Defining the mode of action of cisplatin combined with a phosphoramidate modification of gemcitabine

doi:10.21203/rs.3.rs-4344105/v1

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Defining the mode of action of cisplatin combined with a phosphoramidate modification of gemcitabine

https://doi.org/10.21203/rs.3.rs-4344105/v1

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The combination of gemcitabine with platinum agents is a widely used chemotherapy regimen for a number of tumour types. Gemcitabine plus cisplatin remains the current therapeutic choice for biliary tract cancer. Gemcitabine is associated with multiple cellular drug resistance mechanisms and other limitations and has therefore declined in use. NUC-1031 is a phosphorylated form of gemcitabine, protected by the addition of a phosphoramidate moiety, developed to circumvent the key limitations and generate high levels of the cytotoxic metabolite, dFdCTP. The rationale for combination of gemcitabine and cisplatin is determined by in vitro cytotoxicity. This, however, does not offer an explanation of how these drugs lead to cell death. In this study we investigate the mechanism of action for NUC-1031 combined with cisplatin as a rationale for treatment. NUC-1031 is metabolised to dFdCTP, detectable up to 72 hours post-treatment and incorporated into DNA, to stall the cell cycle and cause DNA damage in biliary tract and ovarian cancer cell lines. In combination with cisplatin, DNA damage was increased and occurred earlier compared to monotherapy. The damage associated with NUC-1031 may be potentiated by a second mechanism, via binding the RRM1 subunit of ribonucleotide reductase and perturbing the nucleotide pools; however, this may be mitigated by increased RRM1 expression. The implication of this was investigated in case studies from a Phase I clinical trial to observe whether baseline RRM1 expression in tumour tissue at time of diagnosis correlates with patient survival.

The combination of gemcitabine with platinum agents is a widely used chemotherapy regimen for a number of tumour types and remains a standard of care in biliary tract cancers (BTC). Biliary tract cancers are uncommon malignancies, affecting less than 3 in 100,000 people in Western countries, often present late and progress rapidly (Khan et al., 2002; Kirstein & Vogel, 2016; Baria et al., 2022). The global standard of care for the first-line treatment of BTC is a combination of gemcitabine and cisplatin (Hyung et al., 2019; Valle et al., 2010; Valle et al., 2016). However, median overall survival on this regimen is very modest at less than 1 year. Ovarian cancers affect 11 in 100,000 females in Europe (6). Gemcitabine and platinum agents were previously one of the chemotherapeutic options for ovarian cancer. However, gemcitabine is associated with acquired resistance and patient relapse (7); therefore, current options include combinations of platinum agents with paclitaxel or PARP inhibitors (8, 9). Survival times remain low with gemcitabine and cisplatin; 11.5- and 15.8-months overall survival for biliary tract and ovarian cancer patients, respectively (Okusaka et al., 2010; Safra et al., 2014; Valle et al., 2010; Valle et al., 2016). Thus, improved treatment options are urgently needed.

In order to assess whether a replacement for gemcitabine would be clinically advantageous, the mode of action of the drug combinations should be understood. Current in vitro screening is generally limited to determining cell death, used to calculate a combination index as a measure of drug synergy (Sakamoto et al., 2019; Tang et al., 2013). This does not encompass drug mechanisms individually or in combination, which might provide more detailed rationale for clinical studies.

Gemcitabine (2', 2'-difluoro 2'deoxycytidine; dFdC) is a fluorinated deoxycytidine analogue approved for treatment of advanced pancreatic, ovarian, bladder, breast, and non-small lung cell carcinoma (14, 15). Anti-proliferative effects occur via incorporation of its triphosphate form, di-fluoro-deoxycytidine triphosphate (dFdCTP), into elongating DNA strands, in place of endogenous non-fluorinated deoxycytidine triphosphate (dCTP). This occludes extension of the growing chain, known as masked chain termination (16, 17). A second cytotoxic mechanism is exerted via deoxycytidine diphosphate (dFdCDP), which inhibits the enzyme ribonucleotide reductase (RNR), required to convert of nucleotide diphosphates (NDPs) to their deoxy- form (dNDPs) (18). The anti-cancer activity of gemcitabine is limited by shortcomings associated with breakdown, transport, and activation. Gemcitabine is metabolised by cytidine deaminase (CDA) extracellularly and intracellularly, rendering it inactive. The anti-proliferative effects of gemcitabine are dependent on active cellular import via nucleoside transporters hENT and hCNT, and intracellular activation by the enzyme deoxycytidine kinase (dCK). Patients often show an initial response to gemcitabine but develop resistance due to intrinsic or acquired mechanisms, including downregulation of hENT and hCNT and regulated inhibition of dCK (19), Similarly, decreased dCK expression has been observed in colon and gastric cancers and cholangiocarcinoma (20, 21).

Cisplatin (cis-diamminedichloroplatinum (II)) is used to treat several solid tumours including lung, bladder, cervical, ovarian, head and neck, nasopharyngeal and oesophageal. Cisplatin forms intra-strand and inter-strand crosslinks between DNA bases. This interferes with DNA polymerases on the replication forks and forms bulky lesions in G1 and G2 phases, interrupting transcription (Clauson et al., 2013; Guainazzi & Schärer, 2010; Wagner & Karnitz, 2009). These lesions can be repaired by nucleotide excision repair (NER); by the Fanconi Anaemia core complex (FA) for inter-strand crosslink (ICL) repair; or by homologous recombination (HR), during S and G2 phases in a BRCA-2 dependent manner (Duan et al., 2020; Hashimoto et al., 2016; Kim et al., 2008; Kiss et al., 2021; Wagner & Karnitz, 2009). Detection of DNA lesions and initiation of the associated repair pathways is co-ordinated with cell cycle progression and post-translational modifications of proteins involved in repair. This is governed by the DNA Damage Response (DDR) which initiates the phosphorylation cascade of multiple downstream cell cycle regulation and repair proteins to trigger cell cycle checkpoint arrests (McHugh & Sarkar, 2006; Chatterjee & Walker, 2017). Incorporation of dFdCTP into DNA during the repair process may disrupt these repair pathways. Thus, the combination of cisplatin and dFdCTP may stimulate an increased DNA damage response and impede cell cycle progression, resulting in synergistic effects that triggers apoptosis in cancer cells.

NUC-1031 is a phosphorylated form of gemcitabine, specifically designed to overcome the key limitations associated with gemcitabine and produce higher concentrations of the active anti-cancer metabolite dFdCTP inside the tumour cell. NUC-1031 is comprised of pre-activated gemcitabine monophosphate (dFdCMP) protected by a phosphoramidate moiety (a specific combination of aryl, ester and amino acid groups). The phosphoramidate moiety enables NUC-1031 to enter cancer cells independent of the hENT1 and hCNT1 transporters. Once NUC-1031 has entered the cell, the protective group is cleaved off and releases dFdCMP, obviating the need for the activating enzyme, dCK, which drives the rate-limiting phosphorylation of gemcitabine. dFdCMP is rapidly converted to dFdCDP and then dFdCTP. Moreover, NUC-1031 is not subject to breakdown by CDA, thus bypassing another key limitation (Slusarczyk et al., 2014).NUC-1031 has been investigated in clinical trials for ovarian (NCT03146663) and biliary tract cancers (NCT04163900)

We hypothesized that a mechanistic approach to deciding how to combine drugs would be more clinically relevant than in vitro IC₅₀ synergy models. Based on individual mechanisms of action, the combination of NUC-1031 and cisplatin is anticipated to increase DNA damage and affect cell cycle progression, resulting in greater cellular apoptosis compared to either agent alone.

NUC-1031 forms dFdCTP which is incorporated into DNA

The kinetics of NUC-1031 cellular uptake and activation was assessed by liquid chromatography mass spectrometry through analysing intracellular levels of the active metabolite dFdCTP and its incorporation into DNA (in the form of dFdC) in intrahepatic cholangiocarcinoma (HuCCT1) cells (Fig. 1. left) and ovarian cancer (A2780) cells (Fig. 1. right). HuCCT1 cells were treated for 24 hours, and A2780 for 2 hours at either IC₂₅ or IC₅₀ doses, based on cell cytotoxicity assays (Fig.S1). The drug was removed after either 24 or 2 hours and dFdCTP was detectable up to 96 hours post-treatment. In HuCCT1 cells, a peak of 220 nmol dFdCTP /10⁶ cells was observed 24 hours post-treatment, diminishing rapidly towards 48 hours and detectable up to 96 hours (Fig. 1A, left). The levels of detectable dFdCTP in A2780 cells (Fig. 1.A, right) increased from 44 nmol/10⁶ cells at 2 hours up to 350 mol/10⁶ cells at 48 hours post-treatment.

To determine the extent of incorporation of the active metabolite, DNA was extracted and hydrolysed to release deoxynucleotide monophosphates. A dephosphorylation step was performed due to concerns regarding stability of this form (Jansen et al., 2009); therefore, the nucleosides dFdC and dG were used as surrogates for the incorporation of the active metabolite (Fig. 1B). Levels of dFdC were normalised to dG as the complementary base for dC and dFdC (Bapiro et al., 2011). This was performed by simultaneous quantification of both analytes, a method independent of hydrolysis efficiency (Wickremsinhe et al., 2010). Incorporation into DNA ranged from 0.33 nmol dFdC/µmol dG at 24 hours to 0.9 nmol dFdC/µmol dG at 72 hours post-treatment in HuCCT1 cells (Fig. 1B, left). In A2780 cells, this ratio increased from 0.2 nmol dFdC/µmol dG at 6 hrs post treatment to a maximum of 0.6 nmol/µmol dG at 48 hours post-treatment (Fig. 1B, right). As incorporation of nucleotides into DNA is related to progression through S phase, we hypothesised that the prolonged incorporation of dFdCTP was related to the proportion of cells in this phase in the cell cycle.

NUC-1031 arrests cells in S phase of cell cycle and elicits a DNA damage response

The effect of incorporation of dFdCTP into DNA on cell cycle progression was measured by flow cytometry. Shifts in proportions of cells in different phases of the cell cycle were observed in response to NUC-1031 in both HuCCT1 and A2780 cells. NUC-1031 increased the proportion of cells in S phase in both cell lines (Fig. 2A); maximal proportions occurred at 48 hours in HuCCT1 cells and 24 hours in A2780 cells. Up to 48% and 42% of HuCCT1 cells treated with 500 nM and 1 µM, respectively, were in S phase, compared to 20% with DMSO at 48 hours (Fig. 2B). In A2780 cells, 30% and 47% of cells were in S phase, respectively, for equipotent doses at 24 hours, compared to 20% with DMSO, with decreases occurring in G0/G1 populations. This was followed by a return towards a normal cell cycle profile at 96 hours.

NUC-1031-induced DNA damage was quantified by use of fluorescence detection of double-strand and single-strand break responses in both HuCCT1 and A2780 cell lines, normalised to that of DMSO-treated cells over 96 hours post-treatment. The double-strand break response was assessed by the phosphorylation of histone variant H2AX (Ser139), (Fig. 3A); and the DNA damage transducer protein ATM (Ser367) (Fig. 3B). Single strand break exposure was measured by phosphorylation of Checkpoint kinase 1 (Chk1-Ser345) (Fig. 3C). An increase in phosphorylation of these markers, above endogenous levels, occurred from as early as 24 hours and up to 96 hours post-treatment in both HuCTT1 and A2780 cells, following cell cycle arrest.

A combination of NUC-1031 and cisplatin inhibits cell cycle progression and exhibits increased DNA damaging effects in vitro

The in vitro cytotoxicity of combinations of NUC-1031 and cisplatin was determined in HuCCT1 cells, treated with both drugs simultaneously for 24 hours (Fig. S3). The IC₅₀ determined at 96 hours post-treatment revealed additivity but no synergy was present across doses of NUC-1031 and cisplatin assessed using the Bliss and Loewe criteria (32, 33).

Cytotoxic synergy was not demonstrated by Bliss and Loewe models, although evidence of additivity was found. Since the mechanism of NUC-1031 and cisplatin acting individually has been described, we explored the DNA damage and cell cycle effects of both drugs in combination. Combinations of NUC-1031 at either IC₁₀ or IC₂₅ doses and IC₂₅ cisplatin (Fig. S1) were investigated in HuCCT1 cells.

To determine whether the addition of cisplatin potentiates NUC-1031 activity, fluorinated deoxycytidine (dFdC) in DNA was measured in HuCCT1 cells (Fig. 4). At 24 hours post-treatment, DNA incorporation was found to be similar between NUC-1031 single-agent and combination samples. DNA incorporation peaked at 48 hours (0.50 nmol/µmol dG and 0.53 nmol/µmol dG for NUC-1031 single-agent and NUC-1031 plus cisplatin combination, respectively), where incorporation of fluorinated deoxycytidine into DNA is independent of cisplatin despite cisplatin-induced cell cycle arrest. At the lower NUC-1031 dose DNA incorporation was higher in NUC-1031 single-agent samples compared to combination samples (0.28 nmol/µmol dG vs 0.15 nmol/µmol dG).

The DNA damage and cell cycle effects were investigated by flow cytometry in order to determine the mechanistic interaction at these doses. The proportion of cells in S phase increased with single-agent treatment, from 18% in DMSO-treated cells to 39% with NUC-1031 and 35% with cisplatin at 24 hours. The combination treatment did not alter the proportions of cell in S phase at any timepoint when compared to single-agent treatment (Fig. 5A). Single-agent cisplatin increased the proportion of cells in G2 phase, from 22–29% at 24 hours post-treatment and from 25–56% at 48 hours post-treatment (Fig. 5B). The addition of NUC-1031 to cisplatin further increased these proportions to 71% at 48 hours, whereas NUC-1031 alone had no effect on the proportion of cells in G2 phase at the doses tested. The proportion of cells in G1 phase was reduced at 48 hours (Fig. 5C), from 59% in DMSO-treated controls to 47% and 28% with single-agent NUC-1031 and cisplatin, respectively. The combination of NUC-1031 and cisplatin reduced this proportion down to 13% by 48 hours post-treatment, corresponding to the increase in S and G2 phases.

The single strand DNA damage response signalled by phosphorylation of Chk1 (Ser345) was increased by 1.4-fold with NUC-1031 but not cisplatin at 48 hours post-treatment (Fig. 5D), with further increase in response to the combination of NUC-1031 and cisplatin (2-fold increase). The double strand break response increased with combinations of drugs compared to either single agent at 48 hours post-treatment. ATM phosphorylation (Ser367) was similar with cisplatin alone and in combination (2.4 and 2.6-fold respectively) but remained elevated up to 72 hours post-treatment compared to either single agent (Fig. 5E). γH2AX (Ser139) intensity increased at 48 hours post-treatment, from 1.6-fold and 2.2-fold with either NUC-1031 or cisplatin alone, respectively, to 3.3-fold increase with the combination (Fig. 5F).

The Chk1 phosphorylation was found to be synergistic at 48 hours. The increase in double-strand break response, via p-ATM and γH2ax, appears additive. The predominant effect of cisplatin on the cell cycle at 48 hours cell cycle stalling occurs at G2/M phase, with additivity at higher doses of cisplatin and synergy with IC₅₀ doses of NUC-1031.

NUC-1031 binds Ribonucleotide Reductase subunit RRM1

An additional mechanism of NUC-1031, the inhibition of RNR via the RRM1 subunit, was assessed in HuCCT1 and A2780 cell lines. The RRM1 regulatory subunit of RNR contains a binding site for NDPs, to facilitate the conversion to dNTPs. Gemcitabine-derived dFdCDP has been shown to competitively bind to this site (34, 35). The binding of dFdCDP to RRM1 was assessed by the presence of a band at the 110 kDa position on Western blots, at equipotent doses of gemcitabine and NUC-1031, and equivalent to IC₅₀ (Fig. 6A). Bound RRM1 was apparent from 24 to 72 hours post-treatment with gemcitabine in both HuCCT1 and A2780 cell lines (Fig. 6B) and remained visible up to 96 hours post-treatment with NUC-1031. Up to 14.2% and 15.5% of total RRM1 was bound after 24-hour exposure to gemcitabine and NUC-1031, respectively, in HuCCT1 cells; and 18.8% and 22% in A2780 cells. The percentage of RRM1 bound by either gemcitabine of NUC-1031 was not significantly different at 72 or 96 hours (Two-way ANOVA, p > 0.05) in each condition (Fig. S4).

The total RRM1 protein expression was determined by addition of top and bottom band intensities (Fig. 6C). The expression increased at 24 hours in A2780 cells (gemcitabine: 1.28-fold, NUC-1031: 1.12-fold) and in HuCCT1 cells (gemcitabine: 1.27-fold, NUC-1031: 1.70-fold), although the difference in expression with each drug was not significant with NUC-1031 or gemcitabine (Two-way ANOVA, p = 0.499). At 72 hours, total RRM1 expression was reduced with NUC-1031 but not gemcitabine in ovarian A2780 cells to 67%, (p = 0.0477) and to 56% at 96 hours post-treatment (p = 0.0187), but not in HuCCT1 cells.

RRM1 expression in patients from a Phase I study with advanced solid cancers does not correlate with survival

We investigated the presence of RRM1 protein in samples obtained from a Phase I study (NCT01621854) of NUC-1031 in patients with advanced solid tumours by immunohistochemistry (Fig. 7A). Analysis was performed on a total of 35 tissue biopsies available, representing 33 of the 68 patients. Patients were categorised according to RECIST v1.1 criteria (WHO, 2009) and divided in two groups: progressive disease and less than 6 months stable disease (group 1); and greater than 6 months stable disease and partial responders (group 2). Median RRM1 expression was greater in group 1 patient samples, however expression was not significantly raised compared to group 2 (Mann Whitney U = 107, n₁ = 16, n₂ = 19, p > 0.05 two tailed) (Fig. 7B), although had a greater RRM1 histoscores within Q3. Patients with stable disease during the Phase I study were stratified by number of months before progression or death (Fig. 7C), which did not show significant correlation with RRM1 histoscores in these patients, indicated by Spearman r test (p > 0.05 two tailed, r= -0.26, ns) and simple linear regression R² = 0.07.

The combination of gemcitabine and platinum agents is used to treat a variety of solid tumours, including those of the biliary tract and ovary; however, patients tend to develop resistance to gemcitabine (Hagmann et al., 2010; Nakano et al., 2015; Saiki et al., 2012). Gemcitabine may also cause liver toxicity, particularly in BTC patients, where there is association with liver disease and intrahepatic dysfunction. Platinum agents such as cisplatin and carboplatin are often used to treat ovarian cancer; however, carboplatin is ineffective in 2D in vitro models (36). Therefore, due to the similar structure and mechanism of action, cisplatin is commonly used as an in vitro alternative in ovarian cancer cell lines (37, 38). Cisplatin-induced lesions following cell cycle arrest can be repaired by several mechanisms that remove the DNA crosslinks. These processes may be interrupted by the misincorporation of fluorinated deoxycytidine. The mode of action of NUC-1031 and the possibility of synergy in combination with cisplatin was therefore investigated.

NUC-1031 can enter cells and is converted to dFdCTP within 2 hours of exposure, lasting up to 96 hours post-treatment, although direct measurement of NUC-1031 falls below the limit of detection. The active metabolite is incorporated into DNA during S phase, preventing progression of replication forks. Maintenance and recovery of replication forks is necessary for genomic stability and is co-ordinated by the DDR at these sites. Increasing ATM phosphorylation at these sites over time indicates a signal transduction from double-strand break detection. This initiates the phosphorylation cascade of multiple downstream DNA repair proteins (29) and encourages their assembly at the break site, with widespread histone H2AX phosphorylation, promoting the initiation of HR (39). Phosphorylation of Chk1 indicates a signal transduction via p-ATR, from single-strand DNA exposure, at resected DNA damage sites (40). The phosphorylation of these damage markers occurred after an increase in S phase arrest, alongside presence of fluorinated deoxycytidine in DNA. This indicates stalled or collapsed replication forks, where one-ended double strand breaks form (41), increasing single-strand DNA exposure, collapsing into double strand breaks. The observed DNA damaging effect of NUC-1031 may be clinically translatable. This may be the case in ovarian carcinoma, where NUC-1031 is stable in patient plasma and tumour shrinkage is found over a number of months (Kazmi et al., 2021).

The DNA damaging effect over time may be partially due to RRM1 binding and inhibition which can alter deoxynucleotide pools. The intermediate metabolite dFdCDP is known to bind to the alpha subunit of E. Coli RNR, homologous to the human large subunit RRM1 (34, 43). This changes the conformation of RRM1 and can be visualised on Western blots, as seen with NUC-1031 and gemcitabine treatment at equipotent doses. The increased bound portion over a longer timeframe observed with NUC-1031 is due to greater availability of intracellular dFdCDP, as a higher concentration of NUC-1031 was used. Bound RRM1 may inhibit RNR function, disrupting deoxynucleotide pools (data not shown) which, in turn, leads to DNA damage due to reduction in resources for DNA replication. Although a proportion of RRM1 is bound by dFdCDP, for up to 96 hours post-treatment, the cytotoxic implication may be limited. The total RRM1 expression increased in response to NUC-1031 and gemcitabine at 24 and 48 hours, which offsets the effect of RRM1 binding at these early timepoints. Low RRM1 expression in patient tissues has been suggested as a good prognostic factor for overall survival rates in BTC and ovarian cancer patients previously treated with gemcitabine and cisplatin (34, 44). BTC and ovarian cancer patients with higher RRM1 expression in response to gemcitabine were reported to have lower overall survival, by 4 months and 6 months, respectively (44, 45). Correlation has also been reported in non-small cell lung cancer and pancreatic cancer studies (46, 47). We, however, find no conclusive correlation between RRM1 expression and survival across multiple cancers from patients enrolled in a Phase I study of NUC-1031.

Synergy in cell cytotoxicity was not observed in vitro in HuCCT1 cells at 96 hours, with either Bliss or Loewe null criteria, also previously reported (48). This is similarly the case for gemcitabine and cisplatin in various BTC cell lines (Sakamoto et al., 2019). Synergy with varying concentrations has been reported with ovarian cancer cell lines (37, 50), inferring variation with cancer type and gene expression. The timepoint at 96 hours based on single agents may not be appropriate as the combination may change the relevant end point for cytotoxicity through a synergistic mechanism. Identifying key mechanisms relating to timeframes of drug activation and lesion repair may explain differences in response to combinations.

The incorporation of dFdCTP into DNA, in combination with cisplatin may stimulate a response from overlapping mechanisms if drugs are available and active intracellularly. Lesions induced by either single agent causes arrest in both S and G2 phases. This may trigger NER and HR pathways, and involve FA pathway componants. The prolonged intracellular availability and DNA incorporation of NUC-1031 may influence the viability of these repair options when combined with cisplatin. Cisplatin alone triggers a DSB response (p-ATM and γH2ax) but not a single-strand break response (p-Chk1) at 48 hours post treatment. These signals cause G2 arrest, preventing M phase entry (Abraham 2001). The timeframe for a DNA damage response to NUC-1031, however, occurs over 72 hours, inferring a period of overlap when combined with cisplatin. This correlates with the increase in damage markers for double and single strand breaks which peak at the earlier timepoint of 48 hours, suggesting NUC-1031 augments the cisplatin lesion repair., NUC-1031 specific DNA incorporation together with DNA strand breaks caused by cisplatin both increase in S and persistent G2 arrest. This may indicate an additive effect with both drugs through different mechanisms. Synergy may occur in p-Chk1 signalling, coinciding with the G2 arrest at 48 hours. This suggests that either new lesions are formed, or activation of an alternate repair pathway utilising p-Chk1, such as the FA pathway. This is associated with cisplatin lesion repair via the ICL pathway (Guervilly et al., 2008). The combination of NUC-1031 and cisplatin may be applicable in patients with the addition of a Chk1 inhibitor to increase the extent of DNA damage induced.

The time-dependent effects of NUC-1031 and cisplatin, based on the activation and incorporation of dFdCTP in vitro, may be relevant in vivo, due to the longevity of intracellular NUC-1031. Assessment of synergy based on mechanism of action offers additional data which may give greater insight into whether the combination of two drugs is clinically relevant.

Cell culture

Cell lines were maintained at 37⁰C, 5% CO₂ and cultured in Roswell Park Memorial Institute (RPMI) 1640 medium, supplemented with 10% foetal bovine serum and 1% penicillin/streptomycin. All lines were routinely tested for Mycoplasma. Work was conducted in a Euroclone class II microbiology safety cabinet. Human intrahepatic cholangiocarcinoma cell line HuCCT1 was obtained from the Japanese Cancer Research Resource Bank (JCRB). Human ovarian epithelial carcinoma cell lines, A2780, PEO1 and PEO4 were gifted from Dr Simon Langdon (University of Edinburgh). Cell lines were stored in liquid nitrogen until use.

Cell cytotoxicity assay

NUC-1031 was provided by NuCana plc. Gemcitabine and cisplatin was purchased from Sigma-Aldrich. The cytotoxicity of these compounds was assessed by a dose-response sulforhodamine B (SRB) assay. The half-maximal inhibitory concentration (IC₅₀) of the drugs that result cell death in vitro was determined for each cell line relative to untreated cells. Cells were plated in 96-well plates at a density of 2000 and 750 cells/well for HuCCT1 and A2780, respectively. Cells were incubated for 48 hours at 5% CO_2, 37⁰C before drug treatment for 2 hours (A2780) or 24 hours (HuCCT1) and replaced with fresh, drug-free media. The IC₅₀ for each drug was measured over 10 concentrations spanning: 0.5–15 µM cisplatin, 10–500 nM gemcitabine, and 0.02-5µM NUC-1031. Cell viability was determined at 96 hours post-treatment by SRB assay as previously described (52)and the optical density recorded on a Biohit BP800 plate reader using a 540 nm filter.

Flow cytometry

1.5x10⁵ HuCCT1 or 1x10⁵ A2780 were seeded in 6 well plates, adhered for 48 hours prior to treatment with NUC-1031 and cisplatin in isolation or in combination for 2 hours (A2780) or 24 hours (HuCCT1). Cells were harvested for flow cytometry at 24-, 48-, 72- or 96-hours post-treatment: Samples were detached, washed in ice-cold TBS and resuspended in a final volume of 200 µL as a single-cell suspension. Cells were fixed and permeabilised by drop-wise addition of 1 ml Ice-cold 70% ethanol under gentle vortex and incubated on ice for 30 mins. Samples were washed twice with 0.5% BSA in TBS and centrifuged at 500xg for 5 mins. Cell counts were adjusted to 2x10⁵ cells to normalise for antibody incubation. Antibodies (APC-Fire750 conjugated-anti-γH2AX Ser139 (BioLegend); PE-conjugated anti-p-Chk1 Ser345 (ThermoFisher Scientific); APC-conjugated anti-p-ATM Ser 1981 (ThermoFisher Scientific) were diluted to 5 µL/million cells in 0.5% BSA in TBS and samples incubated for 1hr at RT. Excess antibody was washed twice in TBS and resuspended in 100 µL. Samples were stained with additional 100 µL 1.5 mM DAPI for 5 mins before flow cytometry using CytoFLex flow cytometer (Beckman-Coulter). Single-labelled and unlabelled samples were used as internal controls for each experiment. Samples were gated for single cell populations and fluorescence intensity for yH2AX, p-ATM, p-Chk1 and DAPI measured in APC-750, APC, PE and PB450 channels respectively. Cell populations, gating and signal intensity was performed and analysed using FlowJo v10.8.1 software.

Western Blotting

HuCCT1 and A2780 cells, were treated with either NUC-1031 or gemcitabine. Lysates were harvested at 24-hour intervals: cells were washed twice in ice-cold PBS and lysed in ice-cold lysis buffer (New England Biolabs) supplemented with cOmplete mini protease inhibitor (Roche), aprotinin (Sigma Aldrich), and phosphatase inhibitor cocktails 2 & 3 (Sigma Aldrich). Samples were sonicated at 4°C, 10s on, 30s off for 3 cycles prior to centrifugation at 13,000xg for 6 mins at 4°C. Protein concentration was determined by Bicinchoninic Acid assay using Pierce BCA protein assay kit (Thermo Scientific, UK) and adjusted to 1 mg/mL with supplemented lysis buffer and 5x Boster’s SDS PAGE reducing Sample Buffer. Adjusted samples were incubated at 95°C for 5 minutes to denature proteins. Protein samples (25 µg) were resolved by 15% SDS Polyacrylamide Gel Electrophoresis, alongside Chameleon Duo protein ladder (LI-COR), then transferred onto nitrocellulose membrane using a Trans-Blot® SD Semi-Dry Transfer Cell. The membranes were blocked with 5% Marvel dried milk in TBS and 1% Tween 20 (Sigma -Aldrich)) for 1 hour at RT. Membranes were incubated with primary antibodies: 1:1000 anti-RRM1 rabbit antibody (Proteintech Europe, catalogue no. 10526-1-AP) and 1:10000 anti- β-actin mouse antibody (LI-COR Biosciences, catalogue no. 926-42212) overnight at 4°C, followed by washes with excess TBS with 1% Tween 20. Membranes were incubated with IRDye® 680CW and IRDye® 800CW conjugated secondary antibodies (1:10000 goat anti-rabbit, (LI-COR Biosciences; catalogue no. 92632211 and 1:10000 donkey anti-mouse, LI-COR Biosciences; catalogue no. 2814912 for 45 minutes at RT. After secondary antibody incubation and washes, membranes were dried in the dark and imaged using Odyssey® CLx Imaging System (Licor). RRM1 and β-actin proteins were visualised and quantified using Image Studio Lite software compatible with the LiCor Odyssey DLx imaging system.

Mass Spectrometry

Preparation of NUC-1031 and dFdCTP extracts

3 x 10⁵ HuCCT1 cells or 1.5 x 10⁵ A2780 cells were plated in 10 cm dishes and grown for 48 hours prior to treatment with different concentrations of NUC-1031 for either 24 or 2 hours, respectively. Cells were washed, detached with trypsin, and washed twice in ice-cold PBS. Pellets were resuspended in 500 µL ice-cold 80% LC-MS grade methanol, vortexed and incubated at -80°C for 20 minutes. Resuspended samples were centrifuged at 14000 RPM for 5 minutes and the supernatant stored at -80°C until analysis by LC-MS/MS. 1 µL of working internal standard solution (10µM ¹³C₁₀¹⁵N₅-dATP) was added to each sample. Samples were mixed thoroughly before drying under a stream of nitrogen gas. All samples were reconstituted in 75 µL of 20% LC-MS grade acetonitrile, vortex mixed and centrifuged for 5 minutes at 14000 rpm. 70 µL of supernatant was transferred to a 96 well plate and analysed by LC-MS. Calibration standards were prepared for NUC-1031 and dFdCTP across the range 0.005–1 nM and 0.5–4000 nM, respectively.

Preparation of hydrolysed DNA samples

Cells were washed with PBS, detached using trypsin and washed twice in ice-cold PBS. Cell suspensions were centrifuged at 3500 RPM for 5 minutes. Pellets were resuspended in 200 µL ice-cold PBS and transferred into 1.5 mL Eppendorf tubes. DNA was extracted using Qiagen DNA mini kit, according to the manufacturer’s recommendations. Extracted DNA was eluted using LC-MS grade water, quantified by Nanodrop and adjusted to 0.25 mg/mL in 25 µL. DNA samples were denatured by heating at 95°C for 5 minutes, and chilled on ice for 3 minutes before hydrolysis. A DNA hydrolysis mix was prepared from 3.75 mL DNA hydrolysis buffer (20mM Tris HCl, pH 7.9; 100mM NaCl; 20mM MgCl₂), 1 µL bacterial alkaline phosphatase (ThermoFisher Scientific), 25 µL deoxyribonuclease (Roche), and 1.5 µL phosphodiesterase I isolated from Crotalus adamanteus venom (ThermoFisher Scientific). 150 µL of DNA hydrolysis mix was used to lyse 5 µg DNA in a thermocycler at 37°C for 18 hours before storing at -20°C until analysis. 1 mL LC-MS grade methanol was added to all DNA samples with 10 µL of an internal working standard solution (0.5 µM ¹⁵N₂¹³C-dFdC, 50 µM ¹⁵N₂¹³C 2’dG) and dried under a stream of nitrogen gas. All samples were reconstituted in 100 µL of LC-MS grade water, vortex mixed and centrifuged for 5 minutes at 14,000 RPM. 90 µL of supernatant was transferred to a 96 well plate and analyzed by LC-MS. Calibration standards were prepared for dFdC and 2’dG across the range 0.01-18 nM and 0.1–180 µM, respectively.

Metabolite analysis by LC-MS and LC-MS/MS

Quantification of NUC-1031, deoxynucleotide triphosphates and DNA bases were carried out by Liquid Chromatography tandem Mass Spectrometry (LC-MS/MS) using a Quadrupole time of flight (Q-TOF) instrument (Acquity H-Class UPLC system coupled to a Waters Xevo G2-XS Q-TOF mass spectrometer). The eluent from the LC system was infused directly into an electrospray ionisation (ESI) source, operated in either positive or negative ionisation mode. Conditions and m/z for each analyte are detailed in Supplementary Table 1. Each sample analysed for NUC-1031 and dFdCTP was injected under two separate analytical conditions and was analysed in either full scan TOF mode (50–800 m/z) or by targeted analysis using multiple reaction monitoring (MRM). Samples measured for DNA were analysed in MRM mode only. Internal controls “single black” included internal standard solution and “double blank” containing only LC-MS grade water were included. All solvents and buffers were of LC-MS grade.

Chromatographic parameters

NUC-1031 analysis was carried out on Waters UPLC BEH C₁₈ 130Å, 1.7 µm, 2.1 mm X 50 mm column at a flow rate of 400 µL min^− 1 at initial conditions of 70% water + 0.1% formic acid, with 30% acetonitrile + 0.1% formic acid until 0.5 min, followed by a linear gradient to 99% acetonitrile + 0.1% formic acid until 3 min and held to 4 min. This was followed by re-equilibration to 70% water + 0.1% formic acid with 30% acetonitrile + 0.1% formic acid to 6 min. Triphosphate analysis (dFdCTP) was carried out on a Waters Atlantis Premier BEH C₁₈ AX 1.7 µm, 2.1 mm X 100 mm column at a flow rate of 300 µL min^− 1 at initial conditions 90% 10 mM ammonium acetate pH 6 with 10% acetonitrile until 0.5 min. This was followed by a step change to 50% 10 mM ammonium acetate (pH6), 40% 10 mM ammonium acetate (pH10), with 10% acetonitrile and re-equilibration to 90% 10 mM ammonium acetate (pH6) with 10% acetonitrile to 7 min. DNA analysis (dFdC and dG) was carried out an ACE Excel 2 AQ 2.1 mm X 100 mm column with a flow rate of 400 µL min^− 1 at initial conditions of 99% water + 0.1% formic acid with 1% acetonitrile + 0.1% formic acid held for 1 min, followed by a linear gradient to 99% acetonitrile + 0.1% formic acid until 8 min and held to 10 min and re-equilibration to 99% water + 0.1% formic acid with 1% acetonitrile + 0.1% formic acid to 14 min. The column temperature for each run was set to maintain 40°C.

Calculations

Calibration lines were plotted linear 1/x² and analyte concentration was calculated either on analyte area or peak area ratio with internal standard using MassLynx (version 4.2). NUC-1031 or dFdCTP abundance was normalised against cell number. DNA incorporation was measured by dFdC in extracted DNA and was normalised to dG, by simultaneous quantification of both analytes (53).

Immunohistochemistry

Study on tissue samples from First-in-Human Phase 1 study - ProGem1 (NCT01621854) patient cohort

Samples were derived from patients with advanced solid tumours were recruited for an open- label, dose escalation Phase I clinical study to assess the safety and anti-tumour activity of NUC-1031. The enrolment of 68 patients commenced on 15th June 2012 and concluded on 26th August 2015, conducted by Blagden et al. (54). Information regarding the clinical trial is available at https://classic.clinicaltrials.gov/ct2/show/NCT01621854. Immunohistochemistry was performed on the 35 available tissue biopsies representing 33 of the 68 patients enrolled in the study. Patient-derived tissue biopsy sections had previously been fixed in 10% formalin and processed according to the routine histopathology standard operating procedure to generate formalin-fixed paraffin-embedded (FFPE) tissue blocks. 3µm sections were previously cut from the FFPE blocks to be stained by immunohistochemistry. Sections were stained for RRM1 expression using a Leica Biosystems BONDIII system. Samples were dewaxed with Leica ‘Dewax Solution’, washed with absolute alcohol, followed by Leica BOND wash buffer. Antigen retrieval was performed at 100°C for 20 mins with Leica BOND ER1 (Epitope Retrieval Solution 1). Endogenous peroxidase activity was blocked by peroxide blocking solution in Bond polymer refined detections kit (Leica, #DS9800). RRM1 (Proteintech Europe, catalogue no. 10526-1-AP) primary antibody was incubated for 30 mins, followed by anti-rabbit HRP (horseradish polymer) polymer for 30 mins. Then DAB (3,3′-Diaminobenzidine) chromogen was utilised to visualise HRP polymer labelled RRM1. Slides were dehydrated in a series of 50%, 80% and twice in 100% ethanol and cleared in xylene three time for 5 mins each and mounted with distyrene-plasticiseran-xylene (DPX). Slides were scanned on a ZEISS Axioscan z1 at 20x brightfield.

Image analysis

Images were analysed with an open access software, QuPath version 0.1.2, for digital quantitative pathology (https://qupath.readthedocs.io/en/stable/docs/intro/citing.html-). Regions of tumour were manually annotated then individual tumour cell nuclei were segmented using the ‘cell detection tool’ in QuPath and DAB intensity was used to perform histoscores within tumour annotations, excluding non-tumour cells. The optical density of DAB intensities of RRM1 was set at 0.2, 0.4, and 0.6 to classify weakly, moderately, and strongly positive cells, respectively. The population of these three grouped cells were converted into a histoscore (Hscore). The histoscore was determined by the equation: Hscore= ((1 × % weakly positive cells)+(2 × % moderately positive cells)+(3 × % strongly positive cells)) to yield a potential range of 0-300 for each annotated tumour region. (Jensen et al., 2016).

Statistical analysis

Histoscores from 35 cases were combined with their corresponding clinical data. Graphs, Spearman r test and simple linear regression were generated using GraphPad Prism 9.

Author contributions

Main manuscript text written by DP. Conceptualization of project by DP, ALD, DJH, and JB. Data collection, optimization, and analysis by DP, ALD, GMZ, IHU, and OJR. Contribution to preliminary experimental data by PM. Resources for mass spectrometry provided by CMC. All authors reviewed the manuscript.

Data availability statement

The datasets generated and/or analysed during the current study are not publicly available due to agreements with funders but are available from the corresponding author on reasonable request.

Competing Interests Statement

DP and GMZ were funded by Nucana plc https://www.nucana.com/. ALD and DJH

partly employed by Nucana plc. OJR, JB were at time of research, fully employed by

Nucana plc. The sponsor did not have any role in the study design, collection or analysis. The

funders approved the decision to publish and provided feedback on the written

manuscript

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Competing interest reported. DP and GMZ were funded by Nucana plc https://www.nucana.com/. ALD and DJH partly employed by Nucana plc. OJR, JB were at time of research, fully employed by Nucana plc. The sponsor did not have any role in the study design, collection or analysis. The funders approved the decision to publish and provided feedback on the written manuscript.

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Defining the mode of action of cisplatin combined with a phosphoramidate modification of gemcitabine

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Abstract

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Introduction

Results

NUC-1031 forms dFdCTP which is incorporated into DNA

NUC-1031 arrests cells in S phase of cell cycle and elicits a DNA damage response

NUC-1031 binds Ribonucleotide Reductase subunit RRM1

Discussion

Materials and Methods

Statistical analysis

Declarations

References

Additional Declarations

Supplementary Files

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