Bhanja, T.T. Tsue, C. Lominska, S. Saha

Study supervision: P. Bhanja, S. Saha

The authors acknowledge the Biospecimen Repository Core Facility, Histopathology Core Facility, and Confocal Microscopy Core Facility at KUMC for their help and support. This work was supported by grants from 1U01AI138323 (to S. Saha), K01DK096032 (to S. Saha), ACS IRG Pilot IRG-15-194-07 (B.F. Kimler.), KUCC support grant P20 168524 (S. Saha), KUCC pilot P20 168524 (to S. Saha).

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.