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Metallodrugs for therapy and imaging : investigation of their mechanism of action Spreckelmeyer, Sarah 2018

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 1     METALLODRUGS FOR THERAPY AND IMAGING – INVESTIGATION OF THEIR MECHANISM OF ACTION   by   SARAH SPRECKELMEYER   Diploma and State Exam, The University of Greifswald and the University of Braunschweig, 2013   A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF DOCTOR OF PHILOSOPHY   in   THE FACULTY OF GRADUATE AND POSTDOCTORAL STUDIES  (Chemistry)  THE UNIVERSITY OF BRITISH COLUMBIA  (Vancouver)  February 2018 © Sarah Spreckelmeyer, 2018      2 		Metallodrugs for Therapy and Imaging:  Investigation of Their Mechanism of Action 	PhD thesis 	to obtain the joint degree of PhD at the  University of Groningen    on the authority of  the Rector Magnificus of the University of Groningen,  Prof. E. Sterken,  and in accordance with  the decision by the College of Deans of the University of Groningen      This thesis will be defended in public on  Friday, 23 February 2018 at 16.15 hours     by  Sarah Spreckelmeyer  born on 8 May 1989  in Osnabrück, Germany       3 Abstract In this thesis, the application of metallodrugs for therapy and imaging was investigated as part of the field of medicinal inorganic chemistry. In the introduction, an overview was given on the application of organic molecules, incorporating a metal or radiometal for either therapeutic or diagnostic purposes, particularly in relation to cancer. It is evident that their mechanism of action, their pharmacokinetic behaviour and biological targets are mostly not fully elucidated yet. Thus, our overall aims included: i) to synthesize new radiopharmaceuticals for either cancer therapy or imaging, and ii) to elucidate the mechanism of cellular uptake and excretion, the anti-cancer activity and the organ toxicity of some new Au containing metallodrugs in comparison to cisplatin. To investigate the toxicity and transport mechanisms of the new cytotoxic organometallic Au(III) compounds, the ex vivo model of precision cut tissue slices and human cell cultures in vitro were used.  The work described in Part A was performed at the University of British Columbia in Vancouver, BC, Canada in the Department of Medicinal Inorganic Chemistry under the supervision of Prof. Chris Orvig. Different radiotracers were developed and characterized by chemical-physical methods. Radiolabeling experiments were also performed at TRIUMF (Canada’s national laboratory for particle and nuclear physics and accelerator-based science) and in vivo animal experiments were conducted at the BC Cancer Agency.  In chapter A1, we report on the synthesis of H4neunpa and its immunoconjugate H4neunpa-trastuzumab (Figure 1) and showed that it can be efficiently radiolabeled with 111In3+ at ambient temperature within 15 min or 30 min, respectively. 111In is a gamma emitter and can thus be used for single photon emission computed tomography (SPECT). The immunoconjugate was further investigated in an in vivo model using HER2/neu positive subcutaneous SKOV-3 ovarian cancer xenografts bearing mice. Unfortunately, our results showed an unexpected lower accumulation of 111In-neunpa-trastuzumab into the tumor compared to the gold-standard 111In-CHX-DTPA-trastuzumab, which was supported by Immuno-SPECT images taken after 1 day, 3 days and 5 day post injection. Consequently, 111In-neunpa-trastuzumab does not seem to be suitable for clinical application, although in vitro experiments (immunoreactivity, radiolabeling efficiencies, stability in human serum) showed similar or superior properties of this chelator compared to the gold-standard. A reason for this lower accumulation might be a difference in internalization process of the chelator-HER2/neu-receptor complex. Finally, radiolabeling of H4neunpa with 177Lu, a therapeutic radiometal, was tried but appeared unsuccessful. Comparison of In3+ (92pm CN=8) and Lu3+ (103pm CN=9)1 leads to the hypothesis that either Lu3+ might be too big for H4neunpa or the preferred coordination number is not saturated because of steric hindrance.          4           Figure  1.  Summarized radioconjugates derived from H4neunpa-p-Bn-NO2.    On the other hand, 225Ac and 213Bi radiolabeling of H4neunpa-p-Bn-NO2 was successful, as described in chapter A2. 225Ac and 213Bi are alpha emitters that can be used for targeted alpha therapy (TAT). Additionally, this chelator could be conjugated to a PSMA targeting molecule, Glu-ureido-Lys, resulting in H4neunpa-PSMA-L (Figure 1). Radiolabeling of H4neunpa-PSMA-L with 111In was disappointingly low, which could possibly be explained by the short distance between Glu-ureido-lys and the chelation cavity of H4neunpa, thereby chelating the 111In by the carboxylic acids of Glu-ureido-lys. Furthermore, metallacage-ligand linkage to H4neunpa and incorporating La3+ was successful and fluorescence spectroscopy with La3+ gave promising results, showing better fluorescent properties of La-neunpa-metallacage-ligand compared to H4neunpa-metallacage-ligand without La3+.  Antimony complexation reactions gave promising results as well, making the complex suitable for targeted radiotherapy, since 119Sb is an Auger emitter. Overall, the results in chapter A1 and chapter A2 show that H4neunpa can be considered as an excellent bifunctional chelator. Modifications can be done easily, depending on the target molecule and radiometal of interest. It was also found that radiolabelling is dependent on the size of the biomolecule that is coupled as targeting moiety. Changing the biomolecule from the antibody trastuzumab to smaller biomolecules with a shorter linker (eg. Glu-ureido-Lys) resulted in different radiolabelling efficiencies. The work on this chelator is currently continued. As the synthesis of H4neunpa is a 10 step reaction with a 2.3 % overall yield and difficult to apply for clinical use, a faster synthetic protocol was developed, by which H4neunpa can be synthesized in only 4 steps (Figure 2).      N N N OHOHOON NHO OHO ONO2H₄neunpa-p-Bn-NO₂H₄neunpa-PSMA-LN NNOHONHOON OHHOO OHN NHSGlu-ureido-lysN NNOHONHOON OHHOO OHNH₄neunpa-p-Bn-trastuzumabNHStrastuzumabN NNOHONHOON OHHOO OHNH₄neunpa-metallacage-ligandNHSmetallacage-ligandChapter	A1Chapter	A2Chapter	A2111In3+ (SPECT)225Ac3+ (Therapy)213Bi3+ (Therapy)natSb3+ (Therapy)177Lu3+ (Therapy)Chapter	A1Chapter	A2Chapter	A2Chapter	A2Chapter	A1 5          Figure  2.  Proposed synthesis route of H4neunpa.   In future work, the neunpa-PSMA-L could be improved by modifications of the length of the linker between Glu-ureido-Lys and H4neunpa. The complexation with radioactive antimony would allow performing radiolabeling experiments with this radioisotope. As 89Zr4+ is a promising radiometal for PET imaging, two hydroxamic acid bearing ligands have been synthesized as described in chapter A3, that were tested for their toxic effects towards several cancer cell lines. The synthesis of these two hydroxamic acid bearing ligands was successful and chemical-physical analysis supported this. Unfortunately, 89Zr radiolabeling was unsuccessful, possibly due to the inflexible hydroxamic acids arms as calculated with density functional theory (DFT). These findings are probably also the reason for the low toxicity of these compounds in cancer cell lines, as they are also not able to bind essential cations (eg. Cu2+ and Fe3+) in a stable manner, which is essential for the toxicity of these compounds. Decreasing emission bands in UV-VIS experiments confirmed the instability of the Fe-complexes. In the future, DFT calculations should be performed before a set of compounds is synthesized, as done for the second generation of hydroxamic acid bearing ligands, as proposed in chapter A3. In the future, after the successful synthesis and characterization of these new hydroxamic acid bearing ligands, radiolabeling experiments with 89Zr as well as evaluation of their stability in human serum needs to be performed.  In chapter A4, we reported on a project which aim was to synthesize a bifunctional chelator H2dedpa with a thiol reactive moiety for conjugation with FXa (factor Xa, a component of the blood coagulation cascade) in order to localize blood clots in patients. Our results showed that the synthesis of a bifunctional chelator H2dedpa that bears a thiol reactive moiety for FXa conjugation was difficult. Three different approaches were unsuccessful, but the fourth attempt was successful and resulted in the synthesis of H2dedpa-acrylate as proven by various chemical-physical techniques. Unfortunately due to time restrictions it was not possible to further investigate this molecule. Part B was investigated at the University of Groningen under the supervision of Prof. Geny Groothuis and co-supervision of Prof. Angela Casini. The anticancer effects on various human cancer cell lines, kidney toxicity and accumulation mechanisms of several novel Au(III) cyclometallated compounds compared to cisplatin were studied. NH2NH NH2NO2BrK2CO3, DMFNa2CO3, CH3CN OOBrPd/C 10%glacialCH3COOHLiOHTHF/H2O 3:1SCCl2 H2O, DCMN N N OHOHOON NHO OHO ONCSN N N OOOON NO OO ONO2N N N OHOHOON NHO OHO ONO2LiOH THF/H2O 3:1N N N OOOON NO OO ONH2N N N OHOHOON NHO OHO ONH27 H4-neunpa-p-Bn-NO26 H4-neunpa-p-Bn-NCS12345OO O NH NHHNN NO OO ONH N HNN NO OO ONO2ACN 6 In part B1, the state of knowledge of transport mechanisms of cisplatin and other metallodrugs was reviewed with the conclusion that there is a substantial lack of knowledge on the accumulation mechanisms of cisplatin and of the new generation anticancer metallodrugs at the molecular level. Such knowledge is necessary to elucidate the balance between activity and toxicity profiles of metal compounds. Furthermore, resistance mechanisms often involve drug transporter expressions in the targeted cells.2,3 Many experiments to study these transport mechanisms were performed in cells in vitro and studies performed in ex vivo or in vivo models are rare. Based on these studies in cell cultures, the transporter proteins OCT2 and CTR1 are hypothesized to be involved in the uptake of cisplatin into the cells, especially in kidney cells and may be responsible for kidney accumulation and severe nephrotoxic side-effects. APT7A/B and MATE are efflux transporters likely to be involved in the efflux of cisplatin and other Pt(II) drugs (Figure 3). Furthermore, other studies also support the involvement of passive diffusion mechanisms.          Figure  3.  Drug transporters possibly involved in cisplatin accumulation.   7 In chapter B2, a set of organometallic Au(III) compounds featuring bidentate C^N type of ligands were synthesized and characterized as well as studied for their anticancer properties in different human cancer cell lines in comparison to non-tumorigenic cells. Among the various Au compounds developed as anticancer agents, the use of cyclometallated complexes is advantageous due to redox and thermodynamic stability. Additionally, their lipophilic character can be tuned by the modification of the ancillary ligands or steric and electronic properties can be easily tuned by modification of the anionic cyclometallated ligands. Overall, our study shows the potential for improvement of the biological properties (like toxicity or PARP-1 inhibition) of organometallic gold-based compounds by tuning their coordination environment by changing the chlorido ligand to a PTA moiety to increase its water solubility or glucose moieties to target GLUT1 transporter.          Figure  4.  Au(III) organometallic complexes discussed in this thesis.  The most active Au(III) compound [Au(pyb-H)(PTA)Cl]PF6 (PTA=1,3,5-triazaphosphaadamantane) was chosen for further characterization of the toxicity and transport mechanisms of the compound in an ex vivo model of precision cut kidney slices (PCKS) in chapter B3. We found, that this novel Au(III) compound (Figure 4) shows also markedly toxic effects on PCKS under the selected experimental conditions after 24h incubation compared to cisplatin. Both compounds induce a concentration dependent decrease in viability of PCKS, which correlated with the Au or Pt content, respectively. The Au(III) compound  showed lower TC50 values compared to cisplatin, being 4.3 ± 0.2 µM and 17 ± 2.0 µM respectively. Additionally, this new experimental metallodrug was studied for its mechanism of transport and cellular accumulation in kidney slices in comparison to cisplatin.  Using cimetidine as an inhibitor for OCT2 and MATE, we showed that there is no evidence that either the Au(III) compound or cisplatin are transported via OCT2 or MATE. In the case of cisplatin, this is in contrast to previously reported results. As commented in chapter B3, this may be due to the important differences between the previously reported cell-based models/assays and our tissue culturing method. Another explanation might be that cimetidine is not only an inhibitor of OCTs and MATEs, but also an H2-receptor antagonist and it can also inhibit some of the drug-metabolizing cytochrome 450 (CYP).4,5 Thus, several transporters and receptors are a target of cimetidine. Consequently is might be possible, that either cimetidine does not have a full effect on the OCTs or MATEs, since other targets have a higher affinity, or other transporters or receptors which can not NAuClNPNNPF6NAuPPh2OOOClPF6[Au(py'-H)(PTA)Cl]PF₆Chapter	B2	&	B3Compound	1Chapter	B4 8 be inhibited by cimetidine might have an effect on cisplatin’s or Au(III) compound’s uptake and consequently toxicity. To evaluate this, further experiments are needed. Firstly, a positive control that proofs cimetidine’s ability to block OCT and MATE transport is necessary. Secondly, a more specific inhibitor of OCTs or MATEs should be studied, but to our knowledge, no specific inhibitor is available yet. Metal quantification in tissues was also achieved by ICP-MS, while histomorphology studies allowed providing evidence of the damage of specific cell types, namely distal tubular cells compared to proximal tubular cells for cisplatin. Since the Au(III) compound seems to be very toxic, we need to prevent these toxic effects by targeting it more specifically to the cancer tissue. This can be tried by linking it to peptides or antibodies with affinity for the cancer cells as described for radiopharmaceuticals.  In chapter B4, the involvement of OCT2/MATE and CTR1/ATP7A/B in several cancer cell lines was studied for another C^N Au(III) compound, named Au(III) compound 1, containing a  fluorescent coumarin moiety which makes it suitable for fluorescent microscopy. Using cimetidine as an inhibitor for OCT2/MATE and CuCl2 as a competitor for CTR1/ATP7A/B, we studied the influence of these transporters on the Au(III) compound 1 accumulation in A2780 and A2780cisR (A2780 cells resistant for cisplatin) cells compared to cisplatin. The Au(III) compound 1 seems to be more potent than cisplatin as concluded from the IC50 results after 24h and 72h incubation and also showed a higher metal content accumulation via ICP-MS. Co-incubation with CuCl2 increased the toxicity of the Au(III) compound 1 in both A2780 and A2780cisR cells. These findings are supported by an increase in Au and Cu content, leading to the hypothesis that not only Au(III) accumulation but also Cu accumulation might be the reason for the increased toxicity or that CuCl2 inhibits efflux pumps involved in Au(III) accumulation. A direct involvement of the CTR1 or OCT2 in the uptake of the drug could not be shown by inhibition by CuCl2 or cimetidine. Cisplatin showed a similar behavior. After 72h, co-incubation of cisplatin with either cimetidine or CuCl2 resulted in an increase in toxicity, but no increase in Pt content could be observed in both cell lines by ICP-MS. This result, together with the evidence for increased Cu content in A2780 cells, leads to the hypothesis that copper accumulation is the reason for the increased toxicity in these cell lines. Overall, both research parts A and B are examples of multidisciplinary collaborations. For radiopharmaceuticals, an expertise network of radiopharmacists (incl. radiochemists) and nuclear medicine physicians is essential for meeting the needs in clinic and patient care as well as taking benefit from the fundamental knowledge of chemists and biologists. In my opinion, such collaborations are ongoing, but they could be intensified and extended in order to accelerate the development of new and better radiopharmaceuticals and prevent an inefficient use of money. Finally, taking into account the applications of radiotracers in the clinic, macrocyclic chelators like DOTA or NOTA are still preferred over acyclic chelators, since they show better stability in patients or superior tumor uptake.  Concerning transporter studies in vitro, ex vivo and in vivo, specific inhibitors for the tested transporter should be used in order to be able to make clear conclusions about the results of transporter inhibition experiments. It should be kept in mind that drug transporters are expressed differently in species and should be taken into account when translating from animal data to human. Moreover differences in transporter and metabolizing enzyme expression between tissues and cell types should be taken into account when evaluating accumulation and toxic effects in cancer cells and different organs.   To conclude, the topic of metallodrugs for therapy and imaging was successfully investigated in this thesis. Two different parts (A and B) focus in detail on radiopharmaceuticals for imaging/diagnosis of specific cancer types and on the mechanisms of transport as well as toxicity of new experimental Au(III) metallodrugs in vitro as well as ex vivo, respectively.  9  Lay Summary In this thesis, metallodrugs were designed and characterized in order to be used for cancer therapy and diagnosis. The first aim was to synthesize new drugs that incorporate a radiometal (resulting in a so-called radiopharmaceutical) for diagnostic or therapeutic application for the treatment of cancer. Secondly, the toxicity and mechanisms of transport of metallodrugs like cisplatin (a widely used anti-cancer metallodrug containing platinum) and novel gold containing drugs were investigated.  Different techniques were used, like basic organic chemistry, in vitro cell experiments and biological assays, drug transporter competition experiments using cimetidine or CuCl2 as inhibitors or competitors, radiolabeling procedures, ex vivo precision cut tissue slicing and in vivo mice experiments. Transport mechanisms via the organic cation transporter 2 (OCT2)/multi drug extrusion protein (MATE) and copper transporter 1 (CTR1) and ATP7A/B were investigated in vitro and ex vivo. We synthesized and characterized successfully a novel molecule (called H4neunpa) that can bind to several radiometals, like 111In (used in imaging for diagnosis), 225Ac and 213Bi (used for therapy). This molecule is stable in vitro and in vivo and further experiments are currently ongoing to test its application in vivo. Furthermore, we found that gold containing drugs can be as potent as cisplatin against cancer cells in vitro, although their mechanism of action is different, but they are also toxic for healthy kidney tissue. Slight structural modifications in a gold containing molecule can change its transport behavior and beside OCT2, MATE, CTR1 and ATP7A/B, other transporters might be involved in their mechanisms of action.                10 Preface 	This dissertation is formatted in accordance with the regulations of the University of Groningen (The Netherlands) and submitted in partial fulfillment of the requirements for a PhD degree awarded jointly by the University of Groningen and the University of British Columbia.  Versions of this dissertation will exist in the institutional repositories of both institutions.  														 11 Table of contents  Abstract……………………………………………...……………….………………..........……………….…...……………….…...……………….………............3 Lay Summary ……………………………………………………………..………………………...…….……...……………….…...……………….…………………..9 Preface……………………………………………………………..………………………......................……………….…...……………….….....…….…………10 Table of contents…………………………………………………..……………………..……………...……………….…...……………….………...…….…………11    Part A: Vancouver………………………………………………..……………..…..…………..……..…………..……..…………..………………………………………………..…………12 A1: p-NO2-Bn-H4neunpa and H4neunpa-Trastuzumab: Bifunctional Chelator for Radiopharmaceuticals and 111In Immuno-SPECT Imaging…………………………………….…...……………………………….…...……………….…...……………….………….…...……………….…….13  A2: H4neunpa: A Bifunctional Acyclic Chelator with Many Faces………………...……………….……….....……………….…………………..61  A3: Tetrahydroxamic Acid Bearing Ligands: EDTA and DTPA Analogues……...……………….…………….……………….…………….….83  A4: Overcoming the Limitations in Thrombosis Treatment: A Bifunctional Chelator as Positron Emission Tomography-Imaging Probe for Detecting Blood Clots…………………………………………………………………….…...………………..….…………….…….….107      Part B: Groningen……………………………………..…………..……..…………..……..…………..……..…………..…………………………………………………..…..………….126 B1: Cellular Transport Mechanisms of Cytotoxic Metallodrugs: An Overview Beyond Cisplatin…………………………………………….………….……………………...……………….…...……………….………………..………………….…………..127 B2: Exploring the Potential of Gold(III) Cyclometallated Compounds as Cytotoxic Agents: Variations on the C^N Theme………………………………………………………………...……………….…...……………….…...……………….…...……………….…...……………….150 B3: On the Toxicity and Transport Mechanisms of Cisplatin in Kidney Tissues in Comparison to a Gold-based Cytotoxic Agent…………………………………….………...……………….…...……………….…...……………….…...……………….…...……………….………………..179 B4: Investigation of the Molecular Accumulation Mechanisms of an Au(III) Cyclometallated Compound Compared to Cisplatin in vitro: Are OCT2 and CTR1 involved?.......………….… ...……………….… .............................................................198 			12 	Part A                	 13    Chapter A1 p-NO2-Bn-H4neunpa and H4neunpa-Trastuzumab:  Bifunctional Chelator for Radiometalpharmaceuticals and 111In Immuno-SPECT Imaging 							Sarah Spreckelmeyer,a,b Caterina F. Ramogida,c Julie Rousseau,d Karen Arane,c Ivica Bratanovic,c Nadine Colpo,d Una Jermilova,d Gemma M. Dias,d Iulia Dude,d Maria de Guadalupe Jaraquemada-Peláez,a François Bénard,d Paul Schaffer,d Chris Orviga  a Medicinal Inorganic Chemistry Group, Department of Chemistry, University of British Columbia, 2036 Main Mall, Van-couver, British Columbia, V6T 1Z1, Canada b Dept. Pharmacokinetics, Toxicology and Targeting, Research Institute of Pharmacy, University of Groningen, Antonius Deusinglaan 1, Groningen 9713 AV, The Netherlands c Life Sciences Division, TRIUMF, 4004 Wesbrook Mall, Vancouver, British Columbia, V6T 2A3, Canada d BC Cancer Agency, 675 West 10th Avenue, Vancouver, British Columbia, V5Z 1L3, Canada Published in: Bioconj. Chem. 2017, 28, 2145-2159.     	14  1 Abstract Potentially nonadentate (N5O4) bifunctional chelator p-SCN-Bn-H4neunpa and its immunoconjugate H4neunpa-Trastuzumab for 111In radiolabeling are synthesized. The ability of p-SCN-Bn-H4neunpa and H4neunpa-Trastuzumab to radiolabel quantitatively 111InCl3 at ambient temperature within 15 min or 30 min, respectively, is presented. Thermody-namic stability determination with In3+, Bi3+ and La3+ resulted in high pM values. In vitro human serum stability assays have demonstrated both 111In complexes to have high stability over 5 days. Mouse biodistribution of [111In][In(p-NO2-Bn-neunpa)]-, compared to that of [111In][In(p-NH2-Bn-CHX-A"-DTPA)]2-, at 1 h, 4 h and 24 h shows fast clearance of both complexes from the mice within 24 h. In a second mouse biodistribution study, the immunoconjugates 111In-neunpa-Trastuzumab and 111In-CHX-A”-DTPA-Trastuzumab demonstrate a similar distribution profile, but with slightly lower tumor uptake of 111In-neunpa-Trastuzumab compared to 111In-CHX-A”-DTPA-Trastuzumab. These results were also con-firmed by Immuno-SPECT imaging in vivo. These initial investigations reveal the acyclic bifunctional chelator p-SCN-Bn-H4neunpa to be a promising chelator for 111In (and other radiometals) with high in vitro stability, and also show H4neunpa-Trastuzumab to be an excellent 111In chelator, with promising biodistribution in mice.  	 15  2 Introduction Early detection and specific therapy are the key factors for the successful treatment of cancer. 111In (t 1/2 = 2.8 days) and/or 177Lu (t 1/2 = 6.6 days) are important radioisotopes in nuclear medicine that match either the requirements for single photon emission tomography (SPECT) and performing dosimetry, or for therapeutic purposes, respectively. 111In being a cyclotron–produced radiometal (via the 111Cd(p,n)111In reac-tion) emits gamma rays (245 and 171 keV) and Auger electrons. 177Lu being a reactor-produced radiometal (176Lu(n,gamma)177Lu) emits primarily beta particles (490 keV) that can be used for therapy.1 A common method to incorporate metallic radioisotopes (i.e. radiometals) into radiopharmaceuticals is via chelation of the desired radioisotope using a bifunctional chelator (BFC). As implied by the name, BFCs possess two properties – they must chelate the radiometal of interest in a tight and stable metal-ligand complex, and the BFC must incorporate a point of attachment for conjugation to a targeting vector (e.g. biomolecule of interest in disease progression such as a peptide or antibody). Both macrocyclic and acyclic chelators are used in the clinic, and are also of interest in the field of medicinal inorganic chemistry research. The pros and cons of cyclic vs acyclic chelators are widely known and beyond debate.2 Relevant to 111In and 177Lu, macrocy-clic DOTA (1,4,7,10-tetraazacyclododecane-1,4,7,10-tetraacetic acid) is the gold-standard chelator, while acyclic chelator DTPA (diethylenetriamine pentaacetic acid) and chiral analogue CHX-A”-DTPA are ubiquitous in 111In radiopharmaceutical devel-opment (Figure 1). Recent studies developed bifunctional somatostatin analogues of DOTA with increased stability in vivo.3 As an acyclic gold-standard, the commercially available radiopharmaceutical OctreoScan (111In-DTPA octeotride) reached approval in 1994 (Figure 1). Since the success of OctreoScan, several more bifunctional acyclic 111In chelators that contain different biomolecules have been developed, hoping to overcome the limitations of OctreoScan. These include an increased physiological uptake which restricts the detection of small lesions, prolonged imaging protocol and relatively high radiation dose to the patients, as well as low image quality.4 Our group has developed several promising acyclic chelators for 111In and/or 177Lu, based on picolinic acid binding motifs, which we have since dubbed the “pa”-family of chelators.5-8 Of note, octadentate H4octapa (N4O4) and its bifunctional ana-logue p-SCN-Bn-H4octapa showed exceptional complexation properties (quantitative 111In or 177Lu radiolabeling in 10-30 minutes at ambient temperature) and favorable in vivo stability of resulting complexes.9,10 Furthermore, chiral ligands H2CHXdedpa (N4O2) and H4CHXoctapa (N4O4) showed promising 68Ga and 111In radiolabeling properties, respectively, and subsequently impressive stability in human serum.8    Our group continues to design ligands that may incorporate large metal ions (such as radioactive actinides/lanthanides for imaging/therapy), which possess ideal 	16 properties for radiopharmaceutical incorporation, e.g. fast, mild, and quantitative com-plexation of radiometals at low ligand concentrations; formation of resultant thermo-dynamically stable and kinetically inert metal-complexes; and a convenient point of attachment to targeting vectors. Herein, we report the synthesis and characterization of a novel nonadentate (CN = 9) acyclic chelator H4neunpa (N5O4, referred to herein as either p-NO2-Bn-H4neunpa or H4neunpa) and bifunctional analogue p-SCN-Bn-H4neunpa that was designed as a bifunctional analogue of H5decapa (N5O5), reported by our group in 2012.5 The carboxylic acid group on the middle nitrogen atom has been re-placed by p-nitrobenzene-ethylene to keep its symmetry, and act as the bifunctional arm to attach the ligand to a biomolecule through a thiourea bond (Figure 1). We hy-pothesized that the extended diethylenetriamine backbone and nine coordinating at-oms of H4neunpa may favorably form complexes with large metal ions such as In3+ (92 pm, CN = 8)11, Lu3+ (103 pm, CN = 9), or Bi3+ (117 pm, CN = 8). Radiolabeling of 111In and 177Lu to H4neunpa was assessed and compared to gold-standards DOTA and CHX-A”-DTPA, and an in vivo biodistribution study of H4neunpa and CHX-A”-DTPA labeled with 111In was performed. Thermodynamic stability constants of selected metal-neunpa complexes were also determined. Moreover, coupling of the HER2/neu targeting mono-clonal antibody (mAb) Trastuzumab was performed via the reaction between the anti-body’s primary-amine(s) with the isothiocyanate functional group of p-SCN-Bn-H4neunpa. The bioconjugate was labeled with 111In, and in vivo biodistribution and SPECT/CT imaging studies were conducted and compared directly to a 111In-CHX-A”-DTPA-Trastuzumab conjugate.      Figure  1.  Structures of cyclic (DOTA) and acyclic (OctreoScan, CHX-A"-DTPA) com-mercial chelators, and acyclic “pa”-ligands H2CHXdedpa, H4CHXoctapa, H4octapa, H5decapa, and novel nonadentate chelator p-SCN-Bn-H4neunpa discussed in this work.  NH HNN NOHOHOON NNOHONHOONOHOHHOO OON NNOHONHOON OHHOO ONCSH2CHXdedpaH5decapa p-SCN-Bn-H4neunpaN NN NOHOHOOH4CHXoctapaOHOOOHN NN NOHOHOOH4octapaOHOOOHN NNNDOTAN N N NHHOO OOOHOHOOOHOctreoScansomatostatinOOHOHOOOHOHON N N OHHOO OOOHOHOOHOp-SCN-Bn-CHX-A''-DTPANCS	 17 3 Results and Discussion 3.1 Synthesis and characterization of the ligand The synthesis of the previously reported analogue H5decapa used N-benzyl protection, N-alkylation with an alkyl halide, benzyl deprotection via hydrogenation, a second alkyl halide N-alkylation, and finally deprotection in refluxing HCl (6M).10 The N-benzyl protection was found to be the yield-limiting step because the deprotection always resulted in partly eliminating the picolinic acid moieties. The use of O-nitrobenzenesulfonyl (nosyl) was found to give better cumulative yields compared to N-benzyl protection. Based on that, the bifunctional analogue H4neunpa, was synthesized with a general reaction scheme that follows N-nosyl-protection, bifunctionalization on the middle nitrogen atom via N-alkylation, N-alkylation with picolinic acid, nosyl-deprotection with thiophenol, a second alkyl halide N-alkylation and ester-deprotection with LiOH to yield p-NO2-Bn-H4neunpa 6 (Scheme 1). The isothiocyanate (NCS) ana-logue for mAb conjugation, p-SCN-Bn-H4neunpa 9, was synthesized from the intermedi-ate 5 followed by nitro-reduction, ester-deprotection with LiOH and isothiocyanate formation with thiophosgene (Scheme 1).  Starting from the diethylenetriamine backbone, the two primary amines were protected with the 2-nitrobenzenesulfonyl groups to yield compound 1. Compound 1 is highly polar due to the two nosyl groups, thus a highly polar solvent like methanol is needed to separate it from the column. The second step is N-alkylation with 4-(2-bromoethyl)nitrobenzene. In order to maintain symmetry of the ligand, the ideal spot for bifunctionalization is the middle nitrogen. After that, N-alkylation with methyl-6-bromomethyl picolinate5 was performed to yield compound 3. The most challenging step was the nosyl-deprotection, constantly resulting in low yields of compound 4. The deprotected product is unfortunately highly polar and likely adsorbs on the surface of potassium carbonate, as seen by the red color of the salt. It was not possible to remove the large fractions of the deprotected product completely from the salt, which explains the low yield reported in the Experimental Section. Subsequently, alkyl halide N-alkylation was performed to yield product 5 with 71 % yield. p-NO2-Bn-H4neunpa 6 was synthesized in a final step of ester deprotection with LiOH. This compound was further used for radiolabeling experiments as well as potentiometric stability titrations. The 1H NMR spectrum of the final product is shown in Figure 2. p-SCN-Bn-H4neunpa 9 was synthesized starting from the intermediate 5 of the previous reaction route. Reduction of the nitro group with palladium on carbon yielded the amine-functionalized product 7.  The hydrolysis of the two tert-butyl esters and two methyl esters was performed differently from previous reports.5,12 Instead of acidic hydrolysis at high temperatures, compound 8 was synthesized by adding 10 eq. of lithium hydroxide to the reaction mixture at room temperature to yield the product, with a 50 % yield. The final step is the synthesis of the isothiocyanate-functionalized prod-uct 9. This was achieved by the reaction of excess thiophosgene with the aromatic primary amine to yield the final product with a 59 % yield. Overall, the synthesis of p-	18 SCN-Bn-H4neunpa from diethylenetriamine has a cumulative yield of 2.3 %, comparable to the overall synthesis yield of H5decapa (2.5 %).    Scheme 1.  Synthetic scheme for p-SCN-Bn-H4neunpa and p-NO2-Bn-H4neunpa.  NH2NH NH2SO2ClNO2NO2BrSHNa2CO3, THF K2CO3, DMFNa2CO3, DMFN OOBrK2CO3, THFNa2CO3, CH3CNOOBrPd/C 10%glacialCH3COOHLiOHTHF/H2O 3:1SCCl2H2O, DCMN N N OHOHOON NHO OHO ONCSN N N OOOON NO OO ONO2NH N HNN NO OO ONO2N N NN NO OO ONO2O2SO2SNO2 O2NNHN NHNO2O2SO2SNO2 O2NNHNHNHO2SO2SNO2 O2NN N N OHOHOON NHO OHO ONO2LiOHTHF/H2O 3:1N N N OOOON NO OO ONH2N N N OHOHOON NHO OHO ONH26 p-NO2-Bn-H4neunpa9 p-SCN-Bn-H4neunpa1234578	 19  3.2 Synthesis and characterization of non-radioactive metal complexes 3.2.1 NMR Three complexation experiments were performed with La3+, In3+ and Bi3+. 1H NMR spectra of the p-NO2-Bn-H4neunpa ligand precursor, and corresponding La and In complexes can be found in Figure 2. The [La(p-NO2-Bn-neunpa)]- complex shows 1H NMR upfield shifts of the alkyl-region; this effect has been previously observed in our group.13 The aromatic region is more resolved and shows a splitting of the peaks. Inte-gration of all peaks gives the same number of protons compared to the uncomplexed ligand. Furthermore, the HSQC spectra of this complex (Figure S2) shows the same number of carbons compared to the bare ligand, suggesting that there is only one iso-mer in solution. In contrast, the 1H NMR spectrum of [In(p-NO2-Bn-neunpa)]- shows more splitting in the aromatic and alkyl regions. The aromatic peaks are sharp and well resolved and integrating the peaks suggests one major static isomer. In addition, the COSY spectrum of this complex shows clear coupling of several peaks in the complex alkyl region (Figure S12), leading to the assumption there are fluxional isomers in solu-tion. Comparing these results to those with [In(decapa)]2-, which gave a complex 1H NMR spectrum with multiple isomers presumably due to several unbound carbox-ylates10, we can see an improvement in terms of isomerization by replacing one car-boxylate group with the functionalization arm on the middle nitrogen atom of the dieth-ylenetriamine backbone. Due to insolubility of the Bi complex, the 1H NMR spectrum cannot be used for proper assignments (Figure S1).  	20  Figure  2.  1H NMR spectra of A: p-NO2-Bn-H4neunpa (400 MHz, CDCl3, 25 °C); B: [La(p-NO2-Bn-neunpa)]- (400 MHz, CDCl3, 25 °C); C: [In(p-NO2-Bn-neunpa)]- (400 MHz, DMSO-d6, 25 °C).   3.2.2 IR Due to the insolubility of [Bi(p-NO2-Bn-neunpa)]-, an IR experiment on the solid was performed (Figure 3). Shifts of various peaks of the ligand itself compared to the Bi complex can be observed. The OH stretch at 2500 cm-1 disappeared after complexa-tion, suggesting that the carboxylic acids are bound to the metal ion; the carboxyl stretch at 1700 cm-1 disappeared as well, supporting this assumption. The two stretch-es of the nitro functional group (1500 cm-1 and 1400 cm-1) stayed the same. The stretch at 1200 cm-1 in the ligand spectra can be assigned as a C-N stretch that shifts to lower energies (1000 cm-1) when bound to the metal ion.   3.3	CD3OD3.3	CD3ODA:	ligandB:	La	complex2.42.62.83.03.23.43.63.84.04.24.44.64.85.05.25.45.65.86.06.26.46.66.87.07.27.47.67.88.08.28.4f1	(ppm)2.5	DMSO-d6C:	In	complex	 21  Figure  3.  IR spectra of p-NO2-Bn-H4neunpa and [Bi(p-NO2-Bn-neunpa)]-.  3.2.3 Thermodynamic Stability The extended diethylenetriamine backbone, along with the nonadentate N5O4 binding motif of H4neunpa, were specifically designed to accommodate binding of larger metal ions. As such, the protonation constants of H4neunpa as well as the stabil-ity constants of the respective La3+, Bi3+ and In3+ complexes were determined at 25 ºC in 0.16 M NaCl aqueous solution. The stepwise protonation constants (log K) obtained are presented in Table 1 together with protonation and stability constants reported for the related ligands H5decapa, H4octapa, DTPA and CHX-A”-DTPA. A straightforward comparison of the ability of different ligands to coordinate a specific metal ion (rather than the thermodynamic stability constants alone) is the conditional stability constant or pM value. pM is defined as (-log [Mn+]free) and is calculated at specific conditions ( [Mn+] = 1 µM, [Lx-] = 10 µM, pH 7.4 and 25 ºC), taking into consideration both metal-ligand association and ligand basicity. The protonation constants of the new synthe-sized ligand H4neunpa were determined by potentiometric titrations at pH 1.8-11.5 and by combined potentiometric-spectrophotometric titrations16,17 over the pH range 2.5-11.5.       800120016002000240028003200360040005060708090100cm-1Transmittance [%]p-NO2-Bn-H4neunpa[Bi(p-NO2-Bn-neunpa)]-	22   Table  1.  Stepwise Protonation Constants (log KHhL) of H4neunpa (25 ºC, I = 0.16 M NaCl)a a Literature data of related systems are presented for comparison. L = Ligand and charges of ligand species and metal complexes were omitted for simplicity. In Figure S3 are shown the sets of spectra obtained as a function of pH, at 7.18 x 10-4 M ligand concentration. The first and second protonation processes occur at the two terminal amines of the diethylenetriamine backbone (log K1 = 10.92(2) and log K2 = 9.29(2)), as suggested by the appearance of a single isosbestic point at 284 nm between pH 8.33 and 11.32 in the UV-potentiometric titration (Figure S3c). The third protonation process (log K3 = 6.79(2)) is assigned to the central nitrogen atom in the backbone and is supported by the appearance of an isosbestic point at 293 nm in the pH region between 5.39 and 8.33 (Figure S3b). The fourth and fifth protonation pro-cesses are attributed to the picolinate moieties13,18  (log K4 = 4.02(3) and log K5 = 2.97(2)). The UV-potentiometric titration showed also in this case a single isosbestic point at 296 nm for these protonation processes (Figure S3a).  The sixth protonation step is attributable to the carboxylic acid substituent (log K6 = 2.39(5)) and was calcu-lated from potentiometric titrations. The value of log K7 could not be determined, as the value was below the threshold of the electrode (pH < 2). H4neunpa, the bifunctional analogue of the previously reported H5decapa (for which we correct here the protona-tion constants, Table 1) presents overall fairly similar protonation constants, although the fourth and fifth protonation processes attributed to the picolinate moieties differ by 0.41 and 0.49 units respectively. The higher protonation constants in the case of H5decapa could be attributed to the higher negative charge of the ligand. The specia-equilibrium reaction neunpa4- (this work) decapa5- (this work) octa-pa4- 10 DTPA14 CHX-A’’-DTPA14 DOTA15 L + H+ ⇆ HL 10.92(2) 11.03(3) 8.59(4)  11.84 12.30 12.60(1) HL + H+ ⇆ H2L 9.29(2) 9.20(3) 5.59(6)  9.40 9.24 9.70(1) H2L + H+ ⇆ H3L 6.79(2) 6.86(4) 3.77(2) 4.85 5.23 4.50(1) H3L + H+ ⇆ H4L 4.02(3) 4.43(4) 2.77(4) 3.10 3.32 4.14(1) H4L + H+ ⇆ H5L 2.97(2) 3.46(5) 2.79(4) 2.20 2.18 2.32(1) H5L + H+ ⇆ H6L 2.39(5) 2.84(6) ND    H6L + H+ ⇆ H7L ND 2.52(4)     H7L + H+ ⇆ H8L  ND     	 23 tion plots for H4neunpa and H5decapa are shown in Figure S4 in the Supporting Infor-mation. Potentiometric titrations of H4neunpa were carried out in the presence of La3+, Bi3+, and In3+ in order to determine the stability constants of the corresponding metal complexes. For lanthanum, combined potentiometric-spectrophotometric titrations demonstrated that the complexation started from pH 2, based on the distinctive fea-tures of the spectra compared to the electronic spectra of H4neunpa (Figures S3 and S5). The thermodynamic stability of [La(neunpa)]- was determined to be log KML = 19.81(4) and pM = 16. This value is close to the values obtained for [La(octapa)]- log KML = 19.92(6)19 and [La(DTPA)]2- log KML = 19.4820. Similar to the free ligand, the depro-tonation of the [La(H2neunpa)]+ and La(Hneunpa) species is marked by the appearance of a single isosbestic point at 291 nm between the pH range 2.42-8.23 and suggests that the deprotonations occur at the two terminal amines of the diethylenetriamine backbone (Figure S5a). The [La(neunpa)]- species further deprotonates presumably due to the deprotonation of a coordinated water molecule with pK 9.78 to form the mono-hydroxo complexes (Figure S5b). Species distribution diagrams for the lanthanum(III) complexes of H4neunpa are plotted in Figure S6. The thermodynamic stability constant of the bismuth(III) complexes of H4neunpa could not be determined by direct potenti-ometric titrations as this requires the knowledge of the concentration of the free and bound metal ion at equilibrium, and even at pH 2 the Bi(III) complex was already signif-icantly formed. The ligand-ligand competition method using Na2H2EDTA as a known competitor was used to yield the stability constants presented in Table 2 and specia-tion plots in Figure S7. Particularly high thermodynamic stability of [Bi(neunpa)]- was found, log KML = 28.76(9) and pBi = 27. The thermodynamic stability constant of the [Bi(neunpa)]- complex is lower than those of [Bi(DTPA)]2- and [Bi(CHX-A-DTPA)]2- com-plexes14 and lower than that for [Bi(DOTA)]-; however, it is interesting to note that H4neunpa and DOTA have the same pBi3+ value of 27 (Table 2). 	 24 Table  2.   Stepwise Stability Constants (log K) of H4 neunpa complexes with La3+, Bi 3+ and In3+ a equilibrium reaction neunpa4- decapa5- 10 octapa4- DTPA CHX-A’’-DTPA DOTA La3+ + L ⇆ LaL 19.81(4)  19.92(6) 19  19.4820  22.021 LaL + H+ ⇆ LaHL 8.05(5)      LaHL + H+ ⇆ LaH2 L 3.28(6)      LaLOH + H+ ⇆ LaL 9.78(4)      Bi 3+ + L ⇆ BiL 28.76(9)   35.2(4) 14  34.9(4) 14  30.322 BiL + H+ ⇆ BiHL 10.26(5)      BiHL + H+ ⇆ BiH2 L 3.8(1)      BiLOH + H+ ⇆ BiL 10.57(7)      In3+ + L ⇆ InL 28.17(2) 27.56(5) 26.8(1) 10  29.023,24  23.9(1) 24  InL + H+ ⇆ InHL 5.07(2) 5.47(3) 2.9(2) 10     InHL + H+ ⇆ InH2 L 3.40(3) 2.73(6)     InLOH + H+ ⇆ InL 9.41(3) 9.83(7)         pLa3+ 16  19.7    pBi 3+ 27     2725 pIn3+ 23.6 23.1 26.510 25.710   18.810 a Literature data for related systems are presented for comparison. L = Ligand and charges of ligand species and metal complexes were omitted for simplicity. 	 25 Despite the high formation constant of [In(H2neunpa)]2+ log KMLH2 = 36.64(3), the system is well determined by direct potentiometric titration taking advantage of the indium-chloride competing species. The system as in the case of lanthanum(III) and bismuth(III) complexes containing MLH2, MLH, ML and ML(OH) complex species (Figure S8) presented a high log KML = 28.17(2) and pM = 23.6, which is significantly higher than for DOTA (Table 2), slightly higher than for the previously reported H5decapa, 2.1 pM units lower than for DTPA and 2.9 pM units lower than for [In(octapa)]-. To our knowledge thermodynamic formation constants of the [In(CHX-A”-DTPA)]2- have not been yet reported. It is noteworthy that, as with other previously reported ligands10, the trend of the stability constants and pM values and the human serum stability data do not correlate well, and despite the higher pM values for [In(octapa)]- species or [In(DTPA)]2- vs [In(neunpa)]-, [In(neunpa]- showed an exceptional serum stability 97.8(1) % after one day, 5.5 units higher than the [In(octapa)]- complex, 7.9 units higher than the [In(CHX-A”-DTPA)]2- complex and 9.5 units higher than the [In(DTPA)]2- complex.   3.3 Radiolabeling Experiments with Unmodified Chelators  The radiolabeling properties of 177Lu and 111In with H4neunpa were investigat-ed, and compared directly to results obtained for the gold-standards DOTA and CHX-A”-DTPA. Initial radiolabeling experiments revealed that p-NO2-Bn-H4neunpa could quan-titatively complex 111In3+ (radiochemical yield, RCY > 99%) in 10 minutes at room tem-perature (RT), pH 4, at ligand concentrations of 10-4 M. Subsequently, concentration-dependent labeling was performed by decreasing the ligand concentration 10-fold while keeping the 111In activity constant. Quantitative radiolabeling was achieved at ligand concentrations as low as 10-7 M (Figure 4), at 10 min and ambient temperature. At decreasing ligand concentrations of 10-8, 10-9, and 10-10 M, radiochemical yields gradually decreased to 71.1, 10.5, and 1.5%, respectively. These results demonstrate the ability of p-NO2-Bn-H4neunpa to rapidly and efficiently complex 111In in high specific activities at ambient temperatures. H4octapa showed similar radiolabeling efficiencies at 10-7 M, results at lower ligand concentrations are not reported.10 In sharp contrast to the two “pa” ligands is the macrocyclic gold-standard DOTA which is reported to re-quire heating samples at 100°C for 30 minutes to achieve high radiochemical yields.10 The acyclic chelator CHX-A”-DTPA is a relatively recent addition to the list of potential 111In chelators; in contrast to DOTA it can efficiently complex In3+ isotopes at ambient temperatures yet exhibits comparable in vivo stability to DOTA conjugates2,26, making it a more appealing chelator for radiolabeling of heat-sensitive biomolecules such as affibodies or antibodies.27-31 Our initial 111In radiolabeling studies with p-NH2-Bn-CHX-A”-DTPA at ligand concentrations of 10-4 M corroborate the efficient and mild labeling of this ligand which yielded RCYs >99%; however, two evident peaks in the HPLC radio-chromatogram are observed – one major product at 8.6 min and a minor product at 8.0 min (Figure S10), with the ratio between the major and minor product being 7.7. The 	26 appearance of two distinct peaks in the radio-chromatogram may indicate the for-mation of distinct 111In-chelate isomers. Contrary to H4neunpa, at p-NH2-Bn-CHX-A”-DTPA concentrations of 10-7 and 10-8 M, 111In labeling yield decreased to 75.0 and 3.4%, respectively. The ratio of major to minor product in the HPLC radio-chromatogram also changed drastically at lower ligand concentrations, with the ratio being close to unity (0.95) for 10-7 M labeling.    Figure 4.  Radiolabeling results of 111In-p-NO2-Bn-neunpa (10min, RT, pH 4). Unlike the facile labeling kinetics of [111In(p-NO2-Bn-neunpa)]-, initial radiolabel-ing studies with 177Lu were unsuccessful. Attempted 177Lu labeling at ligand concentra-tions of 10-4 M in 10 minutes at room temperature, pH 4 or 5.5, displayed a radiochemi-cal yield of 12.4%; heating the sample to 40 °C for 1 hour did not improve RCY. Con-versely, gold-standard DOTA was quantitatively radiolabeled (RCY > 99%) with 177Lu when heated to 40 °C for 1 hour at the same ligand concentration (10-4 M). The inability of p-NO2-Bn-H4neunpa to complex 177Lu isotopes at mild temperatures (< 40 °C) pre-cluded further study with this isotope, since it was immediately obvious from the initial results that H4neunpa was a poor match for 177Lu and presented no potential ad-vantage compared to the gold-standard DOTA.   3.4 Stability Studies with the Unmodified Chelators In order to probe the kinetic inertness of the [111In(p-NO2-Bn-neunpa)]- complex, a 5 d in vitro competition experiment was performed in the presence of human blood serum. Serum contains many endogenous ligands that can compete for In(III) binding in vivo, such as apo-transferrin and albumin, and any chelate-bound 111In must therefore 0.00010.001 0.01 0.1 1 10 100 1000050100150log [µM]RCY [%]	 27 be sufficiently stable to withstand transchelation to such proteins. The in vitro stability of [111In(p-NO2-Bn-neunpa)]- at 1 h, 1 and 5 d time points was tested alongside gold-standard [111In(p-NH2-Bn-CHX-A”-DTPA)]2- for comparison (Table 3). The [111In(p-NO2-Bn-neunpa)]- complex exhibited exceptional stability, remaining 97.8% intact over 5 days, while the [111In(p-NH2-Bn-CHX-A”-DTPA)]2- complex showed an initial ~8% drop in stabil-ity after 1 h and subsequently stabilized for 5 days to remain 90.1% intact. The initial drop in stability after 1 h may be due to the presence of two isomers in the labeling reaction of p-NH2-Bn-CHX-A”-DTPA (vide supra, major isomer 88.5% and minor isomer 11.5%). Studies with 88Y-CHX-DTPA have demonstrated that thermodynamic stability of the resultant metal complex can be significantly affected by the absolute configuration, possibly due to unfavourable steric hindrance of certain stereoisomers;32 therefore, it is feasible that the minor isomer is kinetically labile with respect to transchelation to serum proteins. Indeed, [111In(p-NO2-Bn-neunpa)]- displayed marginally higher stability than [111In(p-NH2-Bn-CHX-A”-DTPA)]2-, [111In(DOTA)]-, and [111In(octapa)]- after 1 d (97.8 ± 0.1%, 89.9 ± 0.6, 88.3 ± 2.2%, 92.3 ± 0.04%, respectively).    Table  3.  Human serum stability challenge data performed at 37°C  (n = 3), with stabil-ity shown as percentage of intact 111In-complex. Complex 1 h (%) 1 d (%) 5 d (%) [111In(p-NO2-Bn-neunpa)]- 97.9 ± 0.3 97.8 ± 0.1 97.8 ± 0.7 [111In(p-NH2-Bn-CHX-A”-DTPA)]2- 91.8 ± 1.8 89.9 ± 0.6 90.1 ± 0.9 [111In(octapa)]- a 93.8 ± 3.6 92.3 ± 0.04 NDb [111In(DOTA)]- a 89.6 ± 2.1 88.3 ± 2.2 NDb 111InCl3 (control) c 4.0 7.2 3.4 a Mouse serum stability data performed at ambient temperature. b ND = not deter-mined. c n = 1 only.; data included from ref10 for comparison  3.5 Initial Biodistribution Studies  Mouse biodistribution studies over the course of 24 hours (n = 4 each time point) were performed with [111In(p-NO2-Bn-neunpa)]- and [111In(p-NH2-Bn-CHX-A”-DTPA)]2- and the data are summarized in Table 4. Both In-complexes were rapidly ex-creted through the kidneys and activity cleared quickly from all other organs. Notably, uptake of [111In(p-NO2-Bn-neunpa)]- in the intestines was significantly higher than for [111In(p-NH2-Bn-CHX-A”-DTPA)]2- after 15 min (17.9 ± 5.5% ID/g vs 3.6 ± 1.6% ID/g) and 1 h (39.8 ± 2.9% ID/g vs 10.7 ± 1.4% ID/g). One explanation for the difference in intestine uptake is that the mono-anionic 111In-neunpa complex is more lipophilic than the di-	28 anionic 111In-p-NH2-Bn-CHX-A”-DTPA complex, as evinced by shifts in the radio-HPLC retention times (tR = 12.9 min and 8.6 min, respectively) and the absolute logP values of each complex (-1.65 ± 0.04, and -3.85 ± 0.17, respectively), thus shifting the excretion of the radiotracer from renal to intestinal elimination because highly charged polar sub-stances are generally eliminated via the kidneys while less hydrophilic compounds tend to be eliminated via the intestinal tract. Nonetheless, the remaining 111In-complex in the intestines at 1 h was rapidly excreted by 4 h for both complexes, and the uptake in intestines of [111In(p-NO2-Bn-neunpa)]- and [111In(p-NH2-Bn-CHX-A”-DTPA)]2- were no longer statistically different (p > 0.05) at later time points (0.265 ± 0.206% ID/g vs 0.160 ± 0.047% ID/g, for 4 h; 0.216 ± 0.114% ID/g vs 0.129 ± 0.06% ID/g, for 24 h, respective-ly). It has been suggested that administration of an unstable 111In-complex would result in demetalation of the complex in vivo and subsequent accumulation of transchelated or “free” 111In3+ activity in the liver, spleen, and bone over time;33 therefore, the rapid excretion of [111In(p-NO2-Bn-neunpa)]-  and [111In(p-NH2-Bn-CHX-A”-DTPA)]2- from these organs suggests both 111In-complexes are exceptionally robust and stable in vivo  (0.035 ± 0.008% ID/g vs. 0.023 ± 0.006% ID/g for liver; 0.029 ± 0.01% ID/g vs 0.032 ± 0.008% ID/g for spleen; 0.010 ± 0.006% ID/g vs 0.007 ± 0.002% ID/g for bone, at 24 h, respectively). Furthermore, [111In(p-NO2-Bn-neunpa)]- had improved kidney clearance compared to [111In(p-NH2-Bn-CHX-A”-DTPA)]2- at 24 h  (0.077 ± 0.058% ID/g vs 0.301 ± 0.043% ID/g, respectively, p <0.05). Although these initial biodistribution data appear promising it may be that the predicted -1 and -2 charge of the In-neunpa/-CHX-A”-DTPA complexes, respectively, at physiological pH, could be mediating the rapid elimination of the metal-complexes from the body;          	29 Table  4.   Decay corrected % ID/g values from biodistribution of 111In-complexes in healthy NOD.Cg-Prkdcscid Il2rgtm1Wjl/SzJ female NOD.Cg-Prkdcscid Il2rgtm1Wjl/SzJ mice (4 months old), n = 4. organ 15 min  1 h  4 h  24 h   [ 111In][In(p-NO2 -Bn-neunpa)] - Blood 1.979 (0.425)  0.077 (0.007)  0.022 (0.004)  0.0064 (0.0011)  Fat 0.174 (0.113)  0.009 (0.001)  0.0020 (0.0009)  0.0009 (0.0007)  Uterus 1.644 (0.321)  0.101 (0.011)  0.059 (0.072)  0.014 (0.005)  Ovaries 0.983 (0.362)  0.056 (0.034)  0.012 (0.011)  0.0080 (0.0067)  Intestine 17.941 (5.475)  39.760 (2.865)  0.265 (0.206)  0.216 (0.114)  Spleen 0.792 (0.379)  0.073 (0.024)  0.032 (0.027)  0.029 (0.010)  Liver 2.684 (0.190)  0.312 (0.090)  0.071 (0.016)  0.035 (0.008)  Pancreas 0.287 (0.196)  0.026 (0.006)  0.010 (0.006)  0.0047 (0.0023)  Stomach 1.251 (0.364)  0.054 (0.019)  0.012 (0.002)  0.062 (0.019)  Adrenal glands 0.585 (0.089)  0.037 (0.029)  0.012 (0.010)  0.0009 (0.0018)  Kidney 5.681 (1.343)  0.484 (0.322)  0.158 (0.105)  0.077 (0.058)  Lungs 2.695 (0.392)  0.388 (0.526)  0.072 (0.101)  0.056 (0.096)  Heart 0.419 (0.032)  0.075 (0.089)  0.011 (0.007)  0.0061 (0.0103)  Muscle 0.394 (0.101)  0.016 (0.004)  0.0030 (0.0018)  0.0020 (0.0016)  Bone 0.743 (0.351)  0.072 (0.029)  0.0099 (0.0069)  0.0102 (0.0060)  	 30 Brain 0.059 (0.033)  0.012 (0.002)  0.0013 (0.0006)  0.0009 (0.0016)  Tail 4.129 (2.183)  0.143 (0.095)  0.029 (0.023)  0.0078 (0.0060)   [ 111In][In(p-NH2 -Bn-CHX-A"-DTPA)] 2- [ 111In][In(p-NH2 -Bn-CHX-A"-DTPA)] 2- Blood 2.370 (0.221)  0.091 (0.035)  0.013 (0.014)  0.0011 (0.0003)  Fat 0.323 (0.070)  0.016 (0.007)  0.0037 (0.0014)  0.0024 (0.0017)  Uterus 1.643 (0.121)  0.116 (0.045)  0.082 (0.092)  0.035 (0.007)  Ovaries 1.279 (0.177)  0.077 (0.033)  0.024 (0.016)  0.0188 (0.0047)  Intestine 3.644 (1.632)  10.713 (1.428)  0.160 (0.047)  0.129 (0.060)  Spleen 0.627 (0.069)  0.074 (0.031)  0.036 (0.008)  0.032 (0.008)  Liver 3.388 (0.293)  0.271 (0.093)  0.053 (0.005)  0.023 (0.006)  Pancreas 0.539 (0.148)  0.036 (0.017)  0.014 (0.009)  0.0053 (0.0020)  Stomach 1.037 (0.115)  0.058 (0.025)  0.018 (0.003)  0.042 (0.030)  Adrenal glands 0.592 (0.174)  0.064 (0.048)  0.022 (0.003)  0.0156 (0.0043)  Kidney 7.643 (1.741)  1.152 (0.276)  0.632 (0.076)  0.301 (0.043)  Lungs 1.677 (0.227)  0.120 (0.045)  0.023 (0.003)  0.012 (0.002)  Heart 0.697 (0.089)  0.041 (0.013)  0.011 (0.001)  0.0069 (0.0011)  Muscle 0.500 (0.122)  0.022 (0.008)  0.0038 (0.0003)  0.0016 (0.0007)  Bone 0.717 (0.187)  0.057 (0.011)  0.0112 (0.0014)  0.0066 (0.0015)  Brain 0.063 (0.018)  0.017 (0.004)  0.0068 (0.0008)  0.0018 (0.0006)  Tail 3.562 (1.334)  0.349 (0.063)  0.410 (0.498)  0.0505 (0.0324)  	 31 Table  5.  Chemical and in vitro characterization data of 111In-neunpa -/- CHX-A''-DTPA-Trastuzumab radioimmunoconjugates Immunoconjugate 111In-neunpa-Trastuzumab 111In-CHX-A”-DTPA-Trastuzumab Radiolabeling conditions and yield pH 6, r.t., 15 or 30 min, 92.6 % pH 6, r.t., 30 min, 91.6% Chelate/mAb . 5.5 ± 1.1 4.6 ± 0.7 Specific activity (mCi/mg) 28.0 20.8 Immunoreactive fraction (%) >99 >99 Serum stability over 5 days (%) 94.7 % ND  therefore, the In-complexes may not have ample opportunity to dissociate in vivo giving the appearance of a stable complex. In order to further scrutinize the in vivo stability of 111In-neunpa and 111In-CHX-A”-DTPA an immuno-conjugate should be prepared (vide infra) and accordingly, biodis-tribution of each complex can be monitored over the course of several days instead of hours.  3.6 Preparation of Bioconjugates and In Vitro Characterization The promising radiolabeling efficiencies and in vitro kinetic inertness of [111In(p-NO2-Bn-neunpa)]- provided motivation to prepare and test the radiolabeling properties, and in vivo behaviour of the H4neunpa-bioconjugate. The HER2/neu-targeting antibody Trastuzumab was chosen as the biovector because it is well estab-lished to target HER2-expressing tumors such as the SKOV-3 ovarian cancer cell line. To provide a basis for comparison, the gold-standard CHX-A”-DTPA was also conjugat-ed to Trastuzumab and tested in parallel in the radiolabeling and in vivo experiments.  The novel bifunctional chelator p-SCN-Bn-H4neunpa 9 and gold-standard p-SCN-Bn-CHX-A”-DTPA were conjugated to Trastuzumab, by incubation at room temper-ature at 5:1 molar ratio of ligand to antibody under slightly basic conditions (pH 9.0).34 Final immunoconjugates were purified by spin filtration and stored at -20°C until use. A radiometric isotopic dilution assay was employed to determine the number of accessi-ble chelates per antibody; an average of 5.5 ± 1.1 H4neunpa chelates per antibody and 4.6 ± 0.7 CHX-A”-DTPA chelates per antibody were conjugated to Trastuzumab.  	32 Preliminary 111In radiolabeling efficiency of H4neunpa-Trastuzumab was test-ed at pH 5.0, 5.5, and 6.0 in NH4OAc buffer (0.15 M) at RT, and the radiochemical yield (RCY) was assessed at 15 min. Calculated RCYs after 15 min were 15.0, 84.4, 92.6% at pH 5.0, 5.5, or 6.0, respectively (Figure S11). RCY was also assessed after 90 min for pH 5.0 and 6.0 reactions; yields increased to 38% and remained constant at 92% for pH 5.0 and 6.0, respectively. These initial radiolabeling tests suggest an optimal radiolabeling pH of 6.0 for H4neunpa-Trastuzumab, in order to generate 111In-conjugates of high radi-ochemical yield (>90%) and purity in only 15 min at RT. This is in agreement with a solution equilibrium study, which reflects the maximum of the [In(neunpa)]- species formed at pH 6 (see distribution diagram in Figure S7). The kinetic inertness of 111In-neunpa-Trastuzumab was assessed in an in vitro human serum challenge assay at 37°C.  Much like the unconjugated precursor, 111In-neunpa-Trastuzumab was excep-tionally inert to transchelation when incubated with human serum, with 95.0 ± 1.1, 96.0 ± 2.5, 94.7 ± 0.6, and 94.8 ± 1.6% of the 111In-bioconjugate remaining intact after 1, 2, 5, and 7 days, respectively.  111In-labeled Trastuzumab conjugates were then prepared for in vivo studies. Both immunoconjugates were radiolabeled with 111In in NH4OAc buffer (0.15 M, pH 6) for 30 min at RT (Table 5), resulting in exceptionally high radiochemical yields (>90%) and radiochemically pure products (>99% after spin purification) for both 111In-neunpa-Trastuzumab and 111In-CHX-A”-DTPA-Trastuzumab. Final specific activities were de-termined to be 28.0 and 20.8 mCi/mg (1036 and 770 MBq/mg) for 111In-neunpa-Trastuzumab and 111In-CHX-A”-DTPA-Trastuzumab, respectively. In vitro cellular bind-ing assays with SKOV-3 cancer cells showed both 111In-immunoconjugates absolutely reactive towards the tested cell line (>99% immunoreactivity). Both 111In-immunoconjugates have thus the ability to still bind to HER2.   3.7 Biodistribution and SPECT/CT Imaging Studies In order to compare directly the pharmacokinetics of 111In-neunpa-Trastuzumab to 111In-CHX-A”-DTPA-Trastuzumab in vivo, biodistribution and single photon emission computed tomography (SPECT) in conjunction with helical X-ray CT imaging experiments were performed on female mice bearing subcutaneous SKOV-3 ovarian cancer xenografts on the left shoulder. Either tracer was injected via the tail vein (~37 MBq, ~35 – 50 µg, in 200 µL saline), and after 1, 3, and 5 days (n = 4 per time point) the mice were imaged (n = 2, Figure 4) and sacrificed to collect organs and tu-mors to be counted on a calibrated γ-counter.  SPECT/CT overlays of 111In-CHX-A”-DTPA-Trastuzumab and 111In-neunpa-Trastuzumab immunoconjugates are shown in Figure 5 at 1, 3 and 5 days post injec-tion. These images were corrected for decay to allow qualitative comparison for the two radiolabeled immunoconjugates. For 111In-CHX-A”-DTPA-Trastuzumab and 111In-neunpa-Trastuzumab, day 1 images show significant activity in the blood, the heart, the spleen and the tumor. The activity in the blood, the heart and the spleen decreases over 	 33 time. The 111In-CHX-A”-DTPA-Trastuzumab shows a higher activity in the tumor at all three time points, giving highly localized activity to the tumor site. On the other hand, 111In-neunpa-Trastuzumab shows a lower uptake of activity into the tumor at day one post injection. Over time, the activity in the tumor decreased to being barely visible after 5 days post injection. Activity in the tumors for the 111In-neunpa-Trastuzumab is still present at day 3 and 5 post-injection but in order to be able to compare the two tracers, an appropriate scale bar was required to prevent oversaturation of the high uptake of the 111In-CHX-A”-DTPA-Trastuzumab within tumors. Reducing the max value of the scale bar by a factor of 2.8 shows the remaining activity within the tumors for the 111In-neunpa-Trastuzumab (data not shown).  Figure 5.  SPECT/CT overlays of 111In-CHX-A”-DTPA-Trastuzumab (left) and 111In-neunpa-Trastuzumab immunoconjugates. Fused μSPECT/CT images in female mice with subcutaneous SKOV-3 xenografts on left shoulder, imaged at 1, 3 and 5 days post injection. Tumors are highlighted with arrows.  Comparing the biodistribution pattern of 111In-neunpa-Trastuzumab with 111In-CHX-A’’-DTPA-Trastuzumab, both tracer bioconjugates show the same general uptake profile, i.e. significant uptake in blood, spleen, liver, kidney, bone and tumor at day 1 (Figure 6 and Table S1). Three days and 5 days after immunoconjugate injection, the spleen and tumor still have the highest uptake of radiotracer compared to all other 	34 organs, but with significant difference (p < 0.01) between 111In-CHX-A”-DTPA-Trastuzumab and 111In-neunpa-Trastuzumab (49.65 ± 6.79 %ID/g for 111In-CHX-A’’-DTPA-Trastuzumab and 21.47 ± 6.61 %ID/g for 111In-neunpa-Trastuzumab after 5 days in the spleen and 59.14 ± 7.70 %ID/g for 111In-CHX-A’’-DTPA-Trastuzumab and 16.01 ± 2.24 %ID/g for 111In-neunpa-Trastuzumab after 5 days in the tumor). This distribution of antibody-linked tracer is well known and is due to the metabolism and circulation of antibodies (or antibody-chelate conjugates).35      Figure  6.  Biodistribution of 111In-CHX-A’’-DTPA-Trastuzumab compared to 111In-neunpa-Trastuzumab in specific organs. Data are expressed as mean ± SD (n=4). For statistical analysis * (p ≤ 0.05) and ** (p ≤ 0.01), two-way ANOVA. The blood, liver, kidney and bone show the lowest %ID/g regarding all the dif-ferent organs. The blood from 111In-neunpa-Trastuzumab treated mice is cleared faster than the gold-standard 111In-CHX-A’’-DTPA-Trastuzumab between 1 d and 3 d. Addition-ally, 111In-CHX-A’’-DTPA-Trastuzumab shows an increase in accumulation in the tumor over time, whereas 111In-neunpa-Trastuzumab shows a decrease of uptake into the tumor over time, which is consistent with the SPECT/CT overlay observations. Regard-ing the tumor:organ ratios (Figure 7), 111In-neunpa-Trastuzumab and 111In-CHX-A’’-DTPA-Trastuzumab show interestingly only significant different values 5 d after injec-tion for each ratio, tumor:blood, tumor:heart and tumor:muscle. Furthermore, the addi-tion of the several chelating ligands onto Trastuzumab (5.5 ± 1.1 H4neunpa chelates per antibody and 4.6 ± 0.7 CHX-A”-DTPA chelates per antibody) can modify the overall charge of the antibody. Specifically, one negative charge per [In(neunpa)]- complex and Blood Spleen Liver Kidney Bone Tumor020406080Organ%ID/g1d CHX-DTPA-trastuzumab3d CHX-DTPA-trastuzumab5d CHX-DTPA-trastuzumab1d neunpa-trastuzumab3d neunpa-trastuzumab5d neunpa-trastuzumab*********	 35 two negative charges per [In(CHX-A”-DTPA)]2- complex labeled to Trastuzumab is gen-erated; this induces a two-fold increase of negative charge on the CHX-A”-DTPA-Trastuzumab conjugates compared to neunpa-Trastuzumab conjugates, assuming an equal number of accessible chelates are occupied by In3+ in each immunoconjugate. Consequently, this variance in overall charge of the Trastuzumab conjugate might affect the biodistribution of the resultant 111In-tracer. The immunoreactivity results are comparable for H4neunpa- and CHX-A”-DTPA-Trastuzumab conjugates, showing that the reactivity between Trastuzumab and its receptor is not altered due to the structural modification post chelate-conjugation. We wonder if the stability of the Trastuzumab-receptor-complex might not be as stable because of the charge difference discussed before. This could lead to a decreased uptake into the cancer cells. To conclude from these observations, different pharmacokinetic mechanisms for 111In-neunpa-Trastuzumab and 111In-CHX-A’’-DTPA-Trastuzumab might take place after 5 days. The-se differences will be investigated further in order to fully understand the mechanism of tumor uptake.  The slightly inferior uptake for this radiometal-neunpa antibody conjugate is disappointing but the complete chemistry and biology results suggest strongly that H4neunpa is an attractive chelating ligand with a built In conjugatable moiety and should be investigated further with Bi3+ and in other In3+-biovector conjugates.   	36  Figure  7.  Tumor:Organ ratios of CHX-A’’-DTPA and neunpa. Data is expressed as mean ± SD (n=4). For statistical analysis ** (p ≤ 0.01), two-way ANOVA. 1d 3d 5d050100150200250timeRatio %T: Muscle (111In-CHX-A''-DTPA-Trastuzumab)T: Muscle (111In-neunpa-Trastuzumab)**1d 3d 5d0510152025timeRatio %T:Blood(111In-CHX-A''-DTPA-Trastuzumab)T:Blood(111In-neunpa-Trastuzumab)**1d 3d 5d020406080timeRatio %T: Heart (111In-CHX-A''-DTPA-Trastuzumab)T: Heart (111In-neunpa-Trastuzumab)**	 37 4 Summary The acyclic chelator p-NO2-Bn-H4neunpa and the bioconjugated analogue H4neunpa-Trastuzumab (5.5 ± 1.1 chelates per antibody) have been synthesized, char-acterized (HR-ESI-MS, 1H NMR, 13C NMR, 2D-HSQC and cold metal complexation stud-ies) and evaluated via radiolabeling with 111In and 177Lu. Unfortunately, low radiochemi-cal yields of p-NO2-Bn-H4neunpa with 177Lu were obtained (pH 4-5.5, ambient – 40°C, max. RCY 12.4 %). The radiolabeling yields of p-NO2-Bn-H4neunpa and H4neunpa-Trastuzumab with 111In were a great success, >99 % and 92.6 %, respectively. Human serum stability experiments revealed that the [111In(p-NO2-Bn-neunpa)]- complex and 111In-neunpa-Trastuzumab immunoconjugate were 97.8 and 94.7 % intact after 5 days, respectively. H4neunpa-Trastuzumab was highly immunoreactive (>99 %) as indicated by a cellular binding assay. Biodistribution study of [111In(p-NO2-Bn-neunpa)]- in mice showed higher uptake into the intestine within the first hours compared to [111In(CHX-A”-DTPA)]2- due to its higher lipophilicity. Small animal SPECT/CT imaging and biodis-tribution studies of 111In-neunpa-Trastuzumab were performed using female NOD.Cg-Prkdcscid Il2rgtm1Wjl/SzJ mice bearing SKOV-3 xenografts, and it was found that 111In-neunpa-Trastuzumab successfully identified the tumor from surrounding tissues and other organs. Compared to the gold-standard 111In-CHX-A’’-DTPA-Trastuzumab, our immunoconjugate showed slightly lower tumor uptake which decreased over time and a lower tumor:blood ratio after 5 days post injection, although high quality SPECT/CT images were obtained. A different pharmacokinetic behavior of both immunoconju-gates can be the result of different charges on the immunoconjugates. Thermodynamic stability experiments support these findings, since p-NO2-Bn-H4neunpa was found to bind strongly to large, highly charged metal ions like In3+, La3+ and Bi3+. Indeed, these results suggest H4neunpa as a strong Bi(III) chelator and, considering the higher 3.6 units pM value respect to its In(III) complex, it could be of interest for Bi(III) isotopes (212Bi and 213Bi) in targeted alpha therapy (TAT). These encouraging results suggest H4neunpa and its immunoconjugate have promise for studies with other radiometals and targeting vectors. These experiments are currently underway.   5 Experimental Materials and Methods All solvents and reagents were from commercial sources (Sigma Aldrich, TCI) and were used as received unless otherwise noted. p-NH2-Bn-CHX-A”-DTPA and p-SCN-Bn-CHX-A”-DTPA were purchased from Macrocyclics (Dallas, TX) and used as received. Human serum was purchased frozen from Sigma Aldrich. 1H and 13C NMR spectra were recorded at room temperature on a Bruker AV400 instrument; the NMR spectra are expressed on the δ (ppm) scale and are referenced to the residual solvent signal of the deuterated solvent. All spectra were recorded with sweep widths of 0-14 ppm or -20-	38 220 ppm for 1H and 13C NMR, respectively, and deviations in the presented spectra are magnifications for visualization purpose only. Assignments of the peaks in the NMR spectra are approximate. Mass spectrometry was performed on a Waters ZQ spec-trometer equipped with an electrospray source. The HPLC system used for purification of ligands and precursors consisted of a Waters 600 controller equipped with a Waters 2487 dual λ absorbance detector connected to a Phenomenex synergi hydro-RP 80Å 250mm x 21.1 mm semi-preparative column. Analysis of 111In and 177Lu radiolabeled chelate complexes was carried out using a Phenomenex Synergi 4 µ Hydro-RP 80 Å analytical column (250 mm x 4.60 mm 4 µm) using an Agilent HPLC system equipped with a model 1200 quaternary pump, a model 1200 UV absorbance detector (set at 250 nm), and a Raytest Gabi Star NaI(Tl) detector. The radiochemical purity and specific activity of the final 111In radioimmunoconjugates was determined by  using a size-exclusion chromatography (SEC) column (Phenomenex, BioSep-SEC-s-3000) on an Agilent HPLC system equipped with a model 1200 quaternary pump, a model 1200 UV absorbance detector (set at 280 nm), and a Bioscan (Washington, DC) NaI scintillation detector (the radiodetector was connected to a Bioscan B-FC-1000 flow-count system, and the output from the Bioscan flow-count system was fed into an Agilent 35900E interface, which converted the analog signal to a digital signal). Instant thin layer chromatography paper strips impregnated with silica gel (iTLC-SG, Varian) were used to analyze crude 111In-immunoconjugate labeling reactions and complex stability and counted on either a BioScan System 200 imaging scanner equipped with a BioScan Autochanger 1000 or on a Raytest miniGita with Beta GMC detector radio-TLC plate reader using TLC control Mini Ginastar software. PD-10 desalting columns (Se-phadex G-25 M, 50 kDa, GE Healthcare) and centrifugal filter units with a 50 kDa molec-ular weight cutoff (Ultracel-50: regenerated cellulose, Amicon Ultra 4 Centrifugal Filtra-tion Units, Millipore Corp.) were used for purification and concentration of antibody conjugates.  111InCl3 was cyclotron produced and provided by Nordion as a ~ 0.05 M HCl solution. 177LuCl3 was purchased from Perkin-Elmer and provided as a solution in dilute HCl.  Synthesis of compounds  N,N-(2-Nitrobenzensulfonamide)-1,2-triaminodiethane, 1 Diethylenetriamine (4.19 mL, 38.8 mmol) was dissolved in THF (240 mL) and cooled to 0°C. Sodium carbonate Na2CO3 (9.04 g, 85.3 mmol, 2.2 eq.) was added, fol-lowed by a slow addition of 2-nitrobenzensulfonyl chloride (18.9 g, 85.3 mmol, 2.2 eq.), causing the reaction mixture to turn pale yellow. The reaction mixture was stirred over-night at room temperature. The off-white mixture was filtered to remove sodium car-bonate and the filtrate was rotary evaporated to dryness. The crude product was puri-fied by silica chromatography (CombiFlash Rf automated column system 220 g HP 	 39 silica; solid (pause) preparation; A: hexanes, B: ethyl acetate, C: methanol, 100 % A to 100 % B gradient followed by 100 % C) to yield the product 1 as a yellow-orange solid (88 %, 16.15 g). 1H NMR (400 MHz, acetone-d6, 25°C): 8.13-8.11 (m, 2H), 7.94-7.89 (m, 6H), 3.11 (t, J= 7.32 Hz, 4H), 2.67 (t, J= 5.80 Hz, 4H). 13C NMR (101 MHz, acetone-d6, 25°C): 134.0, 132.7, 130.7, 125.0, 47.7, and 43.1. HR-ESI-MS calcd. for [C16H19N5O8S2+H]+: 474.0753; found 474.0749 [M+H]+.  N,N-(((4-Nitrophenyl)azanediyl)bis(ethane-2,1-diyl))bis(2-nitrobenzenesulfonamide), 2 To a solution of 1 (16.15 g, 34.1 mmol) in DMF (60 mL) was added K2CO3 (6.13 g, 44.3 mmol, 1.3 eq.) and 4-(2-bromoethyl)nitrobenzene (10.20 g, 44.3 mmol, 1.3 eq.). After stirring the reaction mixture for 3 days at 40°C, the bright yellow solution was cooled to room temperature and the excess K2CO3 was removed by centrifugation. After drying the solution in vacuo, the crude dark red product was purified by silica chromatography (Combi Flash Rf automated column system; 80 g HP silica; solid (pause) preparation; A: hexane, B: ethyl acetate, 100% A to 100% B gradient) to yield product 2 as an orange fluffy solid (64.0 %, 13.59 g). 1H NMR (400 MHz, CDCl3, 25°C): 8.11-8.09 (d, J= 8.58 Hz, 2H), 8.08-8.06 (m, 2H), 7.84-7.81 (m, 2H), 7.76-7.73 (m, 4H), 7.32-7.30 (d, d= 8.58 Hz, 2H), 5.68 (s, 2H, NH), 3.07-3.05 (t, d= 5.63, 4H), 2.86-2.82 (t, J= 6.88, 2H), 2.74-2.72 (m, 2H), 2.70-2.67 (t, J= 6.62 Hz, 4H). 13C NMR (101 MHz, CDCl3, 25°C): 158.0, 157.6, 147.5, 146.8, 129.6, 129.5, 124.0, 117.3, 114.4, 55.0, 52.5, 37.6, and 33.5. HR-ESI-MS calcd. for [C24H26N6O10S2+H]+: 623.1230; found 623.1237 [M+H]+.  Dimethyl-6,6-(((((4-nitrophenethyl)azanediyl)bis(ethane-2,1-diyl))bis(((2-nitrophenyl)sulfonyl)azanediyl)) bis(methylene))-dipicolinate, 3 To a solution of 2 (13.59 g, 21.8 mmol) in dry DMF (80 mL) was added methyl-6-bromomethyl picolinate (11.55 g, 50.2 mmol, 2.3 eq) and sodium carbonate (5.32 g, 50.2 mmol, 2.3 eq). The bright orange reaction mixture was stirred at 60°C overnight, filtered to remove excess sodium carbonate, and concentrated in vacuo. The crude product was purified by silica chromatography (CombiFlash Rf automated column system; 2 x 80 g silica; solid (pause) preparation; A: hexane, B: ethyl acetate, 100% A to 100% B gradient) to yield product 3 as an orange/brown oil (70 %, 14 g). 1H NMR (400 MHz, CDCl3, 25°C): 8.02-8.00 (m, 4H), 7.96 (d, J = 7.7 Hz, 2H), 7.78 (t, J = 7.8 Hz, 2H), 7.67-7.60 (m, 6H), 7.54 (d, J = 7.8 Hz, 2H), 7.18 (d, J = 8.4 Hz, 2H), 4.67 (s, 4H), 3.89 (s, 6H), 3.29 (t, J = 6.8 Hz, 4H), 2.58-2.51 (m, 8H). 13C NMR (101 MHz, CDCl3, 25°C): 165.3, 157.0, 148.1, 147.6, 146.4, 138.1, 132.1, 129.7, 126.0, 125.9, 124.4, 124.4, 123.5, 55.3, 53.9, 52.9, 52.7, 46.9, and 33.4. HR-ESI-MS calcd. for [C40H40N8O14S2+H]+: 921.2184; found 921.2184 [M+H]+.  	40 Dimethyl-6,6-(((((4-nitrophenethyl)azanediyl)bis(ethane-2,1-diyl)bis(azanediyl))bis(methylene))dipicolinate, 4 To a solution of 3 (7.48 g, 8.1 mmol) in dry THF (100 mL) was added thiophe-nol (1.91 mL, 18.7 mmol, 2.3 eq.) and potassium carbonate (3.71g, 26.8 mmol, 3.3 eq.). The reaction mixture was stirred at 50°C for 72 hours, changing color to light orange. The excess salts were removed by centrifugation (5 min, 4000 rpm) followed by several washes with DMF. The filtrate was concentrated in vacuo in a (maximum) 50°C water-bath temperature. The resulting crude dark orange oil was purified by neutral alumina chromatography (CombiFlash Rf automated column system; 6 x 40 g neutral alumina; liquid injection A: dichlormethane, B: methanol, 100 % A to 20 % B gradient) to yield product 4 as an orange oil (32.4 %, 1.45 g). 1H NMR (400 MHz, CDCl3, 25°C): 8.01 (d, J = 8.6 Hz, 2H), 7.90 (d, J = 7.6 Hz, 2H), 7.73 (t, J = 7.8 Hz, 2 H), 7.45 (d, J = 7.7 Hz, 2H), 7.28 (d, J = 8.6 Hz, 2H), 3.99 (s, 4H), 3.91 (s, 6H), 2.79-2.72 (m, 12H). 13C NMR (101 MHz, CDCl3, 25°C): 165.6, 158.9, 148.6, 147.4, 164.4, 137.8, 129.7, 126.0, 123.9, 123.7, 55.9, 54.0, 53.0, 52.7, 47.0, and 33.3. HR-ESI-MS calcd. for [C28H34N6O6+H]+: 551.2618; found 551.2617 [M+H]+.  N,N-[(tert-Butoxycarbonyl)methyl-N,N-[6(methoxycarbonyl)pyridine-2-yl]methyl]-N-(4-nitrophenethyl)-1,2-triaminodiethane, 5 To a solution of 4 (1.45 g, 2.6 mmol) in acetonitrile (60 mL) was added tert-butylbromoacetate (894 µL, 6.1 mmol, 2.3 eq.) and sodium carbonate (642 mg, 6.1 mmol, 2.3 eq.). The reaction mixture was stirred at 60°C overnight, filtered to remove excess sodium carbonate and concentrated in vacuo.  The crude product was purified by silica chromatography (CombiFlash Rf automated system; 40g HP silica; A: di-chloromethane, B: methanol, 100% A to 20% B gradient) to yield product 5 as an orange oil (72 %, 1.48 g). 1H NMR (400 MHz, CDCl3, 25°C): 8.11 (d, J = 8.5 Hz, 2H), 8.03 (d, J = 7.7 Hz, 2H), 7.89 (t, J = 7.7 Hz, 2H), 7.52 (d, J = 7.6 Hz, 2H), 7.43 (d, J = 8.5 Hz, 2H), 4.18 (s, 4H), 3.96 (s, 6H), 3.90 (s, 4H), 3.73 (m, 2H), 3.54 (s, 4H), 3.45 (br s, 4H), 3.24 (m, 2H), 1.38 (s, 18H). 13C NMR (101 MHz, CDCl3, 25°C): 168.7, 165.1, 156.7, 147.3, 147.3, 143.8, 139.1, 130.1, 127.5, 125.0, 124.0, 83.2, 57.5, 56.0, 54.5, 53.3, 50.4, 48.8, 29.8, 28.0. HR-ESI-MS calcd. for [C40H54N6O10H]+:779.3980; found 779.3973 [M+H]+.  p-NO2-Bn-H4neunpa · 2.2 HCl · 3.1 H2O, 6 To compound 5 (0.23 g, 0.3 mmol) in THF/H2O (3 mL, 3:1) was added lithium hydroxide (0.07 g, 3.0 mmol, 10 eq.) and the mixture was stirred for 16 h at room tem-perature. Solvents were evaporated and the crude product was purified by semi-preparative reverse-phase (RP) HPLC (10mL/ min, gradient A: 0.1% TFA in deionized water, B: acetonitrile, A: 95% to B: 100% for 25 min., tR= 14.00 min) and the product 6 was obtained as a yellow oil (61 %, 0.12 g). 1H NMR (400 MHz, CDCl3, 25°C): 8.11 (d, J = 8.5 Hz, 2H), 8.03 (d, J = 7.7 Hz, 2H), 7.89 (t, J = 7.7 Hz, 2H), 7.52 (d, J = 7.6 Hz, 2H), 7.43 	 41 (d, J = 8.5 Hz, 2H), 4.18 (s, 4H), 3.90 (s, 4H), 3.73 (m, 2H), 3.54 (s, 4H), 3.45 (br s, 4H), 3.24 (m, 2H). 13C NMR (101 MHz, CDCl3, 25°C): 168.7, 165.1, 156.7, 147.3, 147.3, 143.8, 139.1, 130.1, 127.5, 125.0,124.0, 57.5, 56.0, 54.5, 48.8. HR-ESI-MS calcd. for [C30H34N6O10+H]+: 639.2415; found 639.2415 [M+H]+. Elemental analysis: calcd % for p-NO2-Bn-H4neunpa · 2.2 HCl ·3.1 H2O: C 46.55 N 10.86 H 5.2; found: C 46.72, N 10.64, H 5.37.  N,N-[(tert-Butoxycarbonyl)methyl-N,N-[6(methoxycarbonyl)pyridine-2-yl]methyl]-N-(4-aminophenethyl)-1,2-triaminodiethane, 7 Compound 5 (0.11 g, 0.1 mmol) was dissolved in glacial acetic acid (3 mL) and Pd/C 10% was added, the vessel sealed and purged with H2 gas, charged with a H2 balloon and left to stir for 2 h at room temperature. The reaction mixture was then fil-tered through Celite and concentrated under reduced pressure to yield compound 7. The aromatic amine was confirmed by a purple ninhydrin staining. The solution was filtered and the filtrate was concentrated in vacuo. 1H NMR (400 MHz, MeOD, 25°C): 7.99 (m, 2H), 7.92 (t, J = 7.9 Hz, 2H), 7.62 (d, J = 7.7 Hz, 2H), 6.87 (d, J = 7.9 Hz, 2H), 4.01 (s, 2H), 3.46 (br.4, 2H), 3.13 (m, 4H), 2.79 (m, 4H), 1.41 (s, 18H). 13C NMR (400 MHz, MeOD): 172.3, 166.7, 160.8, 148.3, 139.7, 139.5, 130.4, 128.3, 125.4, 125.2, 116.8, 82.7, 59.4, 56.9, 53.4, 52.3, 50.6, 29.9, 28.4. HR-ESI-MS calcd. for [C40H56N6O8+H]+: 749.4238; found 749.4236 [M+H]+.  p-NH2-Bn-H4neunpa, 8 Compound 6 (0.09 g, 0.13 mmol) was dissolved in THF/H2O (3 mL, 3:1) and lithium hydroxide (0.03 g, 1.26 mmol, 10 eq.) was added. The reaction mixture was left at room temperature for 24 hours. After product formation was confirmed by ESI-MS analysis, the solution was neutralized with 1 M HCl and solvents were concentrated in vacuo. For purification, semi-preperative RP-HPLC (10mL/ min, gradient A: 0.1% TFA in deionized water, B: acetonitrile, A: 95% to B: 100% for 25 min., tR= 11.50 min) was used and product 8 was obtained as a yellow oil (50 %, 0.04 g). 1H NMR (400 MHz, MeOD, 25°C): 8.05-8.04 (d, J = 6.6 Hz, 2H), 7.96-7.94 (d, J = 5.8 Hz, 2H), 7.63-7.61 (d, J = 6.6 Hz, 2H), 7.42-7-40 (d, J = 5.8 Hz, 2H), 7.32 (s, 2H), 4.08 (s, 4H), 3.71 (s, 4H), 3.59 (s, 2H), 3.53 (s, 4H), 3.35 (s, 4H), 3.14 (m, 2H). 13C NMR (400 MHz, MeOD): 173.5, 167.4, 159.0, 148.7, 140.3, 139.1, 131.8, 128.3, 125.6, 124.4, 116.7, 58.7, 56.5, 55.8, 53.1, 50.1, 30.4; 13C-DEPT NMR (400 MHz, MeOD): 140.3↑, 131.5↑, 128.1↑, 125.6↑, 124.2↑, 58.4↓, 56.4↓, 55.5↓, 51.5↓, 49.9↓, 30.2↓. HR-ESI-MS calcd. for [C30H37N6O8+H]+: 609.2673; found 609.2671 [M+H]+.  p-SCN-Bn-H4neunpa, 9 Compound 8 (0.04 g, 0.1 mmol) was dissolved in 0.1 M HCl (1 mL) and di-chloromethane (1 mL). Thiophosgene (0.05 mL, 0.6 mmol, 10 eq) was added and the 	42 solution was stirred vigorously at room temperature overnight in the dark. The solvents were concentrated in vacuo and the product purified by semi-preparative RP-HPLC (10mL/ min, gradient A: 0.1% TFA in deionized water, B: acetonitrile, A: 95% to B: 100% for 25 min., tR= 17.00 min) to yield product 9 as an orange oil (59 %, 0.02 g). 1H NMR (400 MHz, MeOD, 25°C): 8.04-8.02 (d, J = 6.7 Hz, 2H), 7.95-7.92 (t, J = 8.2 Hz, 2H), 7.61-7.57 (t, J = 7.5 Hz, 2H), 7.27-7-25 (d, J = 7.5 Hz, 2H), 7.18-7.16 (d, J = 7.5 Hz, 2H), 4.05 (s, 4H), 3.66 (s, 4H), 3.55 (s, 2H), 3.50 (s, 4H), 3.26 (br s, 4H), 3.07 (m, 2H). 13C NMR (400 MHz, MeOD): 173.5, 167.4, 159.0, 148.7, 140.3, 137.5, 131.5, 128.3, 126.9, 125.6, 115.9, 58.7, 56.5, 55.8, 51.9, 50.1, 30.5; 13C-DEPT NMR (400 MHz, MeOD): 140.3↑, 131.4↑, 128.3↑, 126.9↑, 125.6↑, 58.7↓, 56.35↓, 56.5↓, 51.9↓, 50.1↓, 30.5↓. HR-ESI-MS calcd. for [C31H35N6O8+H]+: 651.2237; found 651.2239 [M+H]+.  Na[La(p-NO2-Bn-neunpa)]  Compound 6 (10.2 mg, 16.0 mmol) was dissolved in water and lanthanum per-chlorate (7.7 mg, 17.6 mmol, 1.1 eq.) was added. The pH was adjusted to 4 using 0.1 M NaOH. The successful La-complexation as a white precipitate was confirmed by HR-ESI-MS immediately after adding La(ClO)4. After centrifugation, the precipitate was washed with water. 1H NMR (400 MHz, DMSO-d6, 25°C): 8.15 (d, 2H), 8.01 (m, 2H), 7.92 (m, 2H), 7.61 (d, J = 7.6 Hz, 2H), 7.50 (d, J = 8.5 Hz, 2H), 4.01 (s, 4H), 3.69 (m, 5H), 3.51 (d, 4H), 3.25 (s, 7H). HSQC (400 MHz, DMSO-d6, 25°C) in Supporting Information. HR-ESI-MS calcd. for [C30H32N6O10La]+: 775.1243; found 775.1236 [M+2H]+.  Na[Bi(p-NO2-Bn-neunpa)]  Compound 6 (20.3 mg, 31.8 mmol) was dissolved in water and bismuth tri-chloride (11.0 mg, 35.0 mmol, 1.1 eq.) was added. The pH was adjusted to 4 using 0.1 M NaOH. The successful Bi-complexation as a white precipitate was confirmed by HR-ESI-MS immediately after adding BiCl3. After centrifugation, the precipitate was washed with water. The Bi-complex is not soluble in any solvent; DMSO-d6 was chosen for NMR analysis. 1H NMR and 13C NMR not measurable due to solubility problems. HR-ESI-MS calcd. for [C30H30N6O10Bi]+: 843.1827; found 843.1835 [M+2H]+.  Na[In(p-NO2-Bn-neunpa)]  In a 20 mL screw cap vial, compound 6 (12 mg, 0.019 mmol) was dissolved in H2O:MeOH (2:1, 1.5 mL). In a separate screw cap vial, [In(ClO4)3]·8H2O (32 mg) was dissolved in dist. water (0.5 mL) to make a stock solution (64 mg/mL). An aliquot (217 μL, 13.8 mg, 0.0249 mmol) of this In(III) stock solution was added to the chelate solu-tion. The pH of the solution was adjusted from pH 1 to pH 5 using 1 N NaOH and 0.1 M HCl. A stir bar was added, the reaction heated to 60°C in a sand bath and stirred for 3 hours with the lid loosely on. The mixture was removed from the heat and allowed to cool to room temperature. A white precipitate had formed, and the solution was then 	 43 centrifuged and washed with dist. water (5 x 1 mL). After drying under high vacuum, the product as a white solid was collected (4 mg, 0.0053 mmol) with an overall yield of 28%. 1H, and COSY NMR (400 MHz, DMSO-d6) potential multiple isomers in solutions, see Figure S12 Supporting Information. HR-ESI-MS calculated for [115InC30H30N6O10+H+Na]+: 773.1038; found: 773.1039 (M+H+Na)+.  Bioconjugation of p-SCN-Bn-H4neunpa and p-SCN-Bn-CHX-A”-DTPA to Trastuzumab.  Trastuzumab (Herceptin, Genentech, San Francisco, CA, USA) was purified us-ing size exclusion columns (PD-10 desalting columns) and centrifugal filter units with a 50 kDa molecular weight cutoff and phosphate buffered saline (PBS, pH 7.4) to remove α-α-trehalose dehydrate, L-histidine, and polysorbate 20 additives. The purified antibody was brought up in PBS at pH 7.4. For each chelate-antibody conjugation, PBS (905 µL, pH adjusted to 9.0 using 0.1 M Na2CO3) and Trastuzumab (Genentech, San Francisco, CA, USA) (4 mg, 75 µL in PBS pH 7.4) was added to a low protein binding Eppendorf tube. To the antibody mixture, 5 equivalents of p-SCN-Bn-H4neunpa or p-SCN-Bn-CHX-A”-DTPA was added, respectively, in small portions (5 x 5 µL in DMSO). The reaction mixture was stirred at ambient temperature overnight, and subsequently purified by centrifugal filtration. The final bioconjugates were stored in 0.25 M sodium acetate at -20°C. Final protein concentration was determined by the Bradford assay.  Chelate number – radiometric isotopic dilution assay.  The number of accessible chelating ligands conjugated per antibody was de-termined using previously described methods.36,37 Briefly, a 1 µCi/uL [111In]InCl3 working solution (non-radioactive In3+ spiked with 111In) was prepared with a final In3+ concen-tration of 500 µM in ammonium acetate buffer (0.15 M, pH 6). In duplicate, for each chelate-antibody conjugate, 50 µg of bioconjugate (30 µL) was prepared into separate 1.5 mL Eppendorf tubes. Aliquots of 20, 25, and 30 µL of the [111In]InCl3 working solu-tion were added to the two chelate-antibody samples. Positive controls containing 50 µg of bioconjugate and 25 µL of buffered 111InCl3 only (no non-radioactive In3+ added) were prepared in duplicate. Negative controls containing 30 µL of PBS, and 25 µL of [111In]InCl3 working solution were prepared in duplicate. Samples were allowed to incu-bate at room temperature overnight, after which time EDTA (50 mM, pH 5) was added at 1/9 of the reaction volume to scavenge any unspecifically bound In3+, and incubated for 15 min. Each reaction mixture was spotted onto iTLC-SG plates, and developed using EDTA (50 mM, pH 5) as mobile phase. Radioactivity on the plate was measured using a radio-TLC plate reader, and number of chelates attached per antibody was calculated using equation (1). !"#$ !ℎ!"#$! =  # !!"#$% !" !"#$%&'$ (!!!!.!)!"!#$ # !"#$%& ×!"#$ !"!!     (1) 	44  111In-Chelate Radiolabeling Studies The ligand p-NO2-Bn-H4neunpa, or gold standard p-NH2-Bn-CHX-A”-DTPA, was made up as a stock solution (1 mg/mL, ~10-3 M) in deionized water. From this stock solution, serial dilutions were prepared to final ligand concentrations of 10-4 M – 10-9 M.  A 100 µL aliquot of each ligand stock (10-3 to 10-9 M) or water (blank control) was add-ed to screw-cap mass spectrometry vials and diluted with sodium acetate buffer (pH 4, 10 mM, 880 µL). An aliquot of diluted 111In stock (20 µL, ~200 µCi) was added to each vial and allowed to radiolabeled at ambient temperature for 10 min, then it was ana-lyzed by RP-HPLC to confirm radiolabeling and calculate yields. For human serum sta-bility studies, undiluted 111InCl3 stock (~20 µL, 5 mCi) was added to the reaction vial containing 10-4 M ligand in sodium acetate buffer. Areas under the peaks observed in the HPLC radio-trace were integrated to determine radiolabeling yields. Elution condi-tions used for RP-HPLC analysis were gradient: A: 0.1% trifluoroacetic acid (TFA) in water, B: acetonitrile; 0 to 100% B linear gradient 20 min, 1 mL/min. [111In(p-NO2-Bn-neunpa)]- (tR = 12.9 min), [111In(p-NH2-Bn-CHX-A”-DTPA)]2- (tR = 8.0 min (minor product); 8.6 min (major product))  “111In3+” (tR = 5.3 min).  Partition Coefficients 111In-labeled complex (30 µL, 20 µCi) was diluted with phosphate buffered sa-line (pH 7.4, 470 µL), and added to 1-octanol (500 µL) in a 1.5 mL Eppendorf tube. Sam-ples were vortexed for 60 seconds and subsequently centrifuged to separate phases (3000 rpm, 5 min). Aliquots (490 µL) of the aqueous and organic phases were diluted in a standard volume (20 mL) of water or acetonitrile, respectively, for measurement in an N-type Co-axial HPGe gamma spectrometer from Canberra fitted with a 0.5 mm berylli-um window and calibrated (energy and efficiency) with a 20 mL 152Eu and 133Ba source. The samples were counted for a minimum of 5 minutes, with a dead time less than 5 %. The amount of 111In-complex  (Bq) in each fraction was quantified using the 171 and 245 keV gamma lines of 111In.   111In-neunpa/CHX-A”-DTPA-Trastuzumab Radiolabeling for In Vivo Studies Aliquots of H4neunpa/CHX-A”-DTPA-Trastuzumab (650 µg) were diluted with ammonium acetate buffer (0.15 M, pH 6) such that the final volume of the reaction was 1 mL, and then 111InCl3 (~20 mCi) was added. The mixtures were allowed to react at ambient temperature for 40 min, and then analysed via iTLC-SG using 50 mM EDTA (pH 5) as eluent; 111In-labelled antibody remained at the baseline, while 111In3+ ions com-plexed as 111In-EDTA and eluted with the solvent front. Radiolabeled immunoconju-gates were then purified by PD-10 SEC columns and centrifugal filtration (50k cut-off). The radiochemical purity of the final radiolabeled bioconjugates was determined using 	 45 SEC-HPLC (using an isocratic gradient of 0.1 M sodium phosphate monobasic dehy-drate, 0.1 M sodium phosphate dibasic dodecahydrate, 0.1 M sodium azide and 0.15 M sodium chloride (pH 6.2-7.0)); the specific activity was calculated by injecting a known activity, and integrating areas under the peaks of the UV-chromatogram measured against a standard curve.   In Vitro Human Serum Stability Data  The procedures of the serum competition studies followed closely those pre-viously published.8,10 The compound [111In(p-NO2-Bn-neunpa)]-, [111In(p-NH2-Bn-CHX-A”-DTPA)]2-, or blank control “111In3+” was prepared using the radiolabeling protocol as described above. In triplicate for the 111In-complex, solutions were prepared in vials containing 330 µL of 111In-complex (~1.6 mCi), 1000 µL of room temperature human serum, and 670 µL of phosphate buffered saline (PBS, pH 7.4) and incubated at 37°C. At time points of 1 h, 1 and 5 days, 400, 400, and 800 µL aliquots of the human serum competition mixture were removed from each vial, respectively, diluted to a total vol-ume of 2.5 mL with PBS, and counted in a Capintec CRC-55tR dose calibrator; this value is recorded as “full activity” to be loaded onto the PD-10 column. The 2.5 mL of reaction mixture was loaded onto a pre-conditioned PD-10 column, and the empty vial was counted again in the dose calibrator – this value was recorded as “residual activi-ty” left in the vial. The loaded effluent was collected in a waste container, and then the PD-10 column was eluted with 3.5 mL of PBS, and collected into a separate vial. The eluent that contained 111In bound/associated with serum proteins (size exclusion for MW < 5000 Da) was counted in the dose calibrator and then compared to the total activity that was loaded onto the PD-10 column to obtain the percentage of 111In that was bound to serum proteins and therefore no longer chelate-bound by the relation-ship: 1 – (eluted activity/(full activity – residual activity) x 100. For serum stability of the radioimmunoconjugates, 111In-neunpa-Trastuzumab was first prepared as de-scribed above by incubating H4neunpa-Trastuzumab (200 µg) and 111InCl3 (~1 mCi) in NH4OAc (0.15 M, pH 6) for 20 minutes at ambient temperature. After confirming a ra-diolabeling yield >95% via iTLC-SG in 50 mM EDTA (pH 5), in triplicate, 111In-neunpa-Trastuzumab (330 µL, ~320 µCi) was incubated with human serum (330 µL) and the mixture was left at 37°C for 168 h (7 d). At time points of 0, 1, 24, 48, 120, and 168 h, aliquots (3 – 5 µL) of the competition mixture was spotted on iTLC-SG plates and de-veloped in 50 mM EDTA (pH 5) as described above.   Cell Culture The human ovarian adenocarcinoma HER2-positive SKOV-3 cells were cul-tured at 37°C with 5% CO2 in Dulbecco's modified Eagle's medium (DMEM, LifeTechnol-ogies, Rockford, IL, USA) containing 2 mM glutamine and supplemented with 10% fetal 	46 bovine serum (Sigma-Aldrich, Oakville, ON, USA) and 100 U/mL of penicillin-streptomycin (LifeTechnologies).  In Vitro Immunoreactivity Assay  The immunoreactive fractions of 111In-neunpa-Trastuzumab and 111In-CHX-A”-DTPA-Trastuzumab were determined according to the Lindmo cell-binding method38 using SKOV-3 cells37,39 as previously described. Briefly, cells were suspended at 0.23 to 2.3 x 106 cells/mL in PBS (pH 7.4). For each tested antibody, 50 μL (from a solution of 0.45 µg of each radioimmunoconjugate diluted in 5 mL of 1% PBS-BSA) was added to each cell concentration tube in duplicate. The radiolabeled immunoconjugates were incubated for 1 h at 37 °C and under gentle agitation. Cells were then pelleted and washed twice with PBS. Each cell-bound activity for the different cell conditions was determined by measuring the 111In amount of activity within the cell pellets using the Wallac WIZARD2 gamma counter with background and decay correction. The bound fraction was determined as a percentage of total added activity according to control samples. Immunoreactive fractions were estimated for conditions representing infinite antigen excess by linear regression analysis of a plot of total/bound activity against 1/[cell concentration]. Results >80% were considered suitable for in vivo imaging.   SKOV-3 Xenograft Mouse Models All experiments were conducted in accordance with the guidelines established by the Canadian Council on Animal Care and approved by the Animal Ethics Committee of the University of British Columbia (protocol # A16-0104). Female NOD.Cg-Prkdcscid Il2rgtm1Wjl/SzJ mice (4 months old) obtained from an in-house breeding colony were subcutaneously injected with 8 x 106 SKOV-3 cells in matrigel (BD Bioscience) on the left flank.  [111In(p-NO2-Bn-neunpa)]- and [111In(p-NH2-Bn-CHX-A”-DTPA)]2- in vivo Biodistribution [111In(p-NO2-Bn-neunpa)]- and [111In(p-NH2-Bn-CHX-A”-DTPA)]2- were prepared according to the radiolabeling protocol above using 10-4 M ligand and ~ 148 MBq (~4 mCi) of 111InCl3 in sodium acetate buffer (10 mM, pH 4). Radiolabeling yields >99% were confirmed by RP-HPLC. Each radiolabeled tracer was diluted with PBS (pH 7.4) to a concentration of 10 MBq/mL (370 µCi/mL). Each mouse was intravenously injected through the tail vein with ~ 1 MBq (100 µL) of the 111In complex and then sacrificed by inhalation of isoflurane followed by CO2 at 15 min, 1 h, 4 h, or 24 h after injection (n = 4 at each time point). Blood was withdrawn by cardiac puncture and tissues of interest including fat, uterus, ovaries, intestine, spleen, liver, pancreas, stomach, adrenal glands, kidney, lungs, heart, muscle, bone (tibia), brain, and tail were harvested, washed in PBS, dried and weighed. Activity of each sample was measured by a calibrated gamma 	 47 counter (Perkin Elmer, Wizard 2 2480) with decay correction. The activity uptake was expressed as a percentage of the injected dose per gram of tissue (%ID/g).  111In-neunpa/CHX-A”-DTPA-Trastuzumab SPECT/CT Imaging and Biodistribution Stud-ies Mice with SKOV-3 ovarian cancer xenografts were administered with ~37 MBq (~1 mCi) of 111In-neunpa-trastuzmab (1.03 MBq/µg [28.0 µCi/µg]) or 111In-CHX-A”-DTPA-Trastuzumab (0.77 MBq/µg [20.8 µCi/µg]) in ~30 µL of PBS (pH 7.4) via tail vein injec-tion. For each radioimmunoconjugate, mice were imaged (n=2) at 1, 3, or 5 day after injection. Image acquisition and reconstruction was performed using the U-SPECT-II/CT (MILabs, Utrecht, The Netherlands). Approximately 5 min prior to SPECT/CT im-age acquisition, mice were anesthetized via inhalation of 2% isoflurane/oxygen gas mixture and placed on the scanner bed. Anesthesia was maintained during imaging as well as body temperature via a heating pad. A 5 min baseline CT scan was obtained for localization with voltage setting at 60 kV and current at 615 µA followed by a static emission scan using an ultra-high-resolution multi-pinhole rat-mouse (1 mm pinhole size) collimator. Data were acquired in list mode, reconstructed using the U-SPECT II software and co-registered for alignment.  SPECT images were reconstructed using maximum-likelihood expectation maximization (3 iterations), pixel-based ordered sub-set expectation maximization (16 subsets) and a post-processing filter (Gaussian blur-ring) of 0.5 mm centered at photopeaks 171 keV and 245 keV with a 20% window width.  Imaging data sets were decayed corrected to injection time and converted to DICOM data for visualization in the Inveon Research Workplace (Siemens Medical Solutions USA, Inc.). As no calibration factor was used or attenuation and scatter correction were performed on the images; they were used only for qualitative comparison between the two tracers and are presented using a min-max min-max scale bar of counts corrected for decay. For biodistribution studies, mice were sacrificed by inhalation of isoflurane followed by CO2 (n = 4 at each time point), blood was withdrawn by cardiac puncture and tissues were collected and processed as described above. Tissues collected in-clude all those listed above in addition to tumor tissue.  Solution Thermodynamics Protonation constants and metal stability constants were calculated from po-tentiometric titrations using a Metrohm Titrando 809 equipped with a Ross combined electrode and a Metrohm Dosino 800. The titration apparatus consisted of a 20 mL and 25 ºC thermostated glass cell and an inlet-outlet tube for nitrogen gas (purified through a 10% NaOH solution) to exclude any CO2 prior and during the course of the titration. The electrode was daily calibrated in hydrogen ion concentrations using a standard HCl as described before13 in order to obtain the calibration parameters E0 and pKw. Solu-	48 tions were titrated with carbonate-free NaOH (0.157 M) that was standardized against freshly recrystallized potassium hydrogen phthalate. Protonation equilibria of the lig-and were studied by titrations of a solution containing H4neunpa 7.18 x 10-4 M at 25 ºC and 0.16 M NaCl ionic strength using a combined potentiometric-spectrophotometric procedure.16,17 Spectra were recorded in the 200–450 nm spectral range with a 0.2 cm path length fiber optic on a Varian Cary 60 UV/Vis Spectrophotometer. In the study of complex formation equilibria, the ligand-metal solutions were prepared by adding the atomic absorption (AA) standard metal ion solutions to a H4neunpa solution of known concentration in the 1:1 metal to ligand molar ratio. The exact amount of acid present in the lanthanum, bismuth and indium standards was determined by Gran’s method40 titrating equimolar solutions of either La(III), Bi(III) or In(III) and Na2H2-EDTA. Ligand and metal concentrations were in the range of 0.7-1.0 mM. Each titration consisted of 100-150 equilibrium points in the pH range 1.8-11.5, equilibration times for titrations were 2 min for pKa titrations and up to 15 min for metal complex titrations. At least two replicate titrations were performed for each individual system. Potentiometric data were processed using the Hyperquad2013 software41 while the obtained spectropho-tometric data were processed with the HypSpec.41 Proton dissociation constants cor-responding to hydrolysis of La(III), Bi(III) and In(III) aqueous ions and the indium-chloride stability constants included in the calculations were taken from Baes and Mesmer.42 The overall equilibrium (formation) constants log β are referred to the over-all equilibria: pM + qH + rL = MpHpLr (the charges are omitted), where p might also be 0 in the case of protonation equilibria and q may be negative. Stepwise equilibrium con-stants log K correspond to the difference in log units between the overall constants of sequentially protonated (or hydroxide) species. The pM values for metal complexes were calculated by using the Hyss software43, from the set of stability constants for each system at pH 7.4 with [L] = 1.0x10-5 M and [M] = 1.0x10-6 M.  6 References  1. Holland, J. P., Williamson, M. 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(1994) Evaluation of the 	 51 serum stability and in vivo biodistribution of CHX-DTPA and other ligands for yttrium labeling of monoclonal antibodies, Journal of nuclear medicine : official publication, Society of Nuclear Medicine 35, 882-889. 33. Ando, A., Ando, I., Hiraki, T., and Hisada, K. (1989) Relation between the location of elements in the periodic table and various organ-uptake rates, Nucl. Med. Biol. 16, 57-80. 34. Vosjan, M. J., Perk, L. R., Visser, G. W., Budde, M., Jurek, P., Kiefer, G. E., and van Dongen, G. A. (2010) Conjugation and radiolabeling of monoclonal antibodies with zirconium-89 for PET imaging using the bifunctional chelate p-isothiocyanatobenzyl-desferrioxamine, Nat. Protoc. 5, 739-743. 35. Tabrizi, M., Bornstein, G. G., and Suria, H. (2010) Biodistribution mechanisms of therapeutic monoclonal antibodies in health and disease, The AAPS journal 12, 33-43. 36. Meares, C. F., McCall, M. J., Reardan, D. T., Goodwin, D. A., Diamanti, C. I., and McTigue, M. (1984) Conjugation of antibodies with bifunctional chelating agents: isothiocyanate and bromoacetamide reagents, methods of analysis, and subsequent addition of metal ions, Anal. Biochem. 142, 68-78. 37. Deri, M. A., Ponnala, S., Kozlowski, P., Burton-Pye, B. P., Cicek, H. T., Hu, C., Lewis, J. S., and Francesconi, L. C. (2015) p-SCN-Bn-HOPO: A Superior Bifunctional Chelator for 89Zr ImmunoPET, Bioconj. Chem. 26, 2579-2591. 38. Lindmo, T., Boven, E., Cuttitta, F., Fedorko, J., and Bunn, P. A., Jr. (1984) Determination of the immunoreactive fraction of radiolabeled monoclonal antibodies by linear extrapolation to binding at infinite antigen excess, J. Immunol. Methods 72, 77-89. 39. Rousseau, J., Zhang, Z., Dias, G. M., Zhang, C., Colpo, N., Bénard, F., and Lin, K.-S. (2017) Design, synthesis and evaluation of novel bifunctional tetrahydroxamate chelators for PET imaging of 89Zr-labeled antibodies, Bioorg. Med. Chem. Let. 27, 708-712. 40. Gran, G. (1952) Determination of the equivalence point in potentiometric titrations. Part II, Analyst 77, 661-671. 41. Gans, P., Sabatini, A., and Vacca, A. (1996) Investigation of equilibria in solution. Determination of equilibrium constants with the HYPERQUAD suite of programs, Talanta 43, 1739-1753. 42. Baes, C. F. J., and Mesmer, R. E. (1976) The Hydrolysis of Cations, Wiley, New York. 43. Alderighi, L., Gans, P., Ienco, A., Peters, D., Sabatini, A., and Vacca, A. (1999) Hyperquad simulation and speciation (HySS): a utility program for the investigation of equilibria involving soluble and partially soluble species, Coord. Chem. Rev. 184, 311-318.      	52     53   Supporting Information Chapter A1 p-NO2-Bn-H4neunpa and H4neunpa-Trastuzumab:  Bifunctional Chelator for Radiometalpharmaceuticals and 111In Immuno-SPECT Imaging      Sarah Spreckelmeyer,a,b Caterina F. Ramogida,c Julie Rousseau,d Karen Arane,c Ivica Bratanovic,c Nadine Colpo,d Una Jermilova,d Gemma M. Dias,d Iulia Dude,d Maria de Guadalupe Jaraquemada-Peláez,a François Bénard,d Paul Schaffer,d Chris Orviga a Medicinal Inorganic Chemistry Group, Department of Chemistry, University of British Columbia, 2036 Main Mall, Vancouver, British Columbia, V6T 1Z1, Canada b Dept. Pharmacokinetics, Toxicology and Targeting, Research Institute of Pharmacy, University of Groningen, Antonius Deusinglaan 1, Groningen 9713 AV, The Netherlands c Life Sciences Division, TRIUMF, 4004 Wesbrook Mall, Vancouver, British Columbia, V6T 2A3, Canada d BC Cancer Agency, 675 West 10th Avenue, Vancouver, British Columbia, V5Z 1L3, Canada      54 	Figure S1.  1H NMR spectrum of Bi-p-NO2-Bn-neunpa complex ([Bi(6)]-, M2) (400 MHz, DMSO-d6, 25 °C).    Figure  S2.  HSQC NMR spectra of p-NO2-Bn-H4neunpa (6) (top) (101 MHz, 400 MHz, MeOD, 25 °C) and La-(p-NO2-Bn-neunpa) complex (bottom) (101 MHz, 400 MHz, DMSO-d6, 25 °C)  2.53.03.54.04.55.05.56.06.57.07.58.0f1	(ppm)2.5	DMSO-d6  55   Figure S3. a) UV spectroscopic titration of 7.18x10-4 M H4neunpa in 0.16 M NaCl between pH 2.5 and 5.39 b) UV spectroscopic titration of 7.18x10-4 M H4neunpa in 0.16 M NaCl between pH 5.39 and 8.33 c) UV spectrophotometric titration of 7.18x10-4 M H4neunpa in 0.16 M NaCl between pH 8.33 and 11.32.     Figure S4. a) Speciation plots for the H4neunpa ligand calculated with the protonation constants in Table 1, [H4neunpa] 2 4 6 8 10 120255075100L5-HL4-H2L3-H3L2-H4L-H5LH6L+% Formation relative to H5decapapHH7L2+b)2 4 6 8 10 120255075100L4-HL3-H2L2-H3L-H4LH5L+H6L2+% Formation relative to H4neunpapHa)250 300 350 4000.00.51.01.52.0pH 8.23AbsorbanceWavelength (nm)pH 2.42 λ = 291 nm250 300 350 4000.00.51.01.52.0pH 10.54AbsorbanceWavelength (nm)pH 8.48λ = 271 nma) b)  250 300 350 4000.00.51.01.52.0pH 5.39AbsorbanceWavelength (nm)λ = 296 nmpH 2.5250 300 350 4000.00.51.01.52.0pH 5.39AbsorbanceWavelength (nm)λ = 293 nmpH 8.33250 300 350 4000.00.51.01.52.0pH 11.32AbsorbanceWavelength (nm)λ = 284 nmpH 8.33a) b) c)   56 = 1.00 x 10-3 M, 25 °C and I = 0.16 M (NaCl). The dash lines indicate the fourth and fifth protonation processes. b) Speciation plots for the H5decapa ligand calculated with the protonation constants in Table 1, [H5decapa] = 1.00 x 10-3 M, 25 °C and I = 0.16 M (NaCl). The solid lines indicate the fourth and fifth protonation processes for comparison with those of H4neunpa ligand.  Figure S5. a) UV spectroscopic titration of 6.52x10-4 M H4neunpa and La3+ 1:1 metal to ligand molar ratio in 0.16 M NaCl between pH 2.42 and 8.23 b) UV spectroscopic titration of 6.52x10-4 M H4neunpa and La3+ 1:1 metal to ligand molar ratio in 0.16 M NaCl between pH 8.48 and 10.54.    Figure S6. Speciation diagram for the La3+-H4neunpa system from potentiometric titrations. [La3+] = [H4neunpa] = 6.52 x 10-4 M, 25 °C and I = 0.16 M (NaCl).   2 4 6 8 100255075100[La(L)(OH)]2-[La(L)]-La(HL)% Formation relative to La3+pHLa3+[La(H2L)]+2 4 6 8 10 120255075100[Bi(OH)4]-Bi(OH)3[Bi(L)(OH)]2-[Bi(L)]-Bi(HL)% Formation relative to Bi3+pH[Bi(H2L)]+  57 Figure S7. Speciation diagram for the Bi3+-H4neunpa system from potentiometric titrations. [Bi3+] = [H4neunpa] = 6.52 x 10-4 M, 25 °C and I = 0.16 M (NaCl).  Figure S8. Speciation diagram for the In3+-H4neunpa system from potentiometric titrations. [In3+] = [H4neunpa] = 6.62 x 10-4 M, 25 °C and I = 0.16 M (NaCl). The dash line indicates the optimal pH for the radiolabeling of H4neunpa-trastuzumab.   Figure S9. RP-HPLC radiochromatograms of [111In(p-NO2-Bn-neunpa)]- labeled at ambient temperature, 10 minutes reaction time in sodium acetate buffer (10 mM, pH 4) and decreasing ligand concentration.  2 4 6 8 100255075100[In(L)(OH)]2-[In(L)]-In(LH)% Formation relative to In3+pH[In(LH2)]+  58 Figure S10. RP-HPLC radiochromatograms of [111In(p-NH2-Bn-CHX-A”-DTPA)]2- labeled at ambient temperature, 10 minutes reaction time in sodium acetate buffer (10 mM, pH 4) and 0.1 (top) or 100 µM (bottom) final ligand concentration.   Figure S11. iTLC-SG radiotraces of 111In-neunpa-trastuzumab labeled at ambient temperature for 15 min, 200 µg of bioconjugate in ammonium acetate buffer (0.15 M) at varying pH values.    Table S1. Biodistribution data of 111In-CHX-A”-DTPA-/neunpa-trastuzumab performed over 5 days in mice bearing SKOV-3 ovarian cancer xenografts (n = 4 per time point) showing %ID/g values %ID/g 111In-CHX-A"-DTPA-trastuzumab 111In-neunpa-trastuzumab Organ 1 d 3 d 5 d 1 d 3 d 5 d Blood 17.0 ± 1.5 10.9 ± 1.8 4.2 ± 1.2 16.2 ± 0.6 6.3 ± 0.6 3.2 ± 0.6 Tumour 41.8 ± 17.9 51.0 ± 7.6 59.1 ± 7.7 27.5 ± 6.2 21.0 ± 10.0 16.0 ± 2.2 Fat 0.6 ± 0.1 0.6 ± 0.3 0.5 ± 0.1 0.9 ± 0.1 0.7 ± 0.2 0.3 ± 0.1 Uterus 15.4 ± 3.2 13.9 ± 1.9 10.7 ± 1.5 12.9 ± 1.2 10.1 ± 1.4 6.6 ± 1.1 Ovaries 10.0 ± 0.9 10.7 ± 1.2 7.5 ± 2.6 10.0 ± 1.2 5.2 ± 1.4 6.7 ± 1.2 Intestine 3.7 ± 0.5 5.5 ± 0.1 4.4 ± 0.3 3.8 ± 0.2 3.6 ± 0.3 2.3 ± 0.2 Spleen 25.6 ± 1.1 55.8 ± 3.8 49.7 ± 6.8 23.5 ± 1.8 27.0 ± 6.8 22 ± 6.6 Liver 8.9 ± 1.0 6.9 ± 0.8 5.2 ± 0.3 6.8 ± 3.0 4.4 ± 0.2 3.5 ± 0.5 Pancreas 1.5 ± 0.2 1.5 ± 0.2 1.0 ± 0.2 1.7 ± 0.1 1.2 ± 0.1 0.8 ± 0.1 Stomach 12.8 ± 19.7 2.7 ± 0.3 2.0 ± 0.3 3.2 ± 0.3 1.9 ± 0.1 1.2 ± 0.2 Adrenal glands 3.8 ± 1.1 3.0 ± 1.0 1.5 ± 0.5 3.7 ± 0.5 1.8 ± 0.3 1.1 ± 0.3 Kidney 5.0 ± 0.3 3.5 ± 0.3 2.0 ± 0.2 6.6 ± 1.0 5.0 ± 0.2 4.3 ± 0.3 Lungs 7.3 ± 0.6 5.4 ± 1.2 3.3 ± 0.7 7.6 ± 0.7 3.7 ± 0.3 2.4 ± 0.3   59 Heart 4.7 ± 0.4 2.9 ± 0.6 1.4 ± 0.4 4.4 ± 0.6 1.7 ± 0.2 1.1 ± 0.1 Muscle 0.9 ± 0.1 1.0 ± 0.2 0.4 ± 0.1 1.4 ± 0.2 0.6 ± 0.2 0.4 ± 0.1 Bone 6.2 ± 4.3 4.2 ± 0.4 2.9 ± 0.4 4.6 ± 0.3 2.9 ± 0.7 2.4 ± 0.2 Brain 0.3 ± 0.04 0.3 ± 0.03 0.1 ± 0.04 0.3 ± 0.04 0.2 ± 0.04 0.1 ± 0.01 Tail 3.4 ± 0.3 2.5 ± 0.4 1.6 ± 0.1 3.2 ± 0.2 2.0 ± 0.2 1.3 ± 0.1      Figure  S12.  COSY NMR spectra of In-(p-NO2-Bn-neunpa) complex (101 MHz, 400 MHz, DMSO-d6, 25 °C)   Figure S13.  Crude reaction mixture iTLC-SG radiotraces of 111In-neunpa/CHX-A”-DTPA-trastuzumab prepared for in vivo studies labeled at ambient temperature for 30 min, 650 µg of bioconjugate in ammonium acetate buffer (0.15 M, pH 6), or 111In3+ control (no chelate-antibody added); iTLC-SG developed in 50 mM EDTA (pH 5).     2.53.03.54.04.55.05.56.06.57.07.58.08.5f2	(ppm)2.53.03.54.04.55.05.56.06.57.07.58.08.5f1	(ppm)  60     		61	   Chapter A2 H4neunpa: A Bifunctional Acyclic Chelator with many Faces    Sarah Spreckelmeyer,a,b Caterina Ramogida,c Hsiou-Ting Kuo,d Ben Woods, e Valery Radchenko,c Maria Guadalupe Jaraquemada Peláez,a Francois Bénard,d Angela Casinie and Chris Orviga  a Medicinal Inorganic Chemistry Group, Department of Chemistry, University of British Columbia, 2036 Main Mall, Vancouver, British Columbia, V6T 1Z1, Canada b Department of Pharmacokinetics, Toxicology and Targeting, Groningen Research Institute of Pharmacy, University of Groningen, Antonius Deusinglaan 1, Groningen 9713 AV, The Netherlands c Life Sciences Division, TRIUMF, 4004 Wesbrook Mall, Vancouver, British Columbia, V6T 2A3, Canada d BC Cancer Agency, 675 West 10th Avenue, Vancouver, British Columbia, V5Z 1L3, Canada e School of Chemistry, Cardiff University, Park Place, CF103AT Cardiff, United Kingdom      		62	1 Abstract Bifunctional chelators are useful tools in nuclear medicine and they also find great application in the relevant field of personalized medicine. We evaluated the nonadentate chelator H4neunpa as a bifunctional chelator for therapeutic as well as diagnostic application. Several conjugates were synthesized. First, H4neunpa was bifunctionalized with a small biomolecule Glu-ureido-Lys, as biovector that targets the prostate-specific membrane antigen (PSMA) for prostate cancer and can be further radiolabelled with 111In for single photon emission computed tomography (SPECT) imaging. H4neunpa was also bifunctionalized with a supramolecular metallacage precursor for cisplatin encapsulation. This probe was further investigated for La3+ complexation in order to use it in the future for fluorescence imaging and in vitro/in vivo tracking of the supramolecular metallacage. Besides modifying the biovector (Glu-ureido-Lys or metallacage), we also achieved promising radiolabeling results with different radiomdoctoretals for therapy (225Ac, 213Bi and natSb). 225Ac and 213Bi are interesting alpha emitters for targeted alpha therapy (TAT). 119Sb is a promising Auger electron emitter for targeted radiotherapy.                    		63	2 Introduction Cancer is one of the leading causes of morbidity and mortality worldwide, with approximately 14 million new cases in 2012, 8.8 million deaths in 2015, and an expected rise by about 70% over the next two decades.1 Of note, in the last decades, the treatment and diagnosis of cancer improved tremendously, but we are also still facing serious obstacles. As individuals are unique, so is each cancer type and differences can be observed on the macroscopic (e. g. tissue invasions, growth rate) as well as microscopic (e. g. protein expression) level. To cope with this, a persodoctornalized treatment that is targeted to the needs of a patient based on his/her own genetic, biomarker, phenotypic, or psychosocial characteristics, is absolutely necessary. Among numerous methods for personalized medicine, the use of a bifunctional chelator (BFC, Figure 1) in nuclear medicine is a promising strategy and various types are clinically used, like 131I-tositumomab and 90Y-ibritumomab that are bearing anti-CD20 antibodies for the treatment of B cell lymphoma and leukemia.2 BFCs consist of a biovector, like small molecules or antibodies, a linker, and a chelator that stably binds the radiometal of choice. The antibody can have either a targeting or therapeutic value and the radiometal of interest can be either for imaging or diagnosis.  Figure		1.		Illustration	of	a	bifunctional	chelator	(BFC).	Before the treatment with these BFCs, the expression level of the CD20 antigen needs to be assessed. If the expression level is sufficient, a therapy with anti-CD20 antibodies is promising. Non-responders are directly sorted out. By this, side-effects are limited and costs are kept as low as possible.  Overall, the aim of this chapter is to show that the bifunctional chelator H4neunpa can easily be modified for multiple purposes in the field of medicinal inorganic chemistry. This chapter is divided into three sub-chapters, each of them consisting of a short introduction and results and discussion part. The experimental part and the summary are combined. Specifically, the first sub-chapter deals with the prostate-specific membrane antigen (PSMA), which is an attractive target for targeted radiotherapy. A bifunctional chelator, H4neunpa, was linked to a PSMA-specific peptide and radiolabeled with 111In. The second sub-chapter describes the linkage of H4neunpa to a metallacage for therapeutic purposes and in the third sub-chapter, radiolabeling of H4neunpa with different radiometals for imaging or therapeutic purposes is investigated. 2.1 Subchapter 1  An attractive target for prostate cancer radiopharmaceuticals is the prostate-specific membrane antigen (PSMA). PSMA is a well-characterized biomarker for imaging prostate cancer. It is a type II membrane-bound, glutamate-preferring carboxypeptidase and is expressed on prostate tissue, with strong overexpression in prostate cancer. The first antibody targeting PSMA that was published was mAb 7E11 which binds to the receptor intracellularly.3 In the late 1990s, ProstaScint (capromab pendetide, EUSA Pharma, Figure 2) was approved by the FDA for imaging prostate cancer. ProstaScint consists of the chelator GYK-DTPA-HCl that chelates the gamma emitter 111In (thus suitable for SPECT imaging) and is linked to the murine monoclonal antibody 7E11 (capromab).4 Beside antibodies (eg. 7E11) that target PSMA, also aptamers and PSMA inhibitors of low molecular weight have gained interest as targeting devices for prostate cancer. PSMA possesses an enzymatic site in its extracellular domain that cleaves endogenous substrates such as N-acetylaspartylglutamic acid (NAAG) and poly-gamma-glutamyl folic acid. The enzymatic site contains two zinc ions, and is composed of two pockets: the glutamate sensing pocket and the non-pharmacophore pocket that contains an arginine rich region.  		64	Small molecules have been designed to inhibit PSMA, containing a zinc-binding moiety, a glutamate moiety that can reside in the glutamate sensing pocket and a lipophilic moiety that can reside in the non-pharmacophore pocket.5 Beside phosphonate-, phosphate-, and phosphoramidates and thiols, ureas play an important role in small molecule design of PSMA inhibitors.5 Recently, lysine-glutamate-urea based small molecules gained a lot of interest in PET imaging (Figure 1), but no SPECT tracer is available up to now.  Herein, we present the synthesis of a Glu-ureido-Lys based small molecule linked to p-SCN-Bn-H4neunpa, which is called H4neunpa-PSMA-L, as it is aimed to bind to PSMA. Additionally, we performed preliminary radiolabeling experiments with 111In as well as determined  stability in human serum and determination of the 111In-neunpa-PSMA-L chelate for SPECT imaging of prostate cancer. The aim of this study was to have a proof-of-principle that also small molecules can easily be linked to H4neunpa without loosing its 111In-chelating properties.  Figure		2.		Compounds	discussed	in	this	work.	2.1.1 Results and Discussion 2.1.1.1 Synthesis p-SCN-Bn-H4neunpa was synthesized using a protocol already published in chapter A1.6 The Glu-ureido-Lys moiety (Scheme S 1) was linked via a reaction of the isothiocyanate of p-SCN-Bn-H4neunpa with the amine functional group of Glu-ureido-Lys to yield H4neunpa-PSMA-L (Figure 3). Characterization of the product was achieved via HR ESI-MS (see Experimental).     Figure  3. Synthesis scheme of H4neunpa-PSMA-L. 2.1.1.2 Radiolabeling with 111In Radiolabeling experiments with 111InCl3 (200 µCi) were performed in a concentration range of 10-8 M to 10-4 M H4neunpa-PSMA-L and a maximum of 82 % radiochemical yield (RCY) was achieved at 10-5 M (Figure 4). Unfortunately, increasing the concentration of H4neunpa-PSMA-L did not lead to an increase in RCY. The reason for missing about 18 % RCY for quantitative radiolabeling might be that the three carboxylic acids of Glu-ureido-Lys interfere with 111In chelation of the H4neunpa moiety. NHNHOHHOO OONH2NNNNCSNNOHOHOOOHOOOHp-Bn-SCN-H4neunpa Glu-ureido-LysN N N NHHOO OOOHOHOOOHProstaScintOOHNHO HNONHHOcapromabOOHNNNNCSNNOHOHOOOHOOOHNHNHOHHOOOOHNNNNHNNNOHOHOOOHOOOHSOHONH NHOHHOO OONH2OHODMF, DIEAH4neunpa-p-Bn-NCSH4neunpa-PSMA-L		65	 Figure		4.		RCY	of	H4neunpa-PSMA	with	111In.	2.1.1.3 Stability of H4neunpa-PSMA-L in human serum The stability of 111In-neunpa-PSMA-L in human serum was assessed at time points 1, 48 and 120h (Table 1). After 1h incubation, the RCY decreased to 67 % (% of t=0 was taken as 100% reference). The RCY decreased further to 60 % after 120h incubation in human serum albumin. 111In-neunpa-PSMA-L is clearly not stable under these conditions. We assume, that the carboxylic acids of Glu-ureido-Lys might interact with the chelation of 111In, since carboxylic acids are in general good chelators for 111In and transchelation with albumin may take place. The resulting 111In-neunpa-PSMA-L complex might be less stable than 111In-neunpa (see chapter A1).7 In addition, the distance between the carboxylic acids of the lysine-glutamate-urea and the neunpa-cavity is very small, favoring the removal of the 111In from the chelator by the carboxylic groups. Table		1.	Stability	of	111In-neunpa-PSMA-L	(10-5M)	in	human	serum,	expressed	as	%	of	the	%RCY	at	t=0.	Time [h] % RCY 0 100 1 66.8 ± 2.3 48 65.2 ± 0.1 120 58.9 ± 0.7  2.2 Subchapter 2 Instead of linking a small molecule like Glu-ureido-Lys to p-SCN-Bn-H4neunpa, a metallacage can be added for imaging purposes (111In, SPECT imaging) or therapeutic purposes. Self-assembled metallacages that incorporate a chemotherapeutic drug like cisplatin may represent an attractive drug-delivery system. Exo-functionalized Pd2L4 cages, formed by self-assembly of the four ligands in the presence of Pd ions, are therefore highly promising metallacages (Figure 5), that are proven to encapsulate cisplatin.8 Not much is known about these fairly new compounds. In order to get more information about the mechanism of action and cellular uptake mechanisms of the metallacages, they can be linked to a fluorescent moiety such as lanthanide complexes. Using this approach, their uptake can be visualized by fluorescence microscopy in vitro, provided that the uptake characteristics of the metallacages will not be changed upon bifunctional chelator (BFC) linkage. The use of a (BFC) like p-SCN-Bn-H4neunpa is very promising for this purpose. p-SCN-Bn-H4neunpa has shown to be a good chelator for various metal ions, like La3+, In3+ and Bi3+.7  10-9 10-8 10-7 10-6 10-5 10-4 10-3020406080100ligand [mol/L]% RCY		66	Lanthanide cations are known for their unique photonic and magnetic properties associated with their f0-f14 electron configuration. They have wide practical applications in many fields including catalysis, additives in glass, photonic applications, as well as luminescent stains for biomedical analysis, medical diagnosis and cellular optical imaging.9 La3+ ([Kr]4d105s25p6, 1.03 Å) and Eu3+ (([Xe]4f6, 0.95 Å) are metal ions that have fluorescent properties. Here, we report on the synthesis of a self-assembled metallacage functionalized with La3+-H4neunpa  to achieve fluorescent properties for cellular imaging.   Figure		5.		Synthetic	approach	to	achieve	metallacage-NH2	by	self-assembly.		2.2.1 Results and Discussion 2.2.1.1 Metallacage exo-functionalization  The synthesis of the BFC p-SCN-Bn-H4neunpa has already been published and described in chapter A1.7 The coupling of the metallacage-NH2 to this BFC can be achieved in two ways: i) The first possibility starts with the metallacage precursor metallacage-ligand-NH2 (Figure 5), which bears an amine functional group. This precursor can be coupled to the NCS functional group of p-SCN-Bn-H4neunpa via a thiourea linkage. Thereafter, the metal ion complexation can be performed and in a next step, the metallacage formation will be achieved as shown in Figure 6.8 ii) The second possibility starts with 4 eq. of p-SCN-Bn-H4neunpa that will be coupled directly to the already self-assembled metallacage-NH2 (Figure 5). After that, the metal ion complexation will be performed. In both cases, after successful metal complexation and cage formation, cisplatin can be incorporated. Here, we describe the results and discussion of the first approach: The metallacage-ligand-NH2 was coupled via the reaction between its amine and the isothiocyanate group from p-SCN-Bn-H4neunpa to form compound 1 (Figure 6). The product was purified by HPLC. In Figure 7, the 1H NMR spectrum of the H4neunpa-cageligand (compound 1) and of p-SCN-Bn-H4neunpa can be seen.  All proton peaks can be properly assigned (Figure 8) with the help of 2D-HSQC and 2D-COSY NMR spectra (Figure S2 – Figure S5). The next step was the metal ion complexation. Since Eu3+ complexation was not successful (no results in ESI-MS, no fluorescence), we performed a La3+ complexation. HR ESI-MS and 1H NMR confirmed the successful complexation reaction (Figure 9). In addition, fluorescence was observed (Figure 10); this will be discussed below in more detail. NNH2Nmetallacage-ligand-NH2NNH2NPdPdNNNH2NH2NNH2NNNmetallacage-NH2[Pd(NCCH3)4][BF4]20.5 eq.rt, 1h, DMSO		67	  Figure		6.		Synthesis	scheme	of	La-neunpa-metallacage	(compound	1,	compound	2	and	compound	3).	 Figure		7.		1H	NMR	spectra	of	(A)	p-Bn-SCN-H4neunpa	and	(B)	compound	1	(H4neunpa-cageligand	)	(400MHz,	MeOD,	25°C).	NNNNCSNNOHOHOOOHOOOHNNNH2 NNNHNNNOHOHOOOHOOOHNNHNSDMSOLa(NO3)3NNNHNNNOHOHOOOHOOOHNNHNSLa123NNNHNNN OHOHOOHOOOHOSNNNHPdPdNNHNHNNNHNNNNNNNHNNOHOHOOHO OOHOSNNNHNNNHOHOOOOHOO OHSNNNHNNNHOHOOOOHOOOHSLaLaLaLa1a[Pd(NCCH3)4[BF4]20.5 eq.rt, 1h, DMSO3.03.54.04.55.05.56.06.57.07.58.08.59.0f1	(ppm)3.31	CD3OD3.03.54.04.55.05.56.06.57.07.58.08.59.0f1	(ppm)3.3	CD3ODA:	p-Bn-SCN-H4neunpaB:	H4neunpa-cageligand		68	  Figure		8.		1H	NMR	peak	assignments	of	compound	1.	2.2.1.2 La3+ complexation reaction The La3+ complexation of compound 1 was performed following a standard protocol.7 HR ESI-MS and the 1H NMR spectrum of the product compared to its precursor show clear changes in the aromatic region as well as in the alkyl region (Figure 9). Peak assignments for the La complex are difficult due to the low concentration of the sample.   Figure		9.	1H	NMR	spectra	of	compound	1	(top,	400	MHz,	MeOD,	25°C)	and	compound	2	(bottom,	400	MHz,	DMSO-d6,	25°C).	NNNHNNNOHOHOOOHOOOHNNHNS11223 3445 6AABBCCDDEEFGGH HI IJJK K2.93.03.13.23.33.43.53.63.73.83.94.04.14.24.34.4f1	(ppm)3.31	CD3OD6.97.07.17.27.37.47.57.67.77.87.98.08.18.28.38.48.58.68.78.88.99.0f1	(ppm)A D B+KJEC	+	F I G H1352462.02.53.03.54.04.55.05.56.06.57.07.58.08.59.09.510.0f1	(ppm)2.322.50	DMSO-d62.662.722.883.884.546.647.217.487.547.778.028.608.789.949.983.043.253.31	CD3OD3.483.583.674.037.247.417.517.597.757.928.028.568.74		69	 Figure		10.		A	NMR	tube	showing	the	fluorescence	of	compound	2.	 2.2.1.3 Fluorescence spectroscopy The solutions of compound 2, metallacage-ligand-NH2 and La-neunpa-NO2 were prepared at a concentration of 10-6 M in DMSO and the fluorescence was measured at λex = 305 nm. In Figure 11, the emission spectra of the compound 2, metallacage-ligand-NH2 and La-neunpa-NO2 is shown. The compound 2 shows high intensity fluorescence at 430 nm (line 2). La-neunpa-NO2 itself shows a weak emission at 360 nm and 430 nm (line 1 and line 2). The addition of La3+ ions results in successive red shifts of the emission peaks to 380 nm (line 1a) and 430 nm (line 2) accompanied by the enhancement of emission intensity. The enhancement of emission intensity was previously described by F. Wang et al., who studied the effect of a set of lanthanides on the fluorescence intensity of carminic acid, and showed that La3+ results in the best enhancement of fluorescence when compared to Dy3+, Tb3+, Gd3+, Eu3+, Y3+ and Sm3+.10     Figure		11.	Emission	spectra	of	compound	2,	metallacage-ligand-NH2	and	La-neunpa-NO2.		 2.3 Subchapter 3 Instead of labelling H4neunpa with a metal to obtain a fluorescent molecule, the BFC can also be loaded with a radioactive metal for therapeutic or diagnostic purposes. Considering the periodic table, a large number of radiometals can be identified for either therapeutic or diagnostic purposes. Here, we discuss 225Ac and 213Bi as alpha emitters for targeted alpha therapy (TAT) and 119Sb as Auger-electron emitter for targeted radiotherapy (TRT). Alpha emitters and Auger electrons have a characteristic linear energy transfer (LET). Alpha emitters such as 225Ac (t1/2 = 10 d), 212Pb (t1/2 = 10.6 h), 213Bi (t1/2 = 45.6 min) are gaining popularity for labeling biomolecules in targeted α-therapy (TAT). An essential 400 500 60002004006008001000wavelength [nm]EmissionCompound 2metallacage-ligand-NH2La-neunpa-NO21 1a 2		70	characteristic of the radiometal for targeted radiotherapy is a short travelling distance in tissue and a radiation with high-energy transfer. 225Ac and 213Bi are attractive radiometals for therapeutic applications due to their high-energy alpha decays (see simplified decay scheme in Figure 12).  119Sb is another interesting isotope that emits Auger-electrons and could be used in targeted radiotherapy of small tumours, micrometastases and single cancer cells. 117Sb can be used in SPECT for visualization of the tumor.11   Figure		12.		Decay	chain	of	225	Ac.	 2.3.1 Results and Discussion 2.3.1.1 Radiolabeling with 225Ac The ISAC (isotope separator and accelerator) facility is a unique and powerful resource that offers the possibility of a superior production method for alpha-emitting isotopes such as 225Ac, 213Bi, and 212Pb. 225Ac and 225Ra were produced in the spallation of an uranium carbide (UCx) target with 480 MeV protons, following a separation of 225Ra/225Ac from other isotopes via a high resolution mass separator. Purification of 225Ac was achieved using solid phase extraction on branched-DGA resin.12 Radiolabeling of p-Bn-NO2-H4neunpa with 225Ac was compared to that of the macrocyclic DOTA. Incubation of 10-5 or 10-4 M of p-Bn-NO2-H4neunpa at ambient temperature for 30 min with 225Ac yielded an 88.8 % RCY and 98.00 % RCY respectively, as determined via iTLC. For DOTA, the same conditions were used and resulted in 0.85 % RCY and 0.96 % RCY, respectively. p-Bn-NO2-H4neunpa has clearly superior radiolabeling kinetics compared to the gold standard DOTA under these conditions. The stability of both 225Ac complexes was determined after 120 min incubation in NH4OAc (0.15 M, pH 5) buffer solution. The RCY of both p-Bn-NO2-H4neunpa and DOTA remained stable over the tested time period (Figure 13).  		71	 Figure		13.		%RCY	of	the	tested	chelators	225Ac-H4neunpa-p-Bn-NO2	and	225Ac-DOTA.	2.3.1.2 225Ac/213Bi iTLC chromatograms In Figure 14, three sets (1, 10 and 100 µg of chelator) of iTLC chromatograms are shown, representing the results of different times of incubation (0, 15, 30, 60 and 120 min) of neunpa with 225Ac3+. The following features are valid for all iTLC chromatograms shown here. The origin (0-30 mm) represents free radiometal (225Ac and daughters). This is due to the use of 10 mM NaOH/ 10% NaCl as the mobile phase, which causes the free metal ions to form hydroxide species that precipitate at the baseline. Between 30 mm and 120 mm, metal complexes can be detected. The black graph represents the chromatogram analyzed directly after finishing the development of the plate of different concentrations of neunpa-NO2 at different times of incubation with 225Ac3+. The blue chromatogram represents the radioactivity patterns on the plates 3 h later.              0 50 100050100Time [min]% RCY Actinium-225neunpa: 10-4 Mneunpa: 10-5 MDOTA 10-4 MDOTA: 10-5 M		72	                      In general, three scenarios of peak development can be anticipated between the black and blue graphs:13  A) If a peak on the plate is only the daughter of 225Ac, 213Bi, then it will decay within the 3h difference between measurements based on the 46 min half-life of 213Bi, and counts will decrease accordingly.  Figure		14.	iTLC chromatograms of neunpa labeled with 225Ac ( (black: 0, 15, 30, 60 and 120 min for 1, 10 or 100 µg chelator; blue: same iTLC plate read 3h later).			0	min	15	min	30	min	60	min	120	min	1	 µg						10	µg						100	µg							050100150050100150Distance eluted [mm]Counts050100150050100150Distance eluted [mm]Counts050100150050100150Distance eluted [mm]Counts050100150050100150Distance eluted [mm]Counts050100150050100150Distance eluted [mm]Counts050100150050100150Distance eluted [mm]Counts050100150050100150Distance eluted [mm]Counts050100150050100150Distance eluted [mm]Counts050100150050100150Distance eluted [mm]Counts050100150050100150Distance eluted [mm]Counts050100150050100150200Distance eluted [mm]Counts050100150050100150200Distance eluted [mm]Counts050100150050100150200Distance eluted [mm]Counts050100150050100150200Distance eluted [mm]Counts050100150050100150200Distance eluted [mm]Counts		73	B) If a peak on the plate is a mixture of 225Ac and 213Bi in equilibrium, the total number of counts will remain more or less constant over a period of time, since as 213Bi decays, more is constantly being produced by the 225Ac decay, hence 225Ac and 213Bi are in equilibrium.  C) If a peak contains primarily 225Ac, then as the 225Ac decays, it forms detectable 213Bi, and we will get an increase in counts over time as 225Ac decays to 213Bi. The counts will increase (so the plate reader will detect the gamma rays from both the 225Ac and the 213Bi resulting in an increase in counts). The counts will increase until the 225Ac reached equilibrium with 213Bi after that point the counts will stay constant as in 'B'. First, we compare the iTLC chromatograms for the samples taken at 0 min of incubation of neunpa with 225Ac at different concentrations of neunpa-NO2 (1, 10 and 100 µg) with each other. The chromatogram of the lowest concentration of neunpa-NO2 (1 µg) shows two black peaks, one at the origin and one at 100 mm distance eluted. The origin peak stays the same in counts during 3h and the 100mm peak decreases completely within 3h. A third blue peak appeared between 30-60 mm distance eluted. The peak at 0-30 mm can be assigned as free 225Ac and 213Bi being in equilibrium (scenario B), the 100 mm peak can be assigned as Bi-neunpa-NO2 complex (scenario A) and the 30-60mm peak as Ac-neunpa-NO2 peak (scenario C).  At 10 µg neunpa-NO2, one origin peak, one possible peak at 50 mm and one distinct peak at 100 mm can be observed. After a 3 h time period, the origin peak decreased, giving assumption that only free 213Bi was at the origin in its hydroxide form. The peak at 50 mm increased in counts and another peak at 80 mm showed up, giving assumption that two Ac-neunpa-NO2 complexes are formed. The 100 mm peak decreased, accounting for a Bi-neunpa-NO2 complex.  At 100 µg neunpa-NO2, three peaks can be seen, similar to the 10 µg chromatogram, one at the origin, one at 50 mm and one at 100 mm. After a 3h time period, the origin peak decreased, the 50 mm peak decreased slightly, making it difficult to assign as pure Ac or Bi complex. Another distinct 80 mm peak showed up, being an Ac-neunpa-NO2 complex and the 100 mm peak decreased completely, accounting for a Bi-neunpa-NO2 complex. Overall, comparing the iTLC chromatograms of different concentrations of neunpa-NO2 at 0 min incubation with each other shows that a Bi-neunpa-NO2 complex is already formed at low concentrations, indicating a high affinity to 213Bi. At higher concentrations, two Ac-neunpa-NO2 complexes can be seen as well as the Bi-neunpa-NO2 complex. Secondly, each row gets analyzed in more detail, showing a fixed neunpa-NO2 concentration after different times of incubation. At 1 µg neunpa-NO2, the 100 mm peak, likely a Bi-neunpa-complex, decreased over the time of incubation. This might be due to the short 213Bi half-life. The peak between 20-60 mm gets into equilibrium after 30 min incubation, since no difference between the black and blue graph can be observed. At 10 µg neunpa-NO2, the counts of each peak decrease over time, due to the half-life of the metals and a new equilibrium between 225Ac and 213Bi. At 100 µg neunpa-NO2, the second Ac-neunpa-NO2 complex at 80 mm is very distinct and is persistent over 120 min, giving assumption to a pure Ac-neunpa-NO2 complex. The 50 mm peak does not change drastically and the black and blue graphs are overlapping, giving assumption to a mixture of Ac-225 and Bi-213 neunpa-NO2 complexes. 2.3.1.3 Sb-complexation  Figure		15.		Antimony	complexes	discussed	in	this	work.	N OOHNNOONHOONOOH2NSbHONHOOHN OSbHNOONHONNNNO2NNOHOHOOOOOOSbDFO-Sb (SAHA)2-Sb neunpa-p-Bn-NO2-Sb		74	In addition to complexation with 225Ac (Figure 14), cold complexation with antimony (Sb) was performed using solid antimony(V)oxide. Desferioxamine (DFO) and suberanilohydroxamic acid (SAHA) consist of hydroxamic acids moieties which are hard donors, meeting theoretically the requirements for complexation of the hard acid Sb. DFO, SAHA and H4neunpa successfully chelated antimony (Figure 15), as proven by HR ESI-MS. DFO and H4neunpa formed a 1:1 complex and SAHA a 2:1 complex. Noteworthy, the HR-ESI-MS showed a single positive charge on each Sb-complex. This suggests that antimony was reduced to Sb(III) upon chelation. The 1H NMR spectrum of Sb-neunpa-p-Bn-NO2 is shown in Figure 16.  Figure		16.	1H	NMR	spectra	of	p-Bn-NO2-neunpa	(top)	and	Sb-neunpa-p-Bn-NO2	(bottom)	(400	MHz,	MeOD,	25°C).	 3 Summary H4neunpa is a very versatile chelator. We showed that H4neunpa could easily be bifunctionalized with Glu-ureido-Lys (Scheme S1) to target the PSMA receptor, which is highly expressed in prostate cancer. Unfortunately, the three carboxylic acids of Glu-ureido-Lys seem to interfere with the radiolabeling properties of H4neunpa, resulting in a decreased RCY compared previously radiolabeled H4neunpa-p-Bn-NO2 and its antibody derivative H4neunpa--trastuzumab.7 We hypothesize, that the distance between Glu-ureido-Lys and H4neunpa is too short and an extension of the distance between the functionalization part and the chelator will improve radiolabeling efficiencies. Different lengths of linker are currently being studied at the BC Cancer Agency in Vancouver with the aim of increasing radiolabeling efficiencies with 111In3+.  In addition, bifunctionalization can be performed with a therapeutic metallacage, which is able to incorporate cisplatin for the treatment of specific cancer types. We successfully synthesized compound 2, the precursor of the metallacage, as proven via various 1D and 2D NMR spectroscopies as well as HR ESI-MS. The formation of the metallacage is still ongoing in a collaboration project between UBC and the University of Cardiff. After the successful H4neunpa-metallacage synthesis, extensive studies are needed to confirm the cell uptake of the new compound compared to the metallacage alone as well as the evaluation of the cytotoxic profile. 3.03.54.04.55.05.56.06.57.07.58.08.59.0f1	(ppm)1.482.463.613.433.954.272.002.145.863.253.31	CD3OD3.653.753.904.044.074.577.547.768.131.62.64.54.23.63.72.12.06.53.31	CD3OD3.653.823.954.124.667.577.597.837.848.158.19		75	Furthermore, H4neunpa-p-Bn-NO2 shows stable 225Ac labeling efficiencies >95 % over 120 minutes incubation at room temperature at a chelator concentration of 10-4 M. DOTA showed only 1 % radiolabeling yield under the same conditions. Conversely, H4neunpa-p-Bn-NO2 forms a 213Bi complex at 10-6 M and additionally, at higher concentrations, two 225Ac complexes. These results are extremely promising, since H4neunpa is able to bind to 225Ac as well as its daughter nuclide 213Bi, which makes it a potent chelator. In order to proof complex stability, the stability should be assessed in serum. These experiments are currently underway at TRIUMF. To conclude, H4neunpa can be labelled with diverse radiometals for therapy (225Ac, 213Bi) and imaging (111In) and it might thus find application as diagnostic or therapeutic bifunctional chelator. Current studies are ongoing to understand the mechanism of action of 111In-neunpa as well as human serum stability experiments for 225Ac experiments.  4 Experimental Materials and Methods All solvents and reagents were from commercial sources (Sigma Aldrich, TCI) and were used as received unless otherwise noted. 1H and 13C NMR spectra were recorded at room temperature on a Bruker AV400 instrument; the NMR spectra are expressed on the δ (ppm) scale and are referenced to the residual solvent signal of the deuterated solvent. All spectra were recorded with sweep widths of 0-14 ppm or -20-220 ppm for 1H and 13C NMR respectively. Assignments of the peaks in the NMR spectra are approximate. Mass spectrometry was performed on a Waters ZQ spectrometer equipped with an electrospray source at the Department of Chemistry, University of British Columbia. The HPLC system used for purification of ligands and precursors consisted of a Waters 600 controller equipped with a Waters 2487 dual λ absorbance detector connected to a Phenomenex synergi hydro-RP 80Å 250mm x 4.60 mm semipreparative column. Fluorescence spectra were recorded on Varian Cary Eclipse fluorescence spectrophotometer. Synthesis of compounds Synthesis of H4neunpa-PSMA-L 10 mg (0.015 mmol) of H4neunpa-p-Bn-NCS7 and 10 mg (0.031 mmol, 2 eq.) of Glu-ureido-Lys (Scheme S1) were dissolved in DMF (2 mL). 26.1 µL (0.150 mmol, 10 eq.) of N,N-diisopropylethylamine (DIEA) was then added and the solution was allowed to stir at room temperature for 2 days. Water (1 ml) was then added to the mixture and the mixture was lyophilized. The crude product was purified by RP-HPLC using a semi-preparative column eluted with 18 % acetonitrile with 0.1 % TFA at a flow rate of 4.5 mL/min. The retention time was 13.6 min, and the yield of the product was 13.7 % (1.9 mg). HR ESI-MS: calcd. [M+H]+ for H4neunpa-PSMA-L C43H55N9O15S 970.3617; found [M+H]+ 970.4766  Labelling of H4neunpa-PSMA-L with non-radioactive Indium (cold standard) H4neunpa-PSMA-L (1.5 mg, 1.5 μmol) was suspended in 200 µL water and 200 µL 0.1 M HCl as well as In(NO3)3 (2.3 mg, 7.5 μmol) were then added. The pH of the reaction mixture was adjusted to 4-5 by using 1 M NaOH to achieve precipitation of the product. After 1 hour of stirring the reaction mixture, the reaction mixture was centrifuged to obtain the product as a white precipitate. After dissolving the crude product in H2O/acetonitrile (2 mL, 1:1) with 0.1 % TFA, the product was purified by RP-HPLC using a semi-preparative column eluted with 18 % acetonitrile with 0.1 % TFA at a flow rate of 4.5 mL/min. The retention time was 11.8 min, and the yield of the product was 43.7 % (0.7 mg). HR ESI-MS: calcd. [M+H]+ for In-H4-neunpa-PSMA C43H52InN9O15S 1081.2342; found [M+H]+ 1081.2025 Radiolabeling of neunpa-PSMA-L with 111In The ligand p-NO2-Bn-H4neunpa-PSMA-L was made up as a stock solution (1 mg/mL, ~10-3 M) in deionized water. From this stock solution, serial dilutions were prepared to final ligand concentrations of 10-4 M – 10-8 M.  A 100 µL aliquot of each ligand stock (10-4 to 10-8 M) or water (blank control) was added to screw-cap mass spectrometry vials and diluted with sodium acetate buffer (pH 4, 10 mM, 880 µL). An aliquot of diluted 111In stock (20 µL, ~200 µCi) was added to each vial and 		76	allowed to radiolabel at ambient temperature for 10 min, then it was analyzed by RP-HPLC to confirm radiolabeling and calculate radiochemical yields. To study the stability of the radiolabelled compound in human serum, undiluted 111InCl3 stock (~20 µL, 5 mCi) was added to the reaction vial containing 10-4 M ligand in sodium acetate buffer. Areas under the peaks observed in the HPLC radio-trace were integrated to determine radiolabeling yields. Elution conditions used for RP-HPLC analysis were gradient: A: 0.1% trifluoroacetic acid (TFA) in water, B: acetonitrile; 0 to 100% B linear gradient 20 min, 1 mL/min. [111In(p-NO2-Bn-neunpa-PSMA)]- (tR = 11.2 min) and  “111In3+” (tR = 3.0 min).   Synthesis of H4neunpa-cageligand, compound 1 H4neunpa-p-Bn-NCS (0.03 g, 0.35 mmol, 1.1 eq.) and metallocage-arm-NH2 (0.010 g, 0.32 mmol) were dissolved in 2 mL dry DMSO. The reaction mixture was stirred overnight at room temperature and solvents were removed in vacuo. The dry crude powder was washed with ethylacetate and acetone, before purification via semi-prep reverse-phase HPLC (10mL/ min, gradient A: 0.1% TFA in deionized water, B: acetonitrile, A: 95% to B: 100% for 25 min.; tR= 14.4 min) to yield the product as an off-white solid (0.01 g, 40.1 %). 1H NMR (400MHz, MeOD): 8.74 (s, 2H), 8.56-8.54 (d, 2H), 8.02-7.97 (m, 4H), 7.92-7.90 (t, 2H), 7.75 (s, 2H), 7.60-7.58 (t, 3H), 7.52-7.50 (m, 2H), 7.41-7.39 (d, 2H), 7.24-7.22 (d, 2H) 13C NMR (100MHz, MeOD): 172.9, 150.6, 147.7, 139.6, 138.7, 130.7, 129.0, 127.4, 126.8, 124.5, 123.9, 123.9, 57.4, 55.1, 54.4, 50.8, 48.5, 47.9 and 29.1. HR-ESI-MS calcd. for [C51H47N9O8S+H]+ 946.3347; found: 947.3359 [M+H]+. Labelling of neunpa-cageligand with La (cold complexation), compound 2 H4neunpa-cageligand (0.01 g, 0.13 mmol) was dissolved in 1 mL dist. H2O and La(ClO4)3 * 6 H2O (0.01 g, 0.14 mmol) was added and the pH adjusted with 0.1 M NaOH to pH 4. The product precipitated as a white solid, filtered off and washed with water.  HR-ESI-MS calcd. for [C51H44LaN9O8S+H]+ 1082.2175; found:1082.2163 [M+H]+ Fluorescence Spectroscopy The emission spectra were recorded on a Varian Cary Eclipse fluorescence spectrophotometer. For each compound, dilutions in DMSO at a concentration of 10 mM were prepared. First UV/vis spectra of the compounds were recorded in DMSO, to determine the wavelength of the absorbance maximum. The measured absorbance wavelength was used as the excitation wavelength for the emission spectra.  Radiolabeling of H4neunpa-p-Bn-NO2 with 225Ac 10 µL of a ligand stock solution (10 and 1 mg/mL in water) was added to 130 µL of NH4OAc (0.15 M, pH 5), to this solution 10 µL of 225Ac3+ (~750 nCi) was added, mixed gently with a pipette, and left to react at room temperature. 5 µL aliquots were removed at 0, 15, 30, 60 min, and 2h or 4 h and spotted on iTLC-SG plates. Plates were developed in 10% NaCl/10mM NaOH. With this mobile phase, 'free' Ac3+ and Bi3+ remain at the base line (Rf = 0), and complexed metal migrates up the plate (Rf > 0). Plates were counted on a TLC plate reader immediately after development, and 3 hours later. Sb-complexation (cold) DFO (5.7 mg) was dissolved in 2 mL dist. H2O, Sb2O5 (s) was added and the pH adjusted to pH 4. A white precipitation was observed. The precipitate was filtered off, washed with water and the Sb-complex was confirmed via HR-ESI-MS calcd. for [C25H46N6O9Sb]+ 695.2364; found: 695.2358.  		77	SAHA (11.1 mg) was dissolved in 2 mL DMSO. After adding Sb2O5 (s), the solution changed its color from colorless to yellow. HR-ESI-MS calcd. for [C28H38N4O6Sb]+ 647.1830; found: 647.1833. H4neunpa-p-Bn-NO2 (5.6 mg) was dissolved in 2 mL dist. H2O and Sb2O5 (s) was added and the pH adjusted to pH 4. White precipitation was observed. The precipitate was filtered off and washed with water. HR-ESI-MS calcd. for [C30H32N6O10Sb]+ 757.1218; found: 757.1212. 5 References  1. http://www.who.int/mediacentre/factsheets/fs297/en/ (accessed 17th June 2017). 2. Bourgeois, M.; Bailly, C.; Frindel, M.; Guerard, F.; Cherel, M.; Faivre-Chauvet, A.; Kraeber-Bodere, F.; Bodet-Milin, C., Radioimmunoconjugates for treating cancer: recent advances and current opportunities. Expert Opin. Biol. Ther. 2017, 17 (7), 813-819. 3. Chang, S. S., Overview of Prostate-Specific Membrane Antigen. Rev. Urol. 2004, 6 (Suppl 10), S13-S18. 4. Bouchelouche, K.; Choyke, P. L.; Capala, J., Prostate Specific Membrane Antigen—A Target for Imaging and Therapy with Radionuclides. Discov. Med. 2010, 9 (44), 55-61. 5. Mease, R. C.; Foss, C. A.; Pomper, M. G., PET Imaging in Prostate Cancer: Focus on Prostate-Specific Membrane Antigen. Curr. Topics Med. Chem. 2013, 13 (8), 951-962. 6. Spreckelmeyer, S.; Ramogida, C. F.; Rousseau, J.; Arane, K.; Bratanovic, I.; Colpo, N.; Jermilova, U.; Dias, G.; Dude, I.; Jaraquemada Pelaez, M. G.; Benard, F.; Schaffer, P.; Orvig, C., p-NO2-Bn-H4neunpa and H4neunpa-Trastuzumab: Bifunctional Chelator for Radiometalpharmaceuticals and 111In Immuno-SPECT Imaging. Bioconjug. Chem. 2017. 7. Spreckelmeyer, S.; Ramogida, C. F.; Rousseau, J.; Arane, K.; Bratanovic, I.; Colpo, N.; Jermilova, U.; Dias, G.; Dude, I.; Jaraquemada Peláez, M. d. G.; Benard, F.; Schaffer, P.; Orvig, C., p-NO2-Bn-H4neunpa and H4neunpa-Trastuzumab: Bifunctional Chelator for Radiometalpharmaceuticals and 111In Immuno-SPECT Imaging. Bioconjug. Chem. 2017. 8. Schmidt, A.; Molano, V.; Hollering, M.; Pothig, A.; Casini, A.; Kuhn, F. E., Evaluation of New Palladium Cages as Potential Delivery Systems for the Anticancer Drug Cisplatin. Chemistry 2016, 22 (7), 2253-6. 9. Zhao, Q.; Liu, X.-M.; Li, H.-R.; Zhang, Y.-H.; Bu, X.-H., High-performance fluorescence sensing of lanthanum ions (La3+) by a polydentate pyridyl-based quinoxaline derivative. Dalton Trans. 2016, 45 (26), 10836-10841. 10. Wang, F.; Huang, W.; Li, K.; Li, A.; Gao, W.; Tang, B., Study on the fluorescence enhancement in Lanthanum(III)–carminic acid–cetyltrimethylammonium bromide system and its analytical application. Spectrochimica Acta Part A: Molecular and Biomolecular Spectroscopy 2011, 79 (5), 1946-1951. 11. Thisgaard, H.; Jensen, M., Sb119—A potent Auger emitter for targeted radionuclide therapy. Med. Phys. 2008, 35 (9), 3839-3846. 12. Robertson, A. K. H.; Ramogida, C. F.; Rodríguez-Rodríguez, C.; Stephan, B.; Peter, K.; Vesna, S.; Paul, S., Multi-isotope SPECT imaging of the 225 Ac decay chain: feasibility studies. Physics Med. Biol. 2017, 62 (11), 4406. 13. Chappell, L. L.; Deal, K. A.; Dadachova, E.; Brechbiel, M. W., Synthesis, Conjugation, and Radiolabeling of a Novel Bifunctional Chelating Agent for 225Ac Radioimmunotherapy Applications. Bioconj. Chem. 2000, 11 (4), 510-519. 				78	   Supporting Information Chapter A2 H4neunpa: A Bifunctional Acyclic Chelator with many Faces    Sarah Spreckelmeyer,a,b Caterina Ramogida,c Ting Kuo,d Ben Woods, e Valery Radchenko,c Maria Guadalupe Jaraquemada Peláez,a Francois Bénard,d Angela Casinie and Chris Orviga a Medicinal Inorganic Chemistry Group, Department of Chemistry, University of British Columbia, 2036 Main Mall, Vancouver, British Columbia, V6T 1Z1, Canada b Department of Pharmacokinetics, Toxicology and Targeting, Groningen Research Institute of Pharmacy, University of Groningen, Antonius Deusinglaan 1, Groningen 9713 AV, The Netherlands c Life Sciences Division, TRIUMF, 4004 Wesbrook Mall, Vancouver, British Columbia, V6T 2A3, Canada d BC Cancer Agency, 675 West 10th Avenue, Vancouver, British Columbia, V5Z 1L3, Canada e School of Chemistry, Cardiff University, Park Place, CF103AT Cardiff, United Kingdom 			79	 Scheme S 1. Synthesis of Glu-ureido-lys (here: HTK-01068). Synthesis of of HTK-01018: A solution of L-glutamic acid di-tertbutyl ester hydrochloride (1.5 g, 5.07 mmol) and triethylamine (2.31 mL, 16.63 mmol) in CH2Cl2 (40 mL) was cooled to −78 °C in a dry ice/acetone bath. Triphosgene (525 mg, 1.77 mmol) dissolved in CH2Cl2 (10 mL) was added dropwise to the reaction. After the addition was complete, the reaction was allowed to warm to room temperature and stirred for 30 minutes. H-Lys(cbz)-OtBu hydrochloride (1.5 g, 4.06 mmol) was then added to the reaction mixture, followed by triethylamine (566 μL, 4.06 mmol). After stirred overnight for 17 h, the reaction mixture was diluted with CH2Cl2 (50 mL) and washed with H2O (60 mL × 2). The organic phase was then dried over anhydrous magnesium sulfate and concentrated under reduced pressure. The residue was purified by chromatography on silica gel eluted with 3:2 hexane/EtOAc to obtain the desired product HTK-01018 as colorless oil (2.32 g, 92.3 %). Synthesis of of HTK-01027: A solution of HTK-01018 (2.32 g, 4.47 mmol) in MeOH (45 mL) was slowly added Pd/C (117 mg, wet by 5~10 mL MeOH) to the reaction. The reaction mixture was hydrogenated at room temperature under 1 atm. After stirred overnight, the solution was filtered through celite and concentrated under reduced pressure to obtain HTK-01027 as viscous oil (1.81 g). The crude product of HTK-01027 was used in next step without further purification. Synthesis of of HTK-01068: A solution of HTK-01027 (203 mg, 0.32 mmol) in TFA (5 mL) followed by 3% anisole was stirred at room temperature. After 4 h, the reaction mixture was concentrated under reduced pressure. The concentrate diluted with water (1 mL) and extracted with hexane (1 mL × 3) to remove anisole. The water phase was then iced and lyophilized to obtain crude HTK-01068 as a yellow oil. The crude product of HTK-01068 was used in next step without further purification. H2NOOt-BuO Ot-Bu a. Triphosgene    Et3N, CH2Cl2, -78oC, 30 minb. H-Lys(Cbz-OtBu), dropwise     warm to room Temp, Et3N, 17 hrNHNHNHCbzOOt-BuOO Ot-BuOOt-BuNHNHNH2OOt-BuOO Ot-BuOOt-BuPd/CMeOHHTK-01018HTK-01027HClTFA3 % AnisoleNHNHNH2HOOOO OHOHO4 hrHTK-01068			80	  Figure  S1.  13C NMR spectrum of compound 1.   Figure  S1.  2D-HSQC NMR spectrum of the aromatic region of compound 1 (400MHz, MeOD, 25°C).   30405060708090100110120130140150160170180190200f1	(ppm)53.2754.3955.1157.46123.87126.73127.40129.00138.55139.66139.87150.81172.927.17.27.37.47.57.67.77.87.98.08.18.28.38.48.58.68.78.88.9f2	(ppm)120122124126128130132134136138140142144146148150152f1	(ppm){8.75,150.58}{8.57,147.72}{8.05,139.62}{7.94,138.69}{7.59,130.73}{7.25,129.02}{7.76,127.36}{7.61,126.78}{7.42,124.48}{8.04,123.87} {7.53,123.87}			81	  Figure  S2.  2D-HSQC NMR spectrum of the alkyl region of compound 1 (400MHz, MeOD, 25°C).    Figure  S3.  2D-COSY NMR spectrum of the aromatic region of compound 1 (400MHz, MeOD, 25°C). 3.03.13.23.33.43.53.63.73.83.94.04.14.24.3f2	(ppm)24262830323436384042444648505254565860f1	(ppm){4.05,57.44}{3.59,55.06}{3.50,54.42}{3.68,50.82}{3.27,48.52}{3.32,47.93}{3.06,29.08}6.76.86.97.07.17.27.37.47.57.67.77.87.98.08.18.28.38.48.58.68.78.88.99.09.1f2	(ppm)6.66.87.07.27.47.67.88.08.28.48.68.89.09.2f1	(ppm){7.50,8.56}{7.51,8.04}{7.60,7.92}{7.94,7.60}{8.55,7.51} {8.04,7.50}			82	  Figure  S4. 2D-COSY NMR spectrum of the aromatic region of compound 1 (400MHz, MeOD, 25°C). 									6.76.86.97.07.17.27.37.47.57.67.77.87.98.08.18.28.38.48.58.68.78.88.99.09.1f2	(ppm)6.66.87.07.27.47.67.88.08.28.48.68.89.09.2f1	(ppm){7.50,8.56}{7.51,8.04}{7.60,7.92}{7.94,7.60}{8.55,7.51} {8.04,7.50}	 83					 Chapter A3 Tetrahydroxamic Acid Bearing Ligands:  EDTA and DTPA Analogues 													Sarah Spreckelmeyer,a,b Yang Caoa and Chris Orviga  a Medicinal Inorganic Chemistry Group, Department of Chemistry, University of British Columbia, 2036 Main Mall, Vancouver, British Columbia, V6T 1Z1, Canada b Department of Pharmacokinetics, Toxicology and Targeting, Groningen Research Institute of Pharmacy, University of Groningen, Antonius Deusinglaan 1, Groningen 9713 AV, The Netherlands 	84		1 Abstract  Hydroxamic acids are unique cation chelators that find application in the treatment of many diseases. Two tetra hydroxamic acid bearing chelators H4EDT(M)HA and H4EDT(B)HA were successfully synthesized and characterized as metal chelators. They were tested for their anticancer activity and 89Zr radiolabeling properties for positron emission tomography (PET) imaging. Although, Fe3+, Zn2+ and Cu2+ cold metal complexation reactions were achieved as proven by IR, NMR spectroscopies, HR ESI-MS spectrometry and UV-VIS experiments, 89Zr radiolabeling did not lead to any radiolabeling under the tested conditions. With DFT calculations and UV-VIS stability experiments, metal-EDT(M)HA complexes were found unstable due to the inflexibility of the four hydroxamic acid arms and the formation of complexes in a 2:2 ratio of chelator:metal. A second set of hydroxamic acid bearing chelators was designed and evaluated with density functional theory (DFT) and found to have theoretically a better stability of 89Zr radiolabeling. The synthesis of these promising DTPA analogues is currently ongoing. 	 85		 2 Introduction Hydroxamic acids have a long history as biological and chemical agents. In 1869, W. Lossen discovered the first hydroxamic acid, oxalohydroxamic acid.1 Beside their use as insecticides, antimicrobials, plant growth regulators, antioxidants, corrosion inhibitors or redox switches for electronic devices, hydroxamic acids have been identified as a key functional group for the development of pharmacological agents. Several drugs against hypertension, cancer growth, inflammation, infectious agents, asthma and arthritis that contain hydroxamic acid functional groups are used in the clinic. This application versatility is due to their common mechanism of action: their ability to chelate metal ions as well as to form a hydrogen bond at the active site of enzymes. They are selective and potent inhibitors of enzymes such as zinc metalloproteases (e.g. matrix metalloproteinases MMPs, angiotensin converting enzymes ACE, leukotriene A4 hydrolase LTA4), nickel metalloproteases (e.g. urease) and iron metalloproteases (e.g. lipoxygenase 5-LO, peptide deformilase PDF).2 Furthermore, it is suggested that hydroxamic acids are nictric oxide (NO) donors, the reason for their use against hypertension. Consequently, it is beyond dispute that hydroxamic acids are attractive tools in drug-design for a variety of diseases. In addition to their use as therapeutics, hydroxamic acids have been recently found to be attractive compounds for imaging purposes due to their chelation properties of radiometals.3  The chemical structure of hydroxamic acids is RC(O)N(R1)OH, where R1 can be either hydrogen or an alkyl moiety. They belong to a class of organic acids, which are weaker acids (acetohydroxamic acid pKa = 8.70) than structurally related carboxylic acids (acetic acid pKa = 4.75). Hydroxamic acids can either act as a monodentate ligand through the deprotonated hydroxyl (OH) moiety, or as bidentate monoanionic di-oxygen ligands, strongly prone to complex di- and trivalent metal cations. Hydrogen bonds can be formed through their OH group, amine (NH) group and carbonyl (C=O) oxygen. Their iron and copper complexes are highly coloured and are used for spectrophotometric and gravimetric analysis.4 Hydroxamic acids exhibit keto-iminol tautomerism (Figure 1). If the nitrogen is not substituted, hydroxamic acids exist preferably in the keto-Z or keto-E form over the iminol-Z or iminol-E. N-substituted hydroxamic acids on the other hand, can only exist in keto-Z or keto-E form, since they lack the NH group necessary for hydrogen transfer. In solution, the keto-E form seems to be more stable compared to the keto-Z form by 0.8 kcal/mol.4 In order to chelate metal ions, the acid should adopt the required Z-conformation; for that, interconversion between the Z and E conformation must take place by rotation about the C-N bond. The rotational barrier for N-substituted and N-non-substituted hydroxamic acids is between 16.6 kcal/mol and 20.2 kcal/mol.    Figure  1. Keto-iminol tautomerism of hydroxamic acids.  A special class of hydroxamic acid bearing compounds are siderophores (Greek: iron carrier), metal chelators that are low molecular weight agents, expressed by microorganisms in order to scavenge insoluble Fe3+ cations.3 Iron is essential for organisms to maintain DNA synthesis and respiration. The hydroxamic acids bearing agent desferrioxamine B (DFO, Desferal, Figure 2) is FDA approved for the treatment of iron poisoning or thalassemia.5 Due to their strong chelation to metal ions like Fe3+, Cu2+ or Ni2+, siderophores have been recently considered for molecular imaging applications, reviewed by Petrik et al.3  Commonly used radiometals for imaging purposes are 68Ga (t1/2 = 68 min), 11C (t1/2 = 20 min) or 18F (t1/2 = R NHOHOR NHOHOR NOHOHR NOHOHketo-Z keto-Eiminol-Z iminol-E	86		109 min) for PET imaging. The half-lives of these radiometals are quite short. Consequently, a short time for the tracer preparation is thus the limiting step and on-site preparation is often necessary to avoid loss of radioactivity during transportation. Thus, extensive research has been focused on radiometals with longer half-lives for PET imaging. 89Zr is an attractive radionuclide for this purpose, with a half-life of 3.3 days, which suits the biological half-life of biomolecules like antibodies. The development of a stable chelator for 89Zr is a challenging and a highly pursued goal of many research groups. To date, 89Zr-DFO-labelled antibodies are used off-label in the clinic to detect various types of cancer depending on the antibody (e.g. 89Zr-DFO-retuximab for B-cell non-Hodgkin lymphoma), although it bears major drawbacks like high bone uptake of 89Zr due to an unstable metal-ion chelate.6 It is hypothesized, that the preferred coordination number of Zr4+ is 8, thus DFO does not fulfill this requirement with its hexadentate chelating groups (Figure 2).7 Recently, several research groups have developed cyclic and acyclic chelators bearing hydroxamic acid motifs for 89Zr chelation, summarized in Figure 2.8,9,10,11 They were tested for 89Zr radiolabeling in vitro and in vivo. Noteworthy, the acyclic chelator DOTA was successfully labeled with 89Zr and a crystal structure of the “cold” Zr4+ complex was obtained, showing an octadentate binding to the metal centre by four nitrogens and four oxygens.12 Most recently in July 2017, C. Buchwalder et al. published a paper on an octadentate 3,4-HOPO chelator called THPN, that quantitatively complexates 89Zr4+ and shows promising in vitro and in vivo results (Figure 2).13     Figure  2. Hydroxamic acids bearing ligands discussed in this work and macrocyclic DOTA. The hydroxamic acid functional groups are given in red.  Due to the great success of ethylenediaminetetraacetic acid (EDTA) in stable metal complexation reactions14, we report here the synthesis, characterization as well as biological evaluation of the two EDTA analogues with four hydroxamic acid moieties (ethylenediaminetetra(methyl)hydroxamic acid H4EDT(M)HA and NOHOHNN OHONHOONHONCSOHNONHO ON OHOHNNHOONHOONHONH2ODFON NHNNOON OHOONHOOONOHNOON NON OHONHOONHOO N OHNCS-Rousseau et al.9Boros et al.8 Vugts et al.10NNNNOOOOOO OOOHOH HOHODeri et al.7N NNNDOTAOOHOHOOOHOHON NOHNO NHOHNONHNNN NHOOOO OOHOH HOBuchwalder et. al.13	 87		ethylenediaminetetra(benzyl)hydroxamic acid, H4EDT(B)HA) (Figure 3, left side).    Figure  3. Novel tetrahydroxamic acids bearing ligands discussed in this work.  Cold metal complexation reactions with Fe3+, Cu2+ and Zn2+ were studied via HR ESI-MS, IR spectroscopy and UV-VIS spectroscopy. In addition, to study their potential use as anti-cancer agents, the antiproliferative effects of H4EDT(M)HA compared to desferrioxamine (DFO) and suberanilohydroxamic acid (SAHA, Vorinostat) were tested in HT-29, MCF-7 and MCF-10A cell lines. Moreover, their use as 89Zr chelators was assessed in radiolabeling experiments. Additionally, density functional theory (DFT) studies were performed to explain the stability found for the complexes and to contribute to the design of a second generation hydroxamic acids bearing diethylenetriaminepentaacetic acid (DTPA) analogues DTT(ME)HA, DTT(MP)HA, DTT(BE)HA and DTT(BP)HA (Figure 3, right side) as potential 89Zr chelators for PET imaging.     3 Results and Discussion  3.1 Synthesis and characterization Based on the structure of EDTA, the hydroxamic acid analogue ethylenediaminetetra(methyl)hydroxamic acid, H4EDT(M)HA, was synthesized (Scheme 1).  Starting from N-methylhydroxylamine hydrochloride (Scheme 1a), the secondary amine was protected with a tert-butyloxycarbonyl (BOC) group15 1. In a next step, the hydroxyl group was deprotonated with sodium hydride under an inert atmosphere, following a benzylation reaction with benzylbromide to yield 2.16 After deprotection of the BOC group with TFA/DCM 1:1 to yield 3, bromoacetylbromide was added17 to synthesize the “arm“ of NN NONHOONOHOHONHOONN N N NOHOOOHOHOOOHOOHOOHOHOOOHOOHEDTA DTPAH₄EDT(M)HANN NONHOONOHOHONHOOH₄EDT(B)HAn	=	2	DTT(ME)HA				=	3	DTT(MP)HANN NONHOONOHOHONHOOnnnnNHn	=	2	DTT(BE)HA				=	3	DTT(BP)HANN NONHOONOHOHONHOOnnnnNH1-.	generation 2ⁿ:	generation	88		H4EDT(M)HA 4. In a next reaction (Scheme 1b), ethylenediamine was added to the “arm“ 4 to yield the protected intermediate 5. The product was synthesized by palladium-catalyzed debenzylation to produce H4EDT(M)HA 6 in an overall yield of 1.5 %. The yield limiting step is most likely the addition of the “arms” to the backbone, since three different side-products were observed, which are either the mono, di or tri functionalized backbone. The product was fully characterized using 1H NMR (Figure 4), 13C NMR, 2D-HSQC spectroscopy (Figure S 1) and HR ESI-MS.    Scheme 1. Synthesis route of H4EDT(M)HA.  H4EDT(B)HA was synthesized starting with O-benzylhydroxylamine hydrochloride (Scheme 2) and benzaldehyde to form its imine intermediate (not shown). For the reduction of the imine, sodium cyanoborohydride was used at pH 4 to yield 7.18 In a next reaction, bromoacetyl bromide was added to synthesize the “arm” of H4EDT(B)HA 8. After linking the arm to ethylenediamine 9 (Scheme 2b), the O-benzyl groups were removed by palladium catalyzed hydrogenation to produce the final ligand H4EDT(B)HA 10 in an overall yield of 0.35 %. 1H NMR (Figure 4) and 13C NMR confirmed the successful synthesis.             Scheme 2.  Synthesis route of H4EDT(B)HA. HN OH NOHOONOOOHN OTEA, DCM(BOC)2ONaH, DMFBrTFA/DCM (1:1)1 2 3NOBrOBr XOnnn = 1 arm for EDT(M)HA,   4   = 2             DTT(ME)HA,  4a   = 3             DTT(MP)HA, 4bK2CO3, THFa)b) H2N NH2K2CO3, CH3CNNN NONOONOOO NOOPd(OH)2/C, H2MeOHNN NONHOONOHOHO NHOO 5 H4EDT(M)HA,  64H2N OHN OOMeOH, HCl, NaCNBH3N OBrO7Br XOK2CO3, THFnnn = 1 arm for EDT(B)HA,  8   = 2             DTT(BE)HA, 8a   = 3             DTT(BP)HA, 8ba)NN NONOONOOONOO9 H4EDT(B)HA,  10NN NONHOONOHOHONHOOb) 8 H2N NH2K2CO3, CH3CNPd(OH)2/C, H2MeOH	 89		              Figure  4. 1H NMR spectra of H4EDT(M)HA (top, 400 MHz, D2O, 25°C) and H4EDT(B)HA (bottom, 400 MHz, MeOD, 25°C).    3.2 Metal complexation reactions To determine, whether the synthesized ligands H4EDT(M)HA and H4EDT(B)HA are chelators for metal ions, Fe3+, Cu2+ and Zn2+ were chosen as model ions for metal complexation reactions because of their roles in biological processes. As mentioned above, many enzymes bear these metal ions in their active site, which can be chelated by hydroxamic acids. Fe3+ is a hard acid, paramagnetic and forms coloured complexes. Cu2+ is a hard acid and paramagnetic as well. Zn2+ is diamagnetic. As proven by HR ESI-MS (see Materials and Methods section), H4EDT(M)HA and H4EDT(B)HA form metal-ligand complexes in a 1:1 ratio with each of the three metals. 1H NMR spectra of Fe(III)-EDT(M)HA and Cu(II)-EDT(M)HA show the expected changes in chemical shifts (Figure S 2), due to metal complexation. However, the 1H NMR spectra do not allow proper peak assignments due to the paramagnetism of these two metals. Therefore, IR as well as UV VIS spectroscopy is applied for further characterizations. A 1H NMR spectrum of Zn-EDT(M)HA was not recorded, the Cu-EDT(M)HA and Fe-(EDT(M)HA spectra were considered sufficient as a proof of principle.  3.3 Infrared (IR) spectroscopy IR is a useful tool to determine vibrational modes; symmetric and antisymmetric stretching as well as bending modes are the typical vibrational modes that are observed. The carbonyl bond of hydroxamic acids usually show a characteristic peak around 1650 cm-1 in the IR spectra, and they can easily be distinguished from other carbonyl bonds, such as carboxylic acids that show absorption above 1700 cm-1. A successful metal complexation would be indicated by a shift of the characteristic peak of the hydroxamic acids, due to changes in the vibrational modes, to less frequent vibrations.19 For proof of principle, the infrared spectra of the H4EDT(M)HA, Fe-EDT(M)HA and Cu-EDT(M)HA are shown in Figure 5 and the characteristic shifts for the functional groups of the hydroxamic acids are summarized in Table 1. The ligand itself shows a typical carbonyl stretch at 1660 cm-1. After complexation, the carbonyl stretches of the Fe-complex and Cu-complex shift to lower frequencies, 1628 cm-1 and 1634 cm-1, respectively. This behaviour has been reported before for hydroxamic acid complexes with Ni2+, Co2+ and Zn2+.20 This finding indicates a coordination of the ligand with the metal ion through the oxygen of the carbonyl functional group. The symmetric N-O stretch shifts to lower frequencies as well, indicating that the hydroxamate complexation is bidentate. These two results give an indication that the 3.263.634.364.79	D2O2.53.03.54.04.55.05.56.06.57.07.5f1	(ppm)3.31	CD3OD3.374.264.767.33methylenN-CH2-CH2-NN-CH3O-CH2methylenN-CH2-CH2-NbenzylH2O	90		metals are chelated via bidentate bis-oxygen chelation. Moreover, an intense stretch at about 1200 cm-1 was observed for the ligand as well as for the metal complexes, which can be assigned as C-N stretch. This stretch does not shift upon chelation, suggesting that the C-N bond is not involved in the metal complexation. Due to N-substitution of the ligand, the iminol form is not an option, making the involvement of the C-N bond in chelation unlikely (for full spectra, see Supporting Information, Figure S 3).    Figure  5. Partial IR spectra of H4EDT(M)HA, Fe(III)-EDT(M)HA and Cu(II)-EDT(M)HA at 25°C (solid state). 	Table  1. Summarized IR data of the relevant functional groups. Functional group H4EDTMHA [cm-1] Fe(III)-EDTMHA [cm-1] Cu(II)-EDTMHA [cm-1] N-O 1138 1055 1068 C=O 1660 1628 1634  3.4 In vitro cell experiments Based on the successful metal complexation, H4EDT(M)HA was tested for its possible toxic effects towards two cancer cell lines (MCF-7: breast cancer and HT-29: colon cancer) compared to a non-tumorigenic cell line (MCF-10A: epithelial healthy breast cells). We hypothesize that H4EDT(M)HA shows toxicities similar to desferoxamine (DFO, Desferal®) and suberanilohydroxamic acid (SAHA, Vorinostat®) which have been used as controls. DFO has already shown anti-proliferative and cytotoxic effects on several tumour cell lines, due to its chelation ability of iron and thereby removing the iron from an iron-dependent enzyme necessary for cell cycle progression.21 SAHA is an anticancer agent that was approved in 2006 by the FDA. SAHA binds Zn2+ and thereby stays in the active site of histone deacetylases.22 The results of three independent cell experiments with H4EDT(M)HA, DFO and SAHA are shown in Table 2. For DFO the expected low IC50 values of 9.5 ± 2.4 µM, 15.0 ± 2.7 µM and 6.4 ± 0.8 µM were found for HT-29, MCF-7 and MCF-10A cells, respectively.23,24 SAHA shows the highest toxicity, with IC50 values of 1.0 ± 0.3 µM, 0.8 ± 0.5 µM and 1.5 ± 0.5 µM for these cell lines. H4EDT(M)HA shows very low toxicity towards HT-29, MCF-7 and 600800100012001400160018008090100wavenumber [cm-1]Transmittance [%]H4EDT(M)HAFe-EDT(M)HACu-EDT(M)HA	 91		MCF-10A cells with IC50 values above 100 µM. A reliable IC50 determination was not possible above 100 µM of the compound, since these high concentrations required a DMSO concentration above 1.0 % in the final cell incubation medium, which affected the cell viability.  Table  2. Cytotoxicity of DFO, SAHA and H4EDT(M)HA on HT-29, MCF-7 and MCF-10A cells, expressed as IC50 [µM] (n = 3 ± SD).  Cell line/compound HT-29 MCF-7 MCF-10A DFO 9.5 ± 2.4 15.0 ± 2.7 6.4 ± 0.8 SAHA 1.0 ± 0.3 0.8 ± 0.1 1.5 ± 0.5 H4EDT(M)HA >100 >100 >100  The low toxicity of H4EDT(M)HA could have different reasons among which a low stability of the metal-EDT(M)HA complexes or a low uptake into the cells can be considered. Hydroxamic acids are very hydrophilic functional groups and H4EDT(M)HA incorporates four hydroxamic acid moieties. Thus, membrane passage by passive diffusion of H4EDT(M)HA seems unlikely due to its hydrophilicity. Alternatively the complex might cross the cell membrane via channels or carriers. Further experiments are needed to get more information about the uptake mechanisms of H4EDT(M)HA. As mentioned above, instability of the metal-EDT(M)HA complexes might be the reason for the low toxicity. Therefore, the stability of the complexes was determined by UV-VIS spectroscopy.   3.5 Stability determinations of Fe-EDT(M)HA and Fe-EDT(B)HA by UV-VIS spectroscopy UV-VIS spectroscopy is a technique based on electronic transitions in atoms, molecules or ions. Fe(III), in contrast to Zn(II), is a unique cation that changes its colour upon chelation. Thus, the successful iron complexation can easily be seen by a color change of the Fe(III)-solution from yellow to red after adding H4EDT(M)HA or H4EDT(B)HA. The intensely coloured complexes can be studied by UV-VIS spectroscopy. Fe(III) belongs to the d-block elements, with a partly 3d filled orbital shell. Upon complexation, the d orbitals split into two different energy levels, two orbitals with a higher energy level and three orbitals with a lower energy level. Upon absorption of light, an electron is promoted from the lower energy level to the higher energy level. The greater the difference in energy between the levels, the more energy is needed for this promotion. In Figure 6, the changes in UV-VIS spectra of Fe(III)-EDT(M)HA or Fe(III)-EDT(B)HA at 37 °C in water over a time period of 5 days are shown. An absorption maximum at 490 nm for Fe(III)-EDT(M)HA and at 500 nm for Fe(III)-EDT(B)HA correlates well with the red coloured solutions. Fe(III)-EDT(M)HA shows a sharper absorption peak compared to Fe(III)-EDT(B)HA. The UV VIS spectrum of Fe(III)-EDT(B)HA might be broader due to the co-absorbance of the benzyl groups of the chelator. A decrease of the absorbance over 5 days indicates an unstable Fe-complex for both ligands. Notably, the absorbance of Fe(III)-EDT(M)HA decreases faster than that of Fe(III)-EDT(B)HA. Concerning the in vitro IC50 values, the low stability of the Fe(III)-complexes shown here, might at least partly explain the low toxicity in vitro. The cell experiments were conducted after three days, when still 50 % of the Fe-complexes were intact (Figure 6). However we have to consider, that for the cell experiments, other factors may also play a role, like trans-chelation of the Fe-complexes eg. via transferrin. Moreover, the UV VIS experiments were performed in water, with only the tested compound and Fe3+ions present, whereas the cell experiments were performed in more complex cell culture medium.   	92		   Figure  6. UV-VIS spectra of Fe-EDT(M)HA (left) and Fe-EDT(B)HA (right) at 37°C over 5 days in H2O.  3.6 89Zr-radiolabeling Chelation of 89Zr was investigated to explore whether our ligands could be applied for PET imaging.  As mentioned above, hydroxamic acids are hard electron donors, and as such, a good fit for the hard acid cation Zr4+. Zirconium prefers a coordination number of 8, thus the tetrahydroxamic acid H4EDT(M)HA was assumed to be a good candidate for 89Zr radiolabeling. However, we could not observe any labelling of H4EDT(M)HA with 89Zr. Different radiolabeling media (phosphate buffered saline pH 7.4 (PBS) or 0.9 % NaCl solution pH 7.4 (saline)) were applied. The radiochemical yield (RCY) was determined via instant thin layer chromatography (iTLC). Different mobile phases (EDTA pH 7, citrate pH 5.5, DTPA pH 6) were applied for iTLC. Additionally, radio-HPLC was performed as well to get further information. However, 0 % radiolabeling of H4EDT(M)HA with 89Zr was observed whereas labeling of the gold-standard DFO always gave expected good RCYs. An example of an iTLC chromatogram can be found in the Supporting Information Figure S 4.  3.7 Density Functional Theory (DFT) In order to find an explanation for the low stability found in UV-VIS experiments for the Fe-EDT(M)HA complexes, and the lack of complexation with 89Zr, various DFT optimized structures were calculated.   Our aim was to determine, if Fe3+ or Zr4+ complexation can theoretically take place, taking into consideration the bond lengths and angles of the coordinative bonds between the metal and chelator. The results show that the ligand itself shows all four hydroxamic acids arms pointing away from each other, making it impossible to chelate a single metal ion in a 1:1 ratio (Figure 7, left), due to the inflexibility of the arms. Thus, no DFT structure could be calculated in a 1:1 metal-chelator ratio. The second DFT structure in Figure 7 shows a dimer of H4EDT(M)HA with two Fe3+ ions, indicating that a complex can theoretically be formed where the iron is coordinated with only six donor arms of the hydroxamic acids in a 2:2 ratio. The HR ESI-MS does only show the iron-complex in a 1:1 ratio. These findings explain the low stability of metal-EDT(M)HA complexes and can also explain the unsuccessful 89Zr radiolabeling.  In a DFT study of Boros et al.9, hydroxamic acid bearing cyclic DOTA analogues showed higher stability with metal ions, if the hydroxamic acid arms are longer than acetohydroxamic acid and if the cavity is bigger than ethylenediamine. In that case, incorporation of large metal ions like Zr4+ seems feasible. Thus, the chelation of 89Zr by a second set of proposed chelators (Figure 8) with longer hydroxamic acid donor arms was evaluated by DFT. By increasing the size of the backbone from ethylenediamine to diethylenetriamine and by increasing the hydroxamic acid arm length from a methylene group to ethylene or propylene, a more ideal geometry of the Zr-complexes were calculated (see bond lengths in the Supporting Information Figure S 5 and Figure S 6).  Additionally, the middle nitrogen atom can be functionalized without interfering with the chelation properties (Supporting Information, Figure S 7), which makes the new generation of DTPA analogues ideal bifunctional chelators for 89Zr for PET imaging. 400 500 600 700 8000.00.51.01.5wavelength [nm]Absorbance0d1d2d3d4d5d400 600 8000.00.51.01.5wavelength [nm]Absorbance0 d1 d2 d3d4d5d	 93		  Figure 7. DFT optimized structures  of H4EDT(M)HA (left) and [Fe2(EDT(M)HA)2] (right) (grey: carbon, blue: nitrogen, red: oxygen, orange: iron (B3LYP/6-31G* + LANL2DZ and PCM solvation (water) on Gaussian 09).   Figure  8. DFT optimized structure  of Zr-DETT(ME)HA (left) and Zr-DETT(MP)HA (right) (grey: carbon, blue: nitrogen, red: oxygen, light blue: zirconium (B3LYP/6-31G* + LANL2DZ and PCM solvation (water) on Gaussian 09).  Based on the DFT calculations, as is discussed above, we aimed to synthesize four second-generation DTPA analogues. The proposed synthesis routes for these four DTPA analogues diethylenetriaminetetra(methylethyl)hydroxamic acid (H4DTT(ME)HA), diethylenetriaminetetra(methylpropyl)hydroxamic acid (H4DTT(MP)HA), diethylentriaminetetra(phenylethyl)hydroxamic acid (H4DTT(BE)HA and diethylenetriaminetetra(phenylpropyl)hydroxamic acid (H4DTT(BP)HA) are shown in Scheme 3.  	94		  Scheme 3. Proposed synthesis route for f H4DTT(ME)HA, H4DTT(MP)HA, H4DTT(BE)HA and H4DTT(BP)HA.  Regarding H4DTT(ME)HA and H4DTT(MP)HA, their arms were synthesized starting with intermediate 3 (see Scheme 1a) and adding either 4-bromobutyryl chloride or 3-bromopropionyl chloride, respectively to the reaction mixture. The successful synthesis of the two hydroxamic acid arms 4a and 4b was characterized by 1H NMR spectroscopy (Figure 9), 13C NMR spectroscopy (Figure S 8) and HR ESI-MS.      Figure  9. 1H NMR spectra of arms (4a, top) and (4b, bottom), 400 MHz, CDCl3, 25°C.  The next reaction step, the attachment of the “arms” 4a or 4b to the backbone diethylenetriamine via a SN2 reaction, did not result in the anticipated products but always led to the E2 side-products (Figure 10). NN NONOONOOONOOn = 2 precursor of DTT(BE)HA    = 3                     DTT(BP)HAnnnnn = 2 DTT(BE)HA    = 3 DTT(BP)HAb) NHNN NONHOONOHOHONHOOnnnnNHPd(OH)2/C, H2MeOHH2N NHNH2NN NONOONOOONOOn = 2 precursor of DTT(ME)HA    = 3                     DTT(MP)HAnnnnNHn = 2 DTT(ME)HA    = 3 DTT(MP)HANN NONHOONOHOHONHOOnnnnNHa) Pd(OH)2/C, H2MeOHH2N NHNH24a4b8a8b1.82.02.22.42.62.83.03.23.43.63.84.04.24.44.64.85.05.25.45.65.86.06.26.46.66.87.07.27.47.67.88.0f1	(ppm)7.26	CDCl37.26	CDCl3benzylO-CH2 N-CH3Br-CH2-CH2Br-CH2-CH2benzylO-CH2Br-CH2-CH2-CH2 Br-CH2-CH2-CH2Br-CH2-CH2-CH2N-CH3	 95		Unfortunately, the conditions for a SN2 reaction are quite similar to the conditions of an E2 reaction. SN2 reactions require a strong nucleophile and E2 reactions require a strong base. Good nucleophiles are often strong bases. To obtain the required product, a good nucleophile, that is a weak base (eg. K2CO3, NaHCO3), would favour an SN2 reaction over an E2 reaction. The use of polar, aprotic solvents (eg acetone, DMF, acetonitrile and DMSO) might also increase the nucleophilicity and thus increase the rate of SN2. The conditions tried thus far are summarized in the Supporting Information (Table S1). Unfortunately, none of the tried conditions resulted in the anticipated products. Further work to synthesize the appropriate compounds is ongoing.    Figure  10.  E2 reaction schemes that yielded the unwanted side-products.   4 Conclusions  Hydroxamic acids are unique metal ion chelators and can be used for either therapy or diagnosis. Our first aim was the evaluation of new chelators for PET imaging, using 89Zr as radiometal. The synthesis of 89Zr chelating ligands for PET imaging is theoretically a promising goal but appeared challenging in practice. Hydroxamic acid bearing chelators were suggested to be the perfect functional groups for this purpose, as they are hard donors and perfectly fit the hard acid character of 89Zr. Our second aim deals with the evaluation of hydroxamic acid bearing ligands as anticancer agents due to the ability of hydroxamic acids to chelate essential metals like Cu2+, Fe3+ and Zn2+. In this work, two tetrahydroxamic acid bearing EDTA analogues were synthesized, which differ in their N-substitution. They were successfully characterized using 1H NMR, 13C NMR and 2D HSQC NMR spectroscopies. Metal complexations were confirmed via IR spectroscopy, UV-VIS spectroscopy and HR ESI-MS spectroscopy. The stability evaluation of Fe-EDT(M)HA and Fe-EDT(B)HA via UV-VIS shows limited metal complex stability of both ligands, which may explain the relatively low toxicity towards HT-29, MCF-7 and MCF-10A cell lines. DFT calculations further strengthened these findings by showing inflexible hydroxamic acid arms, which makes a metal chelation unlikely to occur. Thus, the synthesized chelators are not suitable as anticancer agents. The same instability hypothesis may explain the 0 % radiochemical yield of 89Zr labeling.  A second set of tetrahydroxamic acid bearing DTPA analogues with diethylentriamine as the backbone and longer hydroxamic acid arms was designed and evaluated by DFT for their potential ability to chelate Zr4+. The DTPA analogues show greater flexibility of the arms, making a metal complexation more favorable. The synthesis of the arms was successful. However, the linkage of the arms to the backbone was restrained by an E2 elimination reaction instead of SN2 substitution reaction thus far. Further experiments are currently underway to improve the synthesis of these promising complexes.  5 Experimental Materials and Methods N OON OOBrN OOBr NOOE2E24a4b	96		All solvents and reagents were from commercial sources (Sigma Aldrich, TCI) and were used as received unless otherwise noted. 1H and 13C NMR spectra were recorded at room temperature on a Bruker AV400 instrument; the NMR spectra are expressed on the δ (ppm) scale and are referenced to the residual solvent signal of the deuterated solvent. All spectra were recorded with sweep widths of 0-14 ppm or -20-220 ppm for 1H and 13C NMR respectively. Assignments of the peaks in the NMR spectra are approximate. Mass spectrometry was performed on a Waters ZQ spectrometer equipped with an electrospray source at the Department of Chemistry, University of British Columbia. The HPLC system used for purification of ligands and precursors consisted of a Waters 600 controller equipped with a Waters 2487 dual λ absorbance detector connected to a Phenomenex synergi hydro-RP 80Å 250mm x 4.60 mm semipreparative column. Infrared spectra were recorded using a Frontier FT-IR spectrometer purchased from PerkinElmer. UV absorbance measurements were recorded on an Agilent Technologies Cary 5000 UV-VIS spectrometer.	Synthesis of compounds N-Methyl-N-Boc- hydroxylamine15 ,1 N-Methylhydroxylamine hydrochloride (0.53 g, 6.4 mmol) was added to dichloromethane (50 mL) and the mixture was cooled to 0°C for 15 min. After that, di-tert-butyl dicarbonate (1.53 g, 7.0 mmol, 1.1 eq) was added in portions to the mixture. Triethylendiamine was added until the solution went clear (1.5 mL) and the reaction mixture was stirred overnight. Dichloromethane was removed by blowing air through the solution without heating, since the product is volatile and has a low boiling point. The resulting colorless oil 1 was dried for one day and used without further purification (70 %, 0.65 g). 1H NMR (400 MHz, CDCl3, 25°C): 3.09-3.06 (m, 3H), 1.38 (s, 9H). 13C NMR (101 MHz, CDCl3, 25°C): 157.3, 81.0, 38.3, and 28.3. HR-ESI-MS: calcd. for [C6H13NO3+H]+: 148.0974; found 148.0973 [M+H]+ 		t-Butyl- N-benzyloxy-N-methylcarbamate16 , 2 Compound 1 (0.65 g, 4.4 mmol) was placed in a two neck round bottom flask that was then evacuated; it was then flushed with argon, dry DMF (10 mL) added and the mixture placed in an ice-bath. Sodium hydride (0.13 g, 5.3 mmol, 1.2 eq.) was slowly added, turning the solution white and the development of foam (H2) was observed.  After 30 minutes, benzylbromide (0.63 mL, 5.3 mmol, 1.2 eq.) was slowly added to the solution, turning the solution clear again. The reaction mixture was stirred for 4 h at room temperature. Hexane was added to the solution and adding solvent dropwise with increasing polarity until no gas formation was observed any longer, destroying unreacted sodium hydride. The solvents were evaporated and the product was dried in vacuo and purified by silica chromatography (CombiFlash Rf automated system; 12 g HP silica; A: hexane, B: ethyl acetate, 100% A to 100% B gradient) to yield product 2 as a yellowish oil (48 %, 0.51 g). 1H NMR (400 MHz, CDCl3, 25°C): 7.46-7.36 (m, 5H), 4.87 (s, 2H), 3.09 (s, 3H), 1.54 (s, 9H). 13C NMR (101 MHz, CDCl3, 25°C): 156.9, 135.6, 129.4, 128.7, 128.4, 81.1, 76.4, 36.8, and 28.3.  HR-ESI-MS: calcd. for [C13H19NO3+H]+: 238.1443; found 238.1443 [M+H]+  O-Benzyl-N-methylhydroxylamine , 3 Compound 2 (0.51 mg, 2.2 mmol) was dissolved in TFA/DCM (1:1, 2 mL) and the reaction mixture stirred at room temperature for 4 h. The solvent was removed under reduced pressure and purified by silica chromatography (CombiFlash Rf automated system; 12 g HP silica; A: hexane, B: ethyl acetate, 100% A to 100% B gradient) to yield product 3 as a colorless oil (97 %, 0.29 g). 1H NMR (400 MHz, CDCl3, 25°C): 11.70 (s, 2H) 7.39 (s, 5H), 5.09 (s, 2H), 2.95 (s, 3H). 13C NMR (101 MHz, CDCl3, 25°C): 135.6, 129.4, 128.7, 128.4, 76.4, and 36.8. HR-ESI-MS: calcd. for [C8H11NO+H]+: 138.0919; found 138.0912 [M+H]+  O-Benzyl-2-bromo-N-methylacetohydroxamic acid17, 4 Compound 3 (0.29 g, 2.1 mmol) and potassium carbonate (0.50 g, 3.6 mmol, 1.75 eq.) were dissolved in dry tetrahydrofuran (3 mL) and after 15 min stirring at 0°C, bromoacetyl bromide (0.22 mL, 2.5 mmol, 1.2 eq.) was 	 97		added slowly to the solution, forming a white cloudy mixture. After 7 h stirring at room temperature, the suspension was filtered and the filtrate evaporated under reduced pressure. The residue was purified by silica chromatography (CombiFlash Rf automated system; 12 g HP silica; A: hexane, B: ethyl acetate, 100% A to 100% B gradient) to yield product 4 as a colorless oil (26 %, 0.14 g). 1H NMR (400 MHz, CDCl3, 25°C): 7.42 (s, 5H), 4.95 (s, 2H), 3.93 (s, 2H), 3.27 (s, 3H). 13C NMR (101 MHz, CDCl3, 25°C): 164.2, 135.6, 129.4, 128.7, 128.4, 76.4, 42.3 and 36.8. HR-ESI-MS: calcd. for [C10H12BrNO2+H]+: 259.0130; found 259.0132 [M+H]+  O-Benzyl-3-bromo-N-methylpropiohydroxamic acid17, 4a Compound 3 (0.5 g, 3.7 mmol) and potassium carbonate (0.89 g, 6.4 mmol, 1.75 eq.) were dissolved in dry tetrahydrofuran (10 mL) and after 15 min stirring at 0°C, 4-bromobutyryl chloride (0.44 mL, 4.4 mmol, 1.2 eq.) was added slowly to the solution, forming a white cloudy mixture. After 7 h stirring at room temperature, the suspension was filtered and the filtrate evaporated under reduced pressure. The residue was purified by silica chromatography (CombiFlash Rf automated system; 12 g HP silica; A: hexane, B: ethyl acetate, 100% A to 100% B gradient) to yield product 4 as a colorless oil (32 %, 0.32 g). 1H NMR (400 MHz, CDCl3, 25°C): 7.39 (s, 5H), 4.95 (s, 2H), 4.85 (s, 2H), 3.56 (t, 2H), 3.22 (s, 3H), 2.95 (t, 2H). 13C NMR (101 MHz, CDCl3, 25°C): 171.9, 134.2, 129.4, 129.2, 128.8, 76.4, 35.6, 33.5 and 26.6. ESI-MS: calcd. for [C11H14BrNO2+H]+: 272.01; found 272.1 [M+H]+  O-Benzyl-4-bromo-N-methylbutylhydroxamic acid17, 4b Compound 3 (0.15 g, 1.1 mmol) and potassium carbonate (0.26 g, 1.9 mmol, 1.75 eq.) were dissolved in dry tetrahydrofuran (10 mL) and after 15 min stirring at 0°C, 3-bromopropionyl chloride (0.15 mL, 1.3 mmol, 1.2 eq.) was added slowly to the solution, forming a white cloudy solution. After 7 h stirring at room temperature, the suspension was filtered and the filtrate evaporated under reduced pressure. The residue was purified by silica chromatography (CombiFlash Rf automated system; 12 g HP silica; A: hexane, B: ethyl acetate, 100% A to 100% B gradient) to yield product 4 as a colorless oil (20 %, 0.06 g). 1H NMR (400 MHz, CDCl3, 25°C): 7.39 (s, 5H), 4.83 (s, 2H), 3.44 (dt, 2H), 3.19 (s, 3H), 2.55 (t, 2H), 2.11 (t, 2H). 13C NMR (101 MHz, CDCl3, 25°C): 173.8, 134.4, 129.4, 129.1, 128.8, 128.6, 128.5, 127.5, 127.0, 76.3, 65.1, 33.8, 30.4, 28.3 and 27.3. ESI-MS: calcd. for [C11H14BrNO2+H]+: 272.01; found 272.1 [M+H]+  N-Benzyl-O-benzylhydroxylamine, 5 O-Benzylhydroxylamine hydrochloride (1.3 g, 8.12 mmol) was dissolved in methanol (20 mL) and triethylamine was added until complete dissolution. Benzaldehyde (0.91 mL, 8.93 mmol, 1.1 eq.) was added at room temperature and after 2 h the imine intermediate was confirmed via ESI-MS. After that, the white mixture was cooled to 0°C and acidified with conc. HCl. Sodium cyano- borohydride (1.53 g, 24.4 mmol, 3 eq.) was added slowly under cooling and the reaction mixture left stirring overnight. After removing the solvents in vacuo, 0.1 M sodium hydroxide solution (10 mL) was added to the residue and extracted with dichloromethane. The organic phase was dried and purified by silica chromatography (CombiFlash Rf automated system; 12 g HP silica; A: hexane, B: ethyl acetate, 100% A to 100% B gradient) to yield product 5 as a white solid (58 %, 1.02 g). 1H NMR (400 MHz, CDCl3, 25°C): 7.39 (m, 10H), 4.62 (s, 4H) 13C NMR (101 MHz, CDCl3, 25°C): 141.0, 128.5, 127.5, 127.1, 64.8, and 64.7. HR-ESI-MS: calcd. for [C14H15NO+H]+: 214.1232; found 214.1233 [M+H]+  O-Benzyl-2-bromo-N-benzyl-acetohydroxamic acid, 6 Compound 5 (1.02 g, 4.7 mmol) and potassium carbonate (0.71 g, 5.2 mmol, 1.1 eq.) were dissolved in dry tetrahydrofuran (20 mL) and after 15 min stirring at 0°C, bromoacetyl bromide (0.45 mL, 5.2 mmol, 1.1 eq.) was added slowly to the solution, forming a white cloudy suspension. After 7 h stirring at room temperature, the suspension was filtered and the filtrate evaporated under reduced pressure. The residue was purified by silica chromatography (CombiFlash Rf automated system; 12 g HP silica; A: hexane, B: ethyl acetate, 100% A to 100% B gradient) to yield product 6 as a white solid (43 %, 0.67 g). 1H NMR (400 MHz, CDCl3, 25°C): 7.42-7.32 (m, 10H), 4.89 (s, 4H), 3.98 (s, 2H). 13C NMR (101 MHz, CDCl3, 25°C): 	98		168.4, 135.7, 133.9, 129.4, 129.3, 128.9, 128.7, 128.6, 128.0, 77.2, 50.6 and 2.9. HR-ESI-MS: calcd. for [C16H16BrNO2+Na]+: 356.0262; found 256.0257 [M+H]+  N,N,N,N-Tetra-(O-benzyl-N-methylacetohydroxamic acid)-diaminoethane , 7 Compound 4 (0.06 g, 0.23 mmol, 4.4 eq.) was dissolved in acetonitrile (1 mL) and potassium carbonate (0.03 g, 0.23 mmol, 4.4 eq.) and ethylenediamine (3.5 μL, 0.05 mmol) were added to the solution. After 16 h stirring at room temperature, the solution was dried under reduced pressure and the residue purified by semi-prep RP HPLC (10 mL/ min, gradient A: 0.1% TFA in deionized water, B: acetonitrile, A: 95 % to B: 80 % for 10 min., followed by A: 20 % to B: 100 % for additional 20 min, tR= 15.25 min) to yield product 5 as a colorless oil (58.7 %, 0.003 g). 1H NMR (400 MHz, CDCl3, 25°C): 7.37 (s, 20H), 4.94 (s, 8H), 3.92 (s, 8H), 3.26 (s, 12H), 3.22 (m, 2H), 3.16 (m, 2H). 13C NMR (101 MHz, CDCl3, 25°C): 168.1, 133.7, 129.4, 128.7, 128.4, 76.3, 33.7 and 25.3. HR-ESI-MS: calcd. for [C42H52N6O8+H]+: 769.3925; found 769.3928 [M+H]+  H4EDT(M)HA (Ethylenediaminetetra(methylene-N-methylhydroxamic acid), 8 Compound 5 (0.03 g, 0.04 mmol) was dissolved in methanol (2 mL) and Pd(OH)2/C (20 w/w %, 0.007 g) was added to the solution. Charging the flask with a H2-filled balloon gave the product 6, after 2 h stirring at room temperature, as a colorless oil (74 %, 0.001 g). 1H NMR (400 MHz, CDCl3, 25°C): 4.36 (s, 8H), 3.63 (s, 4H), 3.26 (s, 12H). 13C NMR (101 MHz, CDCl3, 25°C): 166.5, 55.5, 51.9, 35.6. HR-ESI-MS: calcd. for [C14H28N6O8+H]+: 409.2047; found 409.2047 [M+H]+  N,N,N,N-Tetra-(O-benzyl-N-benzylacetohydroxamic acid)-diaminoethane, 9 Compound 6 (0.67 g, 2.00 mmol, 5 eq.) was dissolved in acetonitrile (15 mL) and potassium carbonate (0.28 g, 2.00 mmol, 5 eq.) and ethylenediamine (26.7 μL, 0.40 mmol) were added to the solution. After 16 h stirring at room temperature, the solution was dried under reduced pressure and the residue purified by semi-prep RP HPLC (10 mL/ min, gradient A: 0.1% TFA in deionized water, B: acetonitrile, A: 50 % to B: 100 % for 20 min., tR= 13.8 min) to yield product 9 as a colorless solid (33.5 %, 0.14 g). 1H NMR (400 MHz, CDCl3, 25°C): 7.28 (m, 40H), 4.75 (s, 16H), 4.20 (s, 8H), 3.23 (s, 4H). 13C NMR (101 MHz, CDCl3, 25°C): 135.5, 133.8, 129.6, 129.2, 128.7, 128.6, 128.0, 77.2, 55.4, 50.6 and 50.1 HR-ESI-MS: calcd. for [C66H68N6O8+H]+: 1073.5177; found 1073.5181 [M+H]+  H4EDT(B)HA, 10 Compound 9 (0.14 g, 0.13 mmol) was dissolved in methanol (5 mL) and Pd(OH)2/C (20 w/w %, 0.067 g) was added to the solution. After charging the flask with a H2-filled balloon, the solution was stirred for 16 h at room temperature. The crude oil was purified by semi-prep RP HPLC (10 mL/ min, gradient A: 0.1% TFA in deionized water, B: acetonitrile, A: 95 % to B: 100 % for 25 min., tR= 20.8 min) as a colorless oil (20.9 %, 0.02 g). 1H NMR (400 MHz, CDCl3, 25°C): 7.33 (m, 20H), 4.76 (s, 8H), 4.26 (s, 8H), 3.37 (s, 4H). 13C NMR (101 MHz, CDCl3, 25°C): 168.4, 135, 8, 128.2, 127.4, 54.6, 51.8 HR-ESI-MS: calcd. for [C38H44N6O8+H]+: 713.3299; found 713.3299 [M+H]+  General procedure for metal complexations H4EDT(M)HA or H4EDT(B)HA was dissolved in 1 mL water at neutral pH, FeCl3  6H2O, ZnSO4  7 H2O or Cu(ClO4)2  6 H2O were added in a 1:1 ratio. Extraction with ethylacetate gave the following metal complexes, analyzed by HR-ESI-MS: Fe-EDT(M)HA calcd. for [C14H25FeN6O8+H]+: 462.1162; found 462.1162 [M+H]+ ; Cu-EDT(M)HA calcd. for [C14H26CuN6O8+H]+: 470.1186; found 470.1186 [M+H]+; Zn-EDT(M)HA calcd. for [C14H26N6O8Zn+H]+: 471.1182; found 471.1182 [M+H]+; Fe-EDT(B)HA calcd. for [C38H41FeN6O8+H]+: 766.2414; found 766.2442 [M+H]+ ; Cu-EDT(B)HA calcd. for [C38H42CuN6O8+H]+: 774.2438; found 774.2440 [M+H]+; Zn-EDT(B)HA calcd. for [C38H42N6O8Zn+H]+: 775.2434; found 775.2439 [M+H]+ 	 99		  DFT calculations Density functional theory (DFT) calculations were carried out using the Gaussian 09 Rev.D01 suite of Programs.25 The B3LYP hybrid functional26 with 6-31G* basis set27 (for C, H, O and N atoms) and the LANL2DZ effective-core pseudopotential28 (for Zr and Fe) was employed to simulate the ground state structures of both the ligands and their metal complexes. For all simulations, solvation effect of water (ε = 78.3553) was implemented using the polarizable continuum model (PCM) on Gaussian 09.29 Optimized structures were confirmed to be the minimum on the potential energy surface by vibrational frequency calculations.  Cell viability assay All three human cell lines were obtained from ATCC, American Type Culture Collection, Manassas, USA. The human colon cancer cell line HT29 (HTB-38), the human breast cancer cell line MCF7 (HTB-22) and the human non-tumorigenic breast epithelial cell line (CRL-10317) were cultured in McCoy’s 5A (Invitrogen 1660082), DMEM (Invitrogen 11965) + 10% FBS or DMEM/F12 (Sigma Aldrich D6421) + Pen/Strep 1x + 2 mM glutamine + hEGF 0.01 µg/mL + hydrocortisone 0.5 µg/mL + human insulin 10 µg/mL, respectively, at 37°C in a humidified atmosphere of 95 % of air and 5 % CO2 respectively. For evaluation of growth inhibition, cells were seeded in 96-well plates (Corning, Fisher Scientific Co Ltd, Edmonton, Canada) at a concentration of 10000 cells/well (MCF7, MCF10A and HT29) and grown for 24 h in complete medium. Solutions of the compounds were prepared by diluting a freshly prepared stock solution (10-2 M in DMSO, or distilled H2O for DFO) of the corresponding compound to water. Afterwards, the intermediate dilutions of the compounds were added to the wells (200 µL) to obtain a final concentration ranging from 0 to 200 µM, and the cells were incubated for 72 h. Following 72 h drug exposure, 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazoliumbromide (MTT) was added to the cells at a final concentration of 0.5 mg ml-1 incubated for 2 h, then the culture medium was removed and the violet formazan (artificial chromogenic precipitate of the reduction of tetrazolium salts by dehydrogenases and reductases) dissolved in DMSO. The optical density of each well (96-well plates) was quantified three times in tetraplicates at 550 nm using a multi-well plate reader, and the percentage of surviving cells was calculated from the ratio of absorbance of treated to untreated cells. The IC50 value was calculated as the concentration reducing the proliferation of the cells by 50 % and it is presented as a mean (± SD) of at least three independent experiments.  		6 References (1) Lossen, W. Justus Liebigs Ann. Chem. 1872, 161, 347-362. (2) Muri, E. M.; Nieto, M. J.; Sindelar, R. D.; Williamson, J. S. Curr. Med. Chem. 2002, 9, 1631-1653. (3) Petrik, M.; Zhai, C.; Haas, H.; Decristoforo, C. Clin. Transl. Imaging 2017, 5, 15-27. (4) Kakkar, R. In Hydroxamic Acids: A Unique Family of Chemicals with Multiple Biological Activities, Gupta, S. P., Ed.; Springer Berlin Heidelberg: Berlin, Heidelberg, 2013, pp 19-53. (5) Chaston, T. B.; Richardson, D. R. Am. J. Hematol. 2003, 73, 200-210. (6) Deri, M. A.; Zeglis, B. M.; Francesconi, L. C.; Lewis, J. S. Nucl. Med. Biol. 2013, 40, 3-14. (7) Van Dongen, G. A.; Huisman, M. C.; Boellaard, R.; Harry Hendrikse, N.; Windhorst, A. D.; Visser, G. W.; Molthoff, C. F.; Vugts, D. J. J. Nucl. Med. Mol. Imaging 2015, 59, 18-38. (8) Deri, M. A.; Ponnala, S.; Zeglis, B. M.; Pohl, G.; Dannenberg, J. J.; Lewis, J. S.; Francesconi, L. C. J. Med. Chem. 2014, 57, 4849-4860. (9) Boros, E.; Holland, J. P.; Kenton, N.; Rotile, N.; Caravan, P. Chem. Plus Chem. 2016, 81, 274-281. (10) Rousseau, J.; Zhang, Z.; Dias, G. M.; Zhang, C.; Colpo, N.; Benard, F.; Lin, K. S. Bioorg. Med. Chem. Lett. 2017, 27, 708-712. 	100		(11) Vugts, D. J.; Klaver, C.; Sewing, C.; Poot, A. J.; Adamzek, K.; Huegli, S.; Mari, C.; Visser, G. W.; Valverde, I. E.; Gasser, G.; Mindt, T. L.; van Dongen, G. A. Eur. J. Nucl. Med. Mol. Imaging 2017, 44, 286-295. (12) Pandya, D. N.; Bhatt, N.; Yuan, H.; Day, C. S.; Ehrmann, B. M.; Wright, M.; Bierbach, U.; Wadas, T. J. Chem. Sci. 2017, 8, 2309-2314. (13) Buchwalder, C.; Rodriguez-Rodriguez, C.; Schaffer, P.; Karagiozov, S. K.; Saatchi, K.; Hafeli, U. O. Dalton Trans. 2017, 46, 9654-9663. (14) Hart, J. R. In Ullmann's Encyclopedia of Industrial Chemistry; Wiley-VCH Verlag GmbH & Co. KGaA, 2000. (15) Carrasco, M. R.; Brown, R. T.; Serafimova, I. M.; Silva, O. J. Org. Chem. 2003, 68, 195-197. (16) Gudmundsdottir, A. V.; Paul, C. E.; Nitz, M. Carbohyd. Res. 2009, 344, 278-284. (17) Esteves, M. A.; Vaz, M. C. T.; Goncalves, M. L. S. S.; Farkas, E.; Santos, M. A. Dalton Trans. 1995, 2565-2573. (18) Grigg, R.; Rankovic, Z.; Thoroughgood, M. Tetrahedron 2000, 56, 8025-8032. (19) Higgins, F. S.; Magliocco, L. G.; Colthup, N. B. Appl. Spectrosc. 2006, 60, 279-287. (20) Shankar, S.; Reddy, J. P.; Rhim, J. W.; Kim, H. Y. Carbohydr. Polym. 2015, 117, 468-475. (21) Valle, P.; Timeus, F.; Piglione, M.; Rosso, P.; di Montezemolo, L. C.; Crescenzio, N.; Marranca, D.; Ramenghi, U. Pediatr. Hematol. Oncol. 1995, 12, 439-446. (22) Richon, V. M. Br. J. Cancer 2006, 95, S2-S6. (23) Hsieh-Ma, S. T.; Shi, T.; Reeder, J.; Ring, D. B. Clin. Immunol. Immunopath. 1996, 80, 185-193. (24) Potuckova, E.; Jansova, H.; Machacek, M.; Vavrova, A.; Haskova, P.; Tichotova, L.; Richardson, V.; Kalinowski, D. S.; Richardson, D. R.; Simunek, T. PloS one 2014, 9, e88754. (25) Frisch, M. J.; Trucks, G. W.; Schlegel, H. B.; Scuseria, G. E.; Robb, M. A.; Cheeseman, J. R.; Scalmani, G.; Barone, V.; Mennucci, B.; Petersson, G. A.; Nakatsuji, H.; Caricato, M.; Li, X.; Hratchian, H. P.; Izmaylov, A. F.; Bloino, J.; Zheng, G.; Sonnenberg, J. L.; Hada, M.; Ehara, M.; Toyota, K.; Fukuda, R.; Hasegawa, J.; Ishida, M.; Nakajima, T.; Honda, Y.; Kitao, O.; Nakai, H.; Vreven, T.; Montgomery Jr., J. A.; Peralta, J. E.; Ogliaro, F.; Bearpark, M. J.; Heyd, J.; Brothers, E. N.; Kudin, K. N.; Staroverov, V. N.; Kobayashi, R.; Normand, J.; Raghavachari, K.; Rendell, A. P.; Burant, J. C.; Iyengar, S. S.; Tomasi, J.; Cossi, M.; Rega, N.; Millam, N. J.; Klene, M.; Knox, J. E.; Cross, J. B.; Bakken, V.; Adamo, C.; Jaramillo, J.; Gomperts, R.; Stratmann, R. E.; Yazyev, O.; Austin, A. J.; Cammi, R.; Pomelli, C.; Ochterski, J. W.; Martin, R. L.; Morokuma, K.; Zakrzewski, V. G.; Voth, G. A.; Salvador, P.; Dannenberg, J. J.; Dapprich, S.; Daniels, A. D.; Farkas, Ö.; Foresman, J. B.; Ortiz, J. V.; Cioslowski, J.; Fox, D. J.; Gaussian, Inc.: Wallingford, CT, USA, 2009. (26) Becke, A. D. J. Chem. Phys. 1993, 98, 5648-5652. (27) Frisch, M. J.; Pople, J. A.; Binkley, J. S. J. Chem. Phys. 1984, 80, 3265-3269. (28) Hay, P. J.; Wadt, W. R. J. Chem. Phys. 1985, 82, 270-283. (29) Scalmani, G.; Frisch, M. J. J. Chem. Phys. 2010, 132, 114110. 		 	 101		 		Supporting Information Chapter A3:  Tetrahydroxamic Acid Bearing Ligands:  EDTA and DTPA analogues 											Sarah Spreckelmeyer,a,b Yang Caoa and Chris Orviga a Medicinal Inorganic Chemistry Group, Department of Chemistry, University of British Columbia, 2036 Main Mall, Vancouver, British Columbia, V6T 1Z1, Canada b Department of Pharmacokinetics, Toxicology and Targeting, Groningen Research Institute of Pharmacy, University of Groningen, Antonius Deusinglaan 1, Groningen 9713 AV, The Netherlands 			  	 	102		 Figure S 1. 2D-HSQC of H4EDT(M)HA (400 MHz, D2O, 25°C)   Figure  S 2. 1H NMR spectra of A: H4EDT(M)HA B: Fe-EDT(M)HA C: Cu-EDT(M)HA  (400 MHz, D2O, 25°C).   0123456789101112f2	(ppm)020406080100120140f1	(ppm){4.28,55.81} {3.55,52.65}{3.19,36.23}4.79	D2O4.79	D2O1.52.02.53.03.54.04.55.05.5f1 (ppm)ABC100020003000400080859095100wavenumber [cm-1]Transmittance [%]EDT(M)HAFe-EDT(M)HACu-EDT(M)HA	 	 103		Figure S 3.  Full IR spectra of H4EDT(M)HA, Fe-EDT(M)HA and Cu-EDT(M)HA.   Figure S 4.  iTLC chromatograms of free 89Zr4+ (black, all radioactivity at about 150 mm), DFO (green, all radioactivity at the origin) and EDT(M)HA (pink, same as control); mobile phase: EDTA pH 7.0.  0 100 200 3000100200300Distance eluted [mm]CountsCtrlDFOEDT(M)HAO-Zr2 bond length [A] O7 2.2269 O9 2.4204 O3 2.2378 O5 2.3933   O8 2.2045 O10 2.1334 	 	104		                 Figure S 5.  DFT optimized structure (B3LYP/6-31G* + LANL2DZ and PCM solvation (water) on Gaussian 09) and bond lengths of Zr-DTT(ME)HA.     O-Zr2 bond length [A] O51 2.2509 O70 2.2275 O3 2.271 O22 2.2294 O6 2.2019 O4 2.1374 	 	 105		  O53 2.199 O65 2.1914 O25 2.209 O6 2.1981 Figure S 6.  DFT optimized structure (B3LYP/6-31G* + LANL2DZ and PCM solvation (water) on Gaussian 09) and bond lenghs of Zr-DTT(MP)HA.  Figure S 7.  DFT optimized structure of bifunctional Zr-DTT(MP)HA (B3LYP/6-31G* + LANL2DZ and PCM solvation (water) on Gaussian 09).   Figure S 8.  13C NMR spectra of arms (4a, top) and (4b, bottom), 400 MHz, CDCl3, 25°C.  253035404550556065707580859095100105110115120125130135f1	(ppm)	 	106			Table S 1.  Conditions tried for adding the "arm" to diethylenetriamine. Attempt Arm Salt Solvent 1 4a 9 eq. K2CO3 Acetonitrile 2 4a 6 eq. NaHCO3 Acetone 3 4b 5 eq. K2CO3 Acetonitrile 4 4b  Acetone (+/- reflux) 5 4b 6 eq. K2CO3 DMF 																							 107	    Chapter A4  Overcoming the Limitations in Thrombosis Treatment:  A Bifunctional Chelator as Positron Emission Tomography-Imaging Probe for Detecting Blood Clots     Sarah Spreckelmeyer,a,b	Frank M. Lee;c Ed Pryzdialc and Chris Orviga a Medicinal Inorganic Chemistry Group, Department of Chemistry, University of British Columbia, 2036 Main Mall, Vancouver, British Columbia, V6T 1Z1, Canada b Department of Pharmacokinetics, Toxicology and Targeting, Groningen Research Institute of Pharmacy, University of Groningen, Antonius Deusinglaan 1, Groningen 9713 AV, The Netherlands c Centre for Blood Research, Life Science Centre, 2350 Health Sciences Mall, Vancouver, BC, V6T 1Z3, Canada       	108	1. Abstract  Nuclear imaging techniques such as Positron Emission Tomography (PET) are useful tools for the non-invasive detection of low concentrations of radiotracers in the body. We aimed to design a bifunctional chelator that contains the well-studied 68Ga chelator H2dedpa and a thiol reactive group for conjugation to the coagulation factor FXa in order to detect and localize blood clots to facilitate surgical removal. In total, four different approaches to obtain the bifunctional chelator were investigated, three approaches using maleimide derivatives for thiol-coupling and one approach using an acrylate derivate. The maleimide functional group obtained in 2 reactions was found to be unstable under basic conditions. Moreover when we tried to synthesize a cyclic maleimide group, ring closure of the maleimide did not occur presumably due to steric hindrance of the carboxylic acids from the picolinic acid moieties. Reaction 4 yielded a promising acrylate analogue of H2dedpa, which showed good reactivity with the thiol group of glutathione. Further experiments have to be conducted to confirm this reactivity and subsequently Ga3+ chelation, radiolabeling experiments with 68Ga3+, stability experiments in human serum as well as phospholipid binding experiments need to be performed to obtain a successful diagnostic for blood clots.                       	 109	2. Introduction Blood clots are a serious health risk due to their ability to block blood flow and cause heart attacks and stroke. These debilitating and life threatening events are the largest healthcare burden on the globe. Several pharmaceuticals for the treatment of blood clots, based on the physiological clot-dissolving protein tissue plasmin activator (tPA) have been developed; however, these FDA-approved drugs exceed the physiological concentration of tPA by many orders of magnitude, which leads to unfavorable systemic effects and haemorrhagic risk. Despite the development of several potential new drugs, which were tested in major clinical trials, advances have been disappointing and the trend has been toward surgical extraction of the clot. Thrombectomy also has limitations, such as bleeding and difficulties in identification of clot location.1  The hemostasis of blood has a complex mechanism for maintaining blood fluidity and conversion to insoluble gel in sites of vascular injury. Blood coagulation (Figure 1) and fibrinolysis are usually in equilibrium, constantly repairing trivial lesions in the body. Upon endothelium injury, loss of the endothelial layer activates platelets to change shape and adhesion properties to form the primary hemostatic plug.  Platelets also release key clotting proteins, such as Factor V, von Willebrand Factor (vWF) and fibrinogen to promote stronger adhesion of platelets to the site of injury.2 An additional flip-flop reaction that exposes negatively charged phospholipids to the outer membrane of platelets provides a surface for the generation of thrombin and fibrin. This mechanism is called secondary hemostasis, which is a carefully controlled proteolytic cascade that forms a clot. The coagulation cascade consists of an intrinsic and extrinsic pathway (not discussed in detail here, but shown for completeness in Figure 1) that both lead to activation of a key serine protease FXa. FXa cleaves prothrombin zymogen in two places, yielding its active form thrombin. Importantly, this process is facilitated by the prothrombinase complex that consists of FXa and FVa assembled on negatively charged phospholipid membranes in the presence of calcium ions. Thrombin then converts fibrinogen to fibrin, the building block of a haemostatic plug. In addition, thrombin activates more platelets with FXIII and consequently more FXa is synthesized locally at the side of injury (Figure 1). When the concentration of FXa surpasses the threshold of physiological anticoagulants, fibrin can generate a clot.3 Due to its key role in coagulation, FXa is a prominent drug target for therapeutic anticoagulants and its mechanism of action is well studied.4 Fibrinolysis is the counterpart of blood coagulation that breaks down the cross-linked fibrin by plasmin. Plasminogen is the zymogen of plasmin and is produced in the liver. It has an affinity for clots and is incorporated into them. Plasminogen is activated to plasmin by tissue plasminogen activator (tPA) or urokinase. tPA is released slowly into the blood stream by damaged endothelium. Additionally, plasmin stimulates the production of tPA and urokinase via a positive feedback mechanism.    Figure  1.  Pathways of the blood coagulation cascade (reproduced with permission from Haematology, 2nd edition (C.J.Pallister and M.S. Watson), © Scion Publishing Ltd.  	110	Early detection of blood clots, and particularly at low concentrations, is challenging with the imaging techniques available nowadays: Magnetic resonance imaging (MRI), ultrasound (US) and computed tomography (CT). These techniques only look at one part of the body at a time, thereby delaying subsequent treatment and increasing the risk for complications. Nuclear imaging techniques (SPECT, PET) have a high sensitivity for the detection of low concentrations of a radiotracer and single whole-body scans can be obtained. Conjugation of radiotracers to biomolecules, which target to the object to be imaged, is a widely established method.5 Applying this approach to the detection of blood clots will help to develop new thrombolytic agents and to pinpoint clots for thrombectomy, thus improving the safety and efficacy of the treatment. Investigation of radiolabeled proteins that bind to blood clotting proteins for tracking blood clots is therefore a worthy endeavour with a great deal of potential for life-saving developments.  Thus far, different tracers have been studied for the diagnosis of blood clots. Most prominent, peptides targeting fibrin or platelet receptors were investigated with 99mTc (e.g. 99mTc-HYNIC-CGPRPPC in Figure 2)6 or 111In for SPECT imaging. In the past 10 years, tracers for PET imaging such as 18F-Fluorodeoxyglucose (18FDG) and most recently 64Cu-fibrin-binding probe 8 (64Cu-FBP8) coupled to 1,4,7-triazacyclononanetriacetic acid (NOTA)7,8 were investigated to detect arterial thrombosis (Figure 2). In the search for a highly localized target other than fibrin, FXa is a promising target, since FXa highly accumulates at the site of clots at the side of endothelial injury due to anionic phospholipid-binding.    Figure  2. Radiotracers discussed in this work.  To our knowledge, there is no literature published on the synthesis of an imaging tracer linked to FXa. Our group has developed numerous ligands with the ability to chelate various radiometals for different purposes. H2dedpa-p-Bn-NH2 (Figure 2) is well studied as a 67Ga chelator for SPECT imaging; Ga3+ has another radioisotope, positron emitter 68Ga, which makes it suitable for PET imaging.  The aim of this study was to design a bifunctional chelator consisting of the 68Ga chelator H2dedpa with a thiol reactive group, in order to conjugate it to the coagulation factor FXa via a thiol functional group inserted on FXa as published by Pryzdial et al.9 for the detection and localization of blood clots to facilitate surgical removal. Thus, we present here the synthesis approaches of bifunctional H2dedpa that bears a thiol reactive functional group. Different reactions were considered, as summarized in Table 1. We selected to use reaction #5 and #8 from Table 1, since they are well known reactions in biochemistry as they can occur under physiological conditions, whereas the other reactions are only used in basic chemistry under non-physiological conditions. Furthermore the maleimide-linkers have been applied frequently and they are commercially available.10,11 The thiol reaction with the electron-withdrawing group (EWG) looks promising in the literature as well and is discussed below in more detail. ONONOHNONHH2NNOOHNOOOHSSNNHNTc 99mLLLL L[99mTc]-HYNIC-CGPRPPC (L= tricine/EDDA)6NNNOHOOOHOOHO(AGADON)2PepNNNOOHOOOHHOO[64Cu]-FBP864Cu64CuOHHOHHOOHOH18FHHOH[18F]-FDGNHHNH2NNNOHOHOOH2dedpa-p-Bn-NH2	 111	 Table  1.  Summarized thiol-reaction with R1-SH as thiol12 (EWG = electron withdrawing group, X= halide) # Functional group Product # Functional group Product 1   6   2      7   3   8   4   9   5           3. Results and Discussion 3.1 Synthesis  R XR SR1 RR S R1SR1RXO RSOR1SOOR SOOR SR1FFFFF FFSFFR1R NOOR NOO S R1R RS R1S SNR S SHNR R1 SEWG EWG S R1	112	 Figure  3.  Approaches for the linkage of H2dedpa to a thiol reactive funtional group.  Four different approaches to synthesize a thiol-reactive H2dedpa analogue were attempted, summarized in Figure 3. In the following, the rationale of the different approaches is described and the results are presented.  Approach 1 was aimed to conjugate the linker smPEG4 that contains a maleimide functional group to H2dedpa-p-Bn-NH2. The linker bears an activated N-hydroxysuccinimide (NHS) ester that can easily react with a primary amine to form an amide bond. The product was synthesized, the obtained structure was confirmed by ESI-MS and 1H NMR spectroscopy (Figure S 1) and further tested for maleimide reactivity (see DTNB assay below).  In approach 2 smTEGK, containing a maleimide group, was conjugated to H2dedpa-p-Bn-NCS. However this approach failed. No product formation was observed. The reason for that might have been lack of product formation, or loss of the product during purification processes that included extraction and HPLC. However, we could not further investigate this due to a limited amount of starting material. For approach 3, H2dedpa-p-Bn-NH2 was used to synthesize the maleimide functional group directly on the primary aromatic amine without a linker (Scheme 1).  H2dedpa-p-Bn-NH2 was synthesized using an established protocol13 (Scheme S1). First a model reaction was used to probe the reaction conditions, using glycine as starting material (Scheme 2). Given successful product formation of the model reaction, the same conditions were used for the reaction between H2dedpa-p-Bn-NH2 and maleic anhydride. The intermediate formation was successful, as indicated by ESI-MS and NMR spectroscopies (Figure S2, Figure S3 and Figure S4), but the ring closure to obtain a maleimide functional group did not occur, presumably because of steric hindrance of the carboxylic acids from the picolinic acid moieties. Two approaches were carried out to solve this problem, refluxing the reaction mixture in H2O, and adding concentrated H2SO4 to protonate the carbonyl group as a catalyst, but neither approaches resulted in ring closure (Scheme 1).  NH HNNOOHNH2NOOHOON OONHNOOOO OO OONHNOOOOOO ONH HNNOOHNHNOOHNH HNNOOHNCSNOOHONHNOOOO OOOHNOHOH2NONHNOOOOOOONHOHONHNH HNNOOHNHNOOHSNH HNNOOHNNOOHOON NNOOHNHNOOHO#1#2#3#4NH HNNOOHNH2NOOHN NNOOHNH2NOOHClOOO ODMAP, DIPEAdry DMSOdry DMSOAcOHH2O, acetoneH2dedpa-p-Bn-NH2 smPEG4H2dedpa-p-Bn-NCS smTEGKH2dedpa-p-Bn-NH2 maleic anhydride	 113	   Scheme 1.  Synthesis of H2dedpa-p-Bn-maleimide #3    Scheme 2 . Model reaction for ring closure for maleimide synthesis.  With approach 4 we succeeded to get promising results. In Scheme 3, the synthesis route of H2dedpa-p-Bn-acrylate is illustrated. This approach was based on the synthesis of intermediate f (Scheme S1) as published by Boros et al. 13 who evaluated the differences on 67Ga radiochemical yield as well as apo-transferrin binding stability between H2RGD-1 and H2RGD-2 and found high radiochemical yields (RCYs) with 67Ga with both ligands. These ligands bear the cyclic RGD peptide either on the ethylenediamine backbone or on the secondary nitrogens, respectively.13 It was found, that for both ligands, quantitative labeling could be achieved, with [67Ga(RGD-1)]+ being more stable after 2h (92 %) than [67Ga(RGD-2)]+ (72%) assessed in transferrin stability experiments.  Here, the nitro functional group of intermediate f was reduced with Pd/C 20% loading and hydrogen for 2 h to yield compound 1 (Scheme 2). The N-benzyl groups stayed intact due to the short reaction time and less Pd/C loading compared to the established protocol. The N-benzyl protection groups were kept intact in order to prevent side-reactions with acryloyl chloride on the secondary amines. After that, the amine functional group was functionalized with acryloyl chloride to an acrylate 2. Deprotection of the methyl esters of the picolinic acids yielded product 3. NHHNH2NNNOHOHOONHHNNHNNOHOHOOOOHONHHNNNNOHOHOOOOmaleic anhydrideH2O refluxAcOHH2dedpa-p-Bn-NH2 H2dedpa-p-Bn-maleimideNH2NHOOHONOOH2O refluxAcOH, argonOOOOHOOHOOHO	114	  Scheme 3.  Synthesis of H2dedpa-N,N'dibenzyl acrylate #4  The product 3 was analyzed by 1H NMR (Figure 4), 13C NMR (Figure S5), 2D-HSQC (Figure S6) spectroscopy and HR ESI-MS. Concerning the 1H NMR spectrum, double bond protons usually show chemical shifts between 4-7 ppm; here, we observed three peaks in that area (6.44-6.39 ppm, 6.23 ppm and 5.80 ppm), each integrating for one proton (Figure 4). These peaks represent the acrylate double bond. In the 13C NMR spectrum (Figure S5), alkenes usually have a chemical shift between 115-140 ppm. Here, 127.1 ppm, 128.7 ppm and 129.1 ppm are the chemical shifts for the acrylate alkene protons. This is due to the negative inductive effect of the carbonyl group. The other peaks are difficult to assign, but the integrations of the hydrogens correspond with the theoretical number of hydrogens.  NNO2NNNOOOONNH2NNNOOOONNHNNNOOOOONNHNNNOHOHOOOPd/C, H2MeOHClOK2CO3,H2O : acetone (1:4)LiOHTHF : H2O (1:3)H2dedpa-N,N-dibenzyl-acrylate, 3f 1 2	 115	  Figure  4.  1H NMR spectrum of compound 3 (400MHz, MeOD, 25°C).  3.1.1. Thiol bioconjugation of compound 3 In the next step, a model reaction was designed to test the thiol bioconjugation reaction ability to the acrylate functional group of compound 3. Glutathione a tripeptide of glutamate, cysteine and glycine was chosen as a thiol-containing protein analogue, due to its structural similarity to proteins. The reaction scheme is given in Figure 5. As starting material, the tri-acrylate functionalized H2dedpa 4 was used that was synthesized from a by-product during the synthesis of intermediate 2. The product 5 precipitated due to its insolubility in hexylamine. Furthermore, during purification with HPLC, the picolinic esters appeared to be cleaved by trifluoroacetic acid (TFA), as suggested by ESI-MS. The product was analyzed by ESI-MS and 1H NMR (Figure 6, where differences between the starting material and the final product are indicated by arrows), 13C NMR and 2D HSQC spectroscopy. The 1H NMR spectrum clearly showed the disappearance of the alkene signals and appearance of additional peaks in the alkyl region for the glutathione protons. The integrations fit to the theoretical values of the number of protons.  To test the reactivity of the acrylate group of compound 3, the compound was incubated with bovine serum albumin (BSA), a thiol-containing protein, and the remaining thiol groups were analysed using DTNB using the same protocol for assessing the reactivity of attempt 1 (see below). Unfortunately, the BSA precipitated in the solvent of the model reaction, making the measurement of the absorbance of DTNB impossible. In the future, the addition of a water/methanol mixture may help to dissolve the BSA. Alternative methods should be developed to validate the reactivity of the acrylate groups with thiol functional groups. 2.53.03.54.04.55.05.56.06.57.07.58.08.59.0f1	(ppm)3.31	CD3OD	116	  Figure  5.   Model reaction for approach 4.   Figure  6.  1H NMR spectra of model reaction of glutathione with #4; starting material (top), product (bottom) (400 MHz, MeOD, 25°C); arrows indicate the differences between the two compounds changes.  3.2 DTNB Assay The product of attempt 1 was further characterized by determining the maleimide reactivity of H2dedpa-smPEG4 compared to the positive control smTEGK (Figure 7). BSA was used as a thiol-containing probe to conjugate to the maleimide functional group of H2dedpa-smPEG4. Unreacted thiol groups of BSA then reacted with 5,5′-dithiobis(2-nitrobenzoic acid) (DTNB) to form a colored compound with absorbance measurable at 412 nm. Thus, a high absorbance indicates a high concentration of unreacted thiols. As seen in Figure 8A, the absorbance decreases with NNNHNNOOOOOOONNHNNNOHOHOOSOOOHNOHNCOOHOCOOHH2NSNHOHNHOOCOHOOCNH2SHNONHCOOHOCOOHH2Nhexylamine/H2O 4:1SHHNOHNCOOHOCOOHH2N45glutathione2.42.83.23.64.04.44.85.25.66.06.46.87.27.68.08.4f1	(ppm)3.31	CD3OD3.31	CD3ODH2OH2O	 117	increasing the concentration of the positive control smTEGK until about 75 µM smTEGK, where the concentration of smTEGK is equal to the concentration of the free thiol groups on BSA. When iodoacetamide (IAA), which also reacts with free thiol groups, was added at the highest concentration of smTEGK, no further decrease was observed indicating that all reactive thiols of BSA have been coupled to smTEGK. Figure 8B shows that the absorbance did not decrease with increasing concentration of H2dedpa-smPEG4. This result suggests that the maleimide functional group on H2dedpa-smPEG4 is not functional. This could be due to the presence of impurities or instability of the maleimide functional group. From these results we conclude, that either the synthesis was not successful or the product is not stable under the applied conditions.   Figure  7.  Structure of positive control smTEGK used in the DTNB assay.    Figure  8.  DTNB results of approach 1 (A: positive control, B: H2dedpa-smPEG4). 4 Conclusions Several approaches were conducted to synthesize a H2dedpa derivative that contains a thiol reactive functional group for functionalization with FXa for detecting blood clots in small concentrations. Unfortunately, the use of the well-studied thiol-reactive maleimide functional group was not successful. However, the synthesis of a H2dedpa-acrylate derivative was achieved and characterized by 1H NMR, 13C NMR and 2D NMR spectroscopies. In addition, a model reaction with the thiol containing glutathione showed promising results concerning thiol-reactivity. In the near future, functionalization with FXa needs to be performed as well as in vitro phospholipid binding studies. 5. Experimental Materials and Methods 1H and 13C nuclear magnetic resonance (NMR) spectra were recorded on Bruker AV400, instrument at ambient temperature; the NMR spectrawere expressed on the δ scale and referenced to residual solvent peaks. Electrospray ionization mass spectrometry (ESI-MS) spectra were recorded on a Micromass LCT instrument also at the Department of Chemistry, University of British Columbia. High-performance liquid chromatography (HPLC) analysis of cold ONHNOOOOOOOHNOHOH2NsmTEGK0 25 50 75 100 125 1500.000.050.100.150.200.25smTEGK [µM]AbsorbanceA+ IAA0 25 50 75 100 125 1500.000.050.100.150.200.25[mTEGdedpa] (µM)AbsorbanceB+ IAA	118	compounds was done on a Phenomenex Synergi 4-μm Hydro-RP 80A column (250 mm x 21.2 mm) on a Waters WE 600 HPLC system equipped with a 2478 dual-wavelength absorbance UV detector run using the Empower software package. Common starting materials such as 6-bromomethylpyridine-2-carboxylic acid methyl ester14, H2dedpa-p-Bn-NH2 and H2dedpa-p-Bn-NCS were synthesized according to the literature.13 Approach 1: H2dedpa-p-Bn-smPEG4 H2dedpa-p-Bn-NH213 (0.008 g, 18.9 mmol, 1.9 eq.) was dissolved in dry DMSO (1 mL) and smPEG4 (0.005 g, 9.9 mmol), N,N-diisopropylethylamine (DIPEA) (0.005 mL, 29.9 mmol, 3 eq) and 4-dimethylaminopyridine (DMAP) (0.003, 29.9 mmol, 3 eq) were added. After 2 h stirring the reaction mixture at room temperature, the crude was purified via semi-prep reverse-phase HPLC (10mL/ min, gradient A: 0.1% TFA in deionized water, B: acetonitrile, A: 95% to B: 100% for 25 min) to obtain the product as a yellow oil (0.005 g, 74.6 %). 1H NMR (D2O, 400MHz) = 8.00-7.98 (d, 2H), 7.93-7.82 (m, 2H), 7.71-7.70 (d, 1H), 7.41-7.39 (d, 2H), 6.96-6.94 (d, 1H), 6.87-6.86 (d, 1H), 6.72-7.71 (d, 1H), 4.22-4.06 (m, 4H), 3.66-6.59 (m, 9H), 3.19 (s, 4H), 3.02 (s, 4H), 2.90 (s. 2H), 2.86 (s. 2H), 2.65 (s, 2H), 2.55 (s, 2H). ESI-MS calcd. for [C41H51N7O12+H]+: 834.3674; found 834.5 [M+H]+  Approach 2: H2dedpa-p-Bn-smTEGK H2dedpa-p-Bn-NCS13 (0.003 g, 7.2 mmol, 1.2 eq) was dissolved in dry DMSO (1 mL). mTEGLys (0.003 g, 6.0 mmol) was added to the solution and the reaction mixture was stirred for 16 h. The crude was purified via semi-preparative reverse-phase HPLC (10mL/ min, gradient A: 0.1% TFA in deionized water, B: acetonitrile, A: 95% to B: 100% for 25 min). No product formation was observed. Approach 3: H2dedpa-p-Bn-maleimide Model reaction15 Glycine (0.10 g, 1.33 mmol) was dissolved in glacial acetic acid (10 mL) and maleic anhydride (0.14 g, 1.47 mmol, 1.1 eq) was added under argon atmosphere. After confirmation of the intermediate by ESI-MS, the solvent was removed in vacuo and dist. water (5 mL) was added to the crude solid. After heating the reaction mixture to 60°C for 10 min, the suspension became clear and was refluxed overnight. The product was isolated as a white precipitate after cooling the solution down (0.19 g, 81.5 %). 1H NMR (DMSO-d6, 400MHz) = 6.06 (s, 2H), 3.68 (s, 2H). 13C NMR (DMSO-d6, 101MHz) = 169.1, 167.2, 135.4 and 39.9. ESI-MS calcd. for [C6H5NO4+H]+: 156.0297; found 156.1 [M+H]+ H2dedpa-p-Bn-maleimide H2dedpa-p-Bn-NH2  (0.007 g, 0.02 mmol) was dissolved in glacial acetic acid (2 mL) and maleic anhydride (0.002 g, 0.02 mmol, 1.1 eq), was added to the reaction mixture. After 16 h stirring of the reaction mixture at room temperature, the intermediate was confirmed via ESI-MS and NMR spectroscopy. The maleimide ring closure reaction (in 2 mL water, refluxing overnight) was unsuccessful.  1H NMR (MeOD, 400MHz) = 8.18-8.13 (m, 4H), 7.84-8.83 (d, 1H), 7.79-7.77 (d, 1H), 7.62-7.61 (d, 2H), 7.32-7.30 (d, 2H), 6.58-6.55 (d, 1H), 6.34-6.31 (d, 1H), 4.76-4.09 (dd, 4H), 3.79-3.69 (m, 2H), 3.59-3.48 (m, 3H), 2.98-2.97 (m, 1H). 13C NMR (MeOD, 101MHz) = 167.0, 164.6, 146.0, 139.6, 133.1, 130.9, 129.7, 126.7, 125.1, 120.9, 58.2, 56.2, 52.5 and 34.3. ESI-MS calcd. for [C27H27N5O7+H]+: 534.1989; found 534.2 [M+H]+ Approach 4: H2dedpa-p-Bn-acrylate Dimethyl 6,6'-(((3-(4-aminophenyl)propane-1,2-diyl)bis(benzylazanediyl))bis(methylene))dipicolinate, 1 	 119	Compound f (Scheme S1) (0.182 g, 0.27 mmol) was dissolved in methanol (10 mL). After adding Pd/C (0.05 g, 0.41 mmol, 1.5 eq) to the solution, the three-neck flask was charged with a H2-balloon and the reaction mixture was stirred for 2h until ESI-MS showed the reduction product 1. After that, the crude was purified via semi-preparative reverse-phase HPLC (10mL/ min, gradient A: 0.1% TFA in deionized water, B: acetonitrile, A: 95% to B: 100% for 25 min) to yield the product as an off-white solid (0.054 g, 31 %).  1H NMR (MeOD, 400MHz) = 8.04 (d, 2H), 8.00-7.92 (m, 16H), 7.00 (d, 2H), 2.98 (s, 6H), 3.94 (s, 4H), 3.65-3.39 (m, 4H), 3.20-2.96 (m, 3H), 2.60-2.55 (m, 2H). ESI-MS calcd. for [C39H41N5O4+H]+: 644.3237; found 644.3 [M+H]+ Dimethyl 6,6'-(((3-(4-acrylamidophenyl)propane-1,2-diyl)bis(benzylazanediyl))bis(methylene))dipicolinate, 2 This synthesis was modified from the literature.16 Potassium carbonate (0.019 g, 0.13 mmol, 1.2 eq) was placed in a three-neck flask, which was evacuated and flushed with argon.  Water  (1 mL) and acetone (4 mL) were added to the reaction flask and cooled to 0°C. A solution of compound 1 (0.072 g, 0,11 mmol) in 1 mL acetone was added to the suspension, followed by a slow addition of acryloyl chloride (10.8 µL, 0.13 mmol, 1.2 eq). The reaction mixture was stirred for 2h to yield a yellow oil (0.077 g, 0.11 mmol, 99 %). The solvents were removed in vacuo and the product 2 was used without further purification since the dominant peak in the ESI-MS was the product peak and HPLC purification was used after the final step. ESI-MS calcd. for [C42H43N5O5+Na]+ 698.3; found: 698.4 [M+H]+ H2dedpa-p-Bn-acrylate, 3 To a solution of compound 2 (0.08 g, 0.11 mmol) in a mixture of THF (7.5 mL) and water (2.5 mL) (3:1), lithium hydroxide (0.026 g, 1.1 mmol, 10 eq.) was added and the reaction mixture was stirred for 6 h at room temperature. The solution was neutralized with 0.1 M HCl and the solvents were removed in vacuo. Compound 3 was obtained as a yellow oil (0.006 g, 0.11 mmol, 8 %) after purification with semi-prep reverse-phase HPLC (10mL/ min, gradient A: 0.1% TFA in deionized water, B: acetonitrile, A: 95% to B: 100% for 25 min) (tR= 18.8 min). 1H NMR (MeOD, 400MHz) = 8.41-6.97 (m, 20H), 6.44-6.39 (m, 1H), 6.23 (d, 1H), 5.80 (d, 1H), 4.72-4.45 (m, 4H), 4.01-3.58 (m, 4H), 3.24-2.70 (m, 5H). 13C NMR (MeOD, 101MHz) = 166.1, 144.6, 144.1, 123.2, 131.1, 129.6, 128.0, 127.9, 122.0, 63.6, 58.3, 53.8, 53.2 and 34.8. HR-ESI-MS calcd. for [C40H39N5O5+H]+: 670.3029; found 670.3022 [M+H]+ Model reaction (glutathione) Dimethyl 6,6'-(((3-(4-acrylamidophenyl)propane-1,2-diyl)bis(acryloylazanediyl))bis(methylene))dipicolinate Potassium carbonate (0.046 g, 0.33 mmol, 1.2 eq) was placed into a flask, which was evacuated and flushed with argon.  Water (1 mL) and acetone (4 mL) were added to the reaction flask and the mixture was cooled to 0°C. A solution of intermediate f (0.129 g, 0.28 mmol) in acetone (1 mL) was added to the suspension, followed by a slow addition of acryloyl chloride (27.1 µL, 0.33 mmol, 1.2 eq.) The reaction mixture was stirred for 2h to yield a yellow oil (0.077 g, 0.11 mmol, 99 %). The solvents were removed in vacuo and the product was purified via semi-prep reverse-phase HPLC (10mL/ min, gradient A: 0.1% TFA in deionized water, B: acetonitrile, A: 95% to B: 100% for 25 min., tR =19.12 min) to yield the product as a white solid (0.07 g, 42%).  1H NMR (MeOD, 400MHz) = 8.04-7.88 (m, 3H), 7.77-7.70 (m, 1H), 7.48-7.39 (m, 3H), 7.29-7.20 (dd, 1H), 7.07-7.00 (m, 2H), 6.81-6.02 (m, 6H), 5.78-5.59 (m, 3H), 4.80-4.57 (m, 4H), 3.98-3.90 (m, 6H), 3.68-3.64 (d, 1H), 2.98-2.92 (m, 2H), 2.66-2.60 (s, 2H). 13C NMR (MeOD, 100MHz) = 165.3, 164.6, 139.3, 139.3, 130.8, 128.1, 122.8, 119.9, 53.7, 51.5, 47.9, 39.0, 35.1. ESI-MS calcd. for [C34H35N5O7+Na]+: 648.2434; found 648.3 [M+H]+ H2dedpa-triglutathione A modified protocol was used.17,18 The starting material 4 (0.07 g, 0.11 mmol) was dissolved in 1 mL hexylamine and 200 µL of methanol were added until complete dissolution. Glutathione (0.11 g, 0.37 mmol, 3.3 eq.) was added to the 	120	solution and methanol was added again until complete dissolution. After 2h stirring of the reaction mixture, a precipitate was formed. The precipitate was filtered off, dried and purified via semi-prep reverse-phase HPLC (10mL/ min, gradient A: 0.1% TFA in deionized water, B: acetonitrile, A: 95% to B: 100% for 25 min., tR =16.86 min) to yield the product 5 as a white solid (0.02 g, 11 %).  1H NMR (MeOD, 400MHz) = 8.03-7.82 (m, 4H), 7.48-7.33 (4H), 7.10-6.89 (m, 2H), 4.73-4.45 (m, 4H), 4.00-3.92 (m, 5H), 3.46-3.37 (m, 5H), 3.29-3.22 (m, 3H), 3.13-2.59 (m, 35H) 13C NMR (MeOD, 100MHz) = 169.2, 138.5, 127.8, 120.2, 120.7, 51.3, 40.9, 38.4, 31.3, 30.8, 29.2. ESI-MS calcd. for [C59H77N14O25S3+H]+: 1479.5260; found 1479.8 [M+H]+ DTNB assay The reactivity of the H2dedpa functionalized compound with the free thiol on bovine serum albumin (BSA) was assessed. BSA (50 μM) was incubated for 15 minutes with different concentrations of the synthesized compound (0, 25, 50, 75, 100 and 150 µM) using a 1 mM stock solution in DMSO of the compound in Tris buffer in a total volume of 100 µL. As a positive control, Lys-PEG4-maleimide was used under the same conditions. Lys-PEG4-maleimide bears a thiol reactive maleimide functional group. Remaining free thiol on BSA was quantified through the addition of 1 mM 5,5′-dithiobis(2-nitrobenzoic acid) (Ellmans’s reagent, DTNB) which reacts with free sulfhydryl groups to yield a mixed disulfide and 2-nitro-5-thiobenzoic acid (NTB), a measurable yellow-colored product detectable at 412 nm. Iodoacetamid (IAA) was used as a negative control that alkylates free thiols. 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L.; Rietz, T. A.; Rotile, N. J.; Day, H.; Looby, R. J.; Ay, I.; Caravan, P. J. Nucl. Med. 2014, 55, 1157-1163. (9) Pryzdial, E. L.; Meixner, S. C.; Talbot, K.; Eltringham-Smith, L. J.; Baylis, J. R.; Lee, F. M.; Kastrup, C. J.; Sheffield, W. P. J. Thromb. Haemost. 2016, 14, 1844-1854. (10) Northrop, B. H.; Frayne, S. H.; Choudhary, U. Polym. Chem. 2015, 6, 3415-3430. (11) Fontaine, S. D.; Reid, R.; Robinson, L.; Ashley, G. W.; Santi, D. V. Bioconjug. Chem. 2015, 26, 145-152. (12) Stenzel, M. H. Macro. Letters 2013, 2, 14-18. (13) Boros, E.; Ferreira, C. L.; Yapp, D. T.; Gill, R. K.; Price, E. W.; Adam, M. J.; Orvig, C. Nucl. Med. Biol. 2012, 39, 785-794. (14) Zeng, X.; Coquiere, D.; Alenda, A.; Garrier, E.; Prange, T.; Li, Y.; Reinaud, O.; Jabin, I. Chemistry 2006, 12, 6393-6402. (15) Song, H. Y.; Ngai, M. H.; Song, Z. Y.; MacAry, P. A.; Hobley, J.; Lear, M. J. Org. Biomol. Chem. 2009, 7, 3400-3406. (16) Chanthamath, S.; Takaki, S.; Shibatomi, K.; Iwasa, S. Angew. Chem. Int. Ed. 2013, 52, 5818-5821. (17) Li, G.-Z.; Randev, R. K.; Soeriyadi, A. H.; Rees, G.; Boyer, C.; Tong, Z.; Davis, T. P.; Becer, C. R.; Haddleton, D. M. Polym. Chem. 2010, 1, 1196-1204. (18) Chan, J. W.; Hoyle, C. E.; Lowe, A. B.; Bowman, M. Macromolecules 2010, 43, 6381-6388. 			121	   Supporting Information Chapter A4 Overcoming the Limitations in Thrombosis Treatment:  A Bifunctional Chelator as Positron Emission Tomography-Imaging Probe for Detecting Blood Clots 						Sarah Spreckelmeyer,a,b Frank M. Lee;c Ed Pryzdialc and Chris Orviga a Medicinal Inorganic Chemistry Group, Department of Chemistry, University of British Columbia, 2036 Main Mall, Vancouver, British Columbia, V6T 1Z1, Canada b Department of Pharmacokinetics, Toxicology and Targeting, Groningen Research Institute of Pharmacy, University of Groningen, Antonius Deusinglaan 1, Groningen 9713 AV, The Netherlands c Centre for Blood Research, Life Science Centre, 2350 Health Sciences Mall, Vancouver, BC, V6T 1Z3, Canada 		122	 Figure  S 1. 1H NMR spectra of #1 (400 MHz, D2O (top) MeOD (bottom), 25°C).   Scheme S 1. Synthesis of H2dedpa-p-Bn-NH2. 4.79	D2O2.53.03.54.04.55.05.56.06.57.07.58.08.5f1 (ppm)4.79	D2OH2dedpa-p-Bn-NH2H2dedpa-p-Bn-smPEG4NH2OHONH2OHOO2NNH2OOO2NNH2NH2OO2NNH2NH2O2NH2SO4, HNO3 MeOH, sat. HCl (g) MeOH, NH3 (g)THF, 1M diboraneNHNHO2NNNO2NNNOOOONHHNH2NNNOOOONHHNH2NNNOHOHOOPd/C, H2AcOH1. benzaldehyde2. NaBH4EtOHpicolinic acidLiOHCH3CNTHF/H2O 3:1a b cdefgh		123	 Figure S 2.  1H NMR spectrum of #3 (400 MHz, MeOD, 25°C).  Figure S 3.  13C NMR spectrum of #3 (101 MHz, MeOD, 25°C). 2.83.03.23.43.63.84.04.24.44.64.85.05.25.45.65.86.06.26.46.66.87.07.27.47.67.88.08.28.4f1	(ppm)1.31.11.61.73.42.94.31.00.92.01.51.01.23.92.983.283.31	CD3OD3.353.483.523.553.593.733.873.896.317.327.60-100102030405060708090100110120130140150160170180190200210f1	(ppm)34.3352.4656.2158.21120.86125.08126.72129.74130.86133.05139.55146.00164.62167.00		124	 Figure S 4.  2D-HSQC spectrum of #3.    Figure  S 5.  13C NMR spectrum of compound 3 (101MHz, MeOD, 25°C).  2.02.53.03.54.04.55.05.56.06.57.07.58.08.59.0f2 (ppm)30405060708090100110120130140f1 (ppm){8.05,139.52} {6.58,133.26}{6.32,130.44}{7.83,127.39}{8.18,125.02}{7.62,122.13}{4.03,60.52}{3.89,52.46}{3.74,48.86}{3.60,48.69}{4.97,48.58}{5.08,47.95}{4.76,46.73}{3.00,34.36}{3.52,33.91}30405060708090100110120130140150160170180190200210f1	(ppm)49.00	CD3OD		125	 Figure  S 6.  2D-NMR HSQC spectrum of compound 3.     2.53.03.54.04.55.05.56.06.57.07.58.08.59.0f2	(ppm)0102030405060708090100110120130140150f1	(ppm){8.41,142.82}{6.40,129.09}{7.31,129.09}{6.24,128.65}{5.77,127.13}{7.80,125.78}{8.29,122.85}{4.70,53.65}{3.89,52.26}{2.90,49.88}{2.86,34.40}   	126			Part B               	 127	   Chapter B1  Cellular Transport Mechanisms of Cytotoxic Metallodrugs: An Overview Beyond Cisplatin 							Sarah Spreckelmeyer,a,b Chris Orvig,b and Angela Casinia a Dept. Pharmacokinetics, Toxicology and Targeting, Research Institute of Pharmacy, University of Groningen, Antonius Deusinglaan 1, 9713 AV Groningen, The Netherlands; E-Mail: a.casini@rug.nl b Medicinal Inorganic Chemistry Group, Department of Chemistry, University of British Columbia, 2036 Main Mall, Vancouver, British Columbia V6T 1Z1, Canada.  Published in: Molecules, 2014, 19, 15584-15610.          	128	1 Abstract The field of medicinal inorganic chemistry has grown consistently during the past 50 years; however, metal-containing coordination compounds represent only a minor proportion of drugs currently on the market, indicating that research in this area has not yet been thoroughly realized. Although platinum-based drugs as cancer chemotherapeutic agents have been widely studied, exact knowledge of the mechanisms governing their accumulation in cells is still lacking, although evidence suggests active uptake and efflux mechanisms to be involved; this may be involved also in other experimental metal coordination and organometallic compounds with promising antitumor activities in vitro and in vivo, such as ruthenium and gold compounds. Such knowledge would be necessary to elucidate the balance between activity and toxicity profiles of metal compounds. In this review, we present an overview of the information available on the cellular accumulation of Pt compounds from in vitro, in vivo and clinical studies, as well as a summary of reports on the possible accumulation mechanisms for different families of experimental anticancer metal complexes (e.g. Ru Au and Ir). Finally, we discuss the need for rationalization of the investigational approaches available to study metallodrug cellular transport.      	 129	2 Introduction  Cisplatin [cis-diamminedichloroPt(II)] (Figure 1) is an important chemotherapeutic drug used in the therapy of a broad spectrum of human malignancies such as ovarian, testicular, head and neck, and lung cancers, and in combination with a wide range of other drugs for the treatment of other malignancies. For this reason, it is one of the most widely utilized antitumor drugs in the world, with annual sales of approximately $500 million (US). Unfortunately, its use is greatly limited by severe dose limiting side effects (nephrotoxicity, ototoxicity, and peripheral neurotoxicity) and intrinsic or acquired drug resistance. Thus, numerous Pt derivatives have been further developed with more or less success to minimize toxic effects. Over the last 30 years, 23 other Pt-based drugs have entered clinical trials with only two of these (carboplatin and oxaliplatin, Fig. 1) gaining international marketing approval, and another three (nedaplatin, lobaplatin and heptaplatin) approved in individual nations.1 Currently, there are only four Pt drugs in the various phases of clinical trial (satraplatin, picoplatin, LipoplatinTM and ProLindacTM).  Over the years, research on innovative anticancer metallodrugs has produced several ruthenium-based compounds as alternatives to Pt compounds. The Ru(III) complex trans-[tetrachloro(DMSO)(imidazole)ruthenate(III)] (NAMI-A) (Fig. 1) was demonstrated to have high selectivity for solid tumor metastases and low toxicity at pharmacologically active doses.2 This is the first ruthenium complex to enter clinical trials. A related Ru(III) compound, indazolium trans-[tetrachlorobis(1H-indazole)ruthenate(III)] (KP1019) and its sodium salt analogue NKP-1339 (Fig. 1), also entered clinical trials after they were found to exhibit cytotoxic activity in vitro in cisplatin-resistant human colon carcinoma cell lines and in vivo in various tumor types.3 Due to its higher water solubility, NKP-1339 has now been selected as a lead candidate for further clinical development.  Besides these coordination compounds, several classes of other metal complexes and organometallic compounds, based on different metals such as Au, Fe, Ag, Ga, Rh, and Ti, exhibit promising anticancer activity at least in preclinical studies.4-8 Noteworthy, Jaouen and co-workers have developed organometallic ferrocene-modified tamoxifens (named ferrocifens, Fig. 1) as estrogen targeting molecules effective in hormone-independent breast cancer cells, where hydroxytamoxifen and ferrocene are inactive.9 In this compound class the β-phenyl ring of tamoxifen has been substituted by a ferrocenyl moiety using classical organic/organometallic synthetic methods. Such structural modifications lead to more lipophilic compounds able to easily cross cell membranes, and, therefore, provided with stronger cytotoxic effects.   For most of all these cytotoxic metal-based compounds the mechanisms leading to their pharmacological and toxicological profiles are still not fully elucidated and different biological targets have been proposed, most of which still need validation. Due to the fact that DNA was identified early as the primary target of Pt(II) anticancer agents - adduct formation causes changes in DNA structure, hindering replication and transcription, which ultimately results in the induction of apoptosis – nucleic acids and their “alkylation” were believed to constitute the main pathway of activity for any cytotoxic metal compound.10      	130	 Figure 1.  Chemical structures of clinical and experimental metal-based anticancer agents.   Therefore, until 2006, only a limited number of biophysical studies encompassed the interactions of anticancer metallodrugs with proteins. These studies mostly concerned the two major serum proteins, albumin and transferrin, involved in the transport of metals and metallodrugs in the bloodstream, as well as metallothioneins, small, cysteine-rich intracellular proteins, primarily involved in storage and detoxification of soft metal ions. Afterwards, the crucial role of the interactions of metallodrugs with protein targets in determining the compounds’ pharmacological action, uptake and biodistribution, as well as their overall toxicity profile, was fully recognized and, as a result, the number of studies increased exponentially.11-13 Nowadays, cisplatin and other metal-based compounds are known to bind to several classes of proteins with different roles, including transporters, antioxidants, electron transfer proteins, DNA-repair proteins, as well as proteins/peptides simply used as model systems to characterize the reactivity of metallodrugs in vitro, but that are also present in vivo.13   Among the various families of investigated proteins, we believe that membrane transporters are of particular relevance. In fact, as essential mediators of specific cellular uptake, they are implicated not only in the pharmacological effects, but also in determining side-effects, metabolism, and excretion of many drugs, including cisplatin and other cytotoxic metal compounds.14 Recent progress has been made in understanding the role of membrane transporters in drug safety and efficacy. In particular, more than 400 membrane transporters organized in two major superfamilies — ATP-binding cassette (ABC) and solute carrier (SLC) — have been annotated in the human genome.15, 16 Many of these transporters have been cloned, characterized and localized to tissues and cellular membrane domains in the human body. In drug development, particular attention has been paid to transporters expressed in the epithelia of the intestine, liver and kidney, and in the endothelium of the blood–brain-barrier (BBB). As a result, a number of studies focus on the interaction of drugs and their metabolites with mammalian transporters present in epithelial and endothelial barriers. Interestingly, clinical pharmacokinetic drug–drug interaction (DDI) studies have suggested that transporters often work together with drug-metabolizing enzymes (DMEs) in drug absorption and elimination.  In spite of their great importance, transport mechanisms of anticancer metallodrugs have not yet been fully elucidated, especially in the case of a new generation of cytotoxic metal complexes. Moreover, the lack of a systematic investigation/approach to study metal compound accumulation in cells/tissues, as well as the generally limited structural information on membrane transporters, prevent the understanding of the complex mechanisms of drug absorption and excretion, particularly in medicinal inorganic chemistry. In this review we present a summary of the literature on the mechanisms of cellular uptake and accumulation for cisplatin, and analogues in the clinic. Most importantly, we will attempt an overview of the studies available for other families of experimental anticancer metal compounds, focusing on ruthenium and gold complexes (both coordination and organometallics) for which some PtCl NH3Cl NH3 PtOOOOH2CCH2H3NH3NH2NNH2PtOOOONHNRuCl ClCl ClSH3CH3CO HNHNNHNRuNNHCl ClCl ClNa+FeO(CH2)2N(CH3)2Cisplatin Carboplatin OxaliplatinNAMI-ANKP-1339Ferrocifen   	 131	studies are available in the literature. We have also considered the case of iridium-based organometallic complexes since recent detailed studies on their possible transport mechanisms have appeared. In the last section, the use of bioactive ligands to enhance the cellular uptake of metal compounds will be presented. 3 Transport processes of metal-based compounds  Initially, passive diffusion through the cellular lipid bilayer was considered to be the dominant process involved in drug uptake and distribution; however, recently the concept of carrier-mediated active uptake of commonly prescribed drugs has become the rule rather than the exception14. Thus, membrane transporters and channels, collectively termed the transportome,17 are increasingly recognized as important determinants of tumour cell chemosensitivity and chemoresistance. For example, reduction in Pt concentration of 20-70% has been observed in cancer cell lines resistant to cisplatin. This reduced accumulation can result from decreased influx or increased efflux, or both. Herein, we overview studies of cellular accumulation of Pt compounds, as well as of new anticancer metal complexes; thus, the few available studies reporting on the possible accumulation mechanisms for different families of experimental anticancer metal complexes (e.g. Ru, Au and Ir) will be summarized. Most importantly, we will attempt a rationalization of the investigational approaches available to study metallodrugs cellular transport and will comment on the relation between compound accumulation and anticancer properties.  3.1 Anticancer Pt drugs  Experimental evidence, well reviewed by Hall et al.,18 has led to the conclusion that cisplatin most likely enters the cell via two pathways: (a) passive diffusion and (b) facilitated and active uptake by a number of transport proteins.19 Membrane transporters of Pt-based anticancer agents determining active Pt uptake and efflux pathways, as well as their clinical significance have also recently been reviewed by Burger et al.20 including Cu transporters (Ctrs) organic cation transporters (OCTs), solute carriers (SLCs) and ATP-binding cassette (ABC) multidrug transporters. Other studies pointed also towards the involvement of different transport mechanisms in the overall biological activities of platinum compounds, including Na+-dependent glucose transport,21 and other ATP-dependent processes beside those regulated by Na+,K+-ATPase;22 however, no actual validation of such mechanisms have been attained so far. Our review presents a selection of in vitro, in vivo and clinical studies that elucidate mechanisms of accumulation for cisplatin and related Pt(II) anticancer drugs, together with information on the structural features and tissue distribution of the mentioned uptake and efflux transporters. When available, information of the reactivity of metal compounds with the protein transporters at a molecular level obtained by biochemical and biophysical methods will be provided.   3.1.1 Cu transporters   According to the Human Genome Organisation (HUGO), human transporters are classified based on their amino acid sequence in 43 solute carrier (SLC) families;23 Cu transporters have been assigned to the SLC31A family. Cu is an essential nutrient for almost all eukaryotic organisms to effect biological processes (e.g. free radical detoxification, mitochondrial respiration, iron metabolism, biosynthesis of neuroendocrine peptides etc.) and is cofactor in many enzymes. Due to the fact that the intracellular form of Cu, Cu+, is highly toxic because it reacts with molecular oxygen or hydrogen peroxide to produce free radicals, Cu homeostasis is guaranteed by a complex network of proteins that bind and deliver Cu+ to the Cu-dependent proteins and protect cells from the harmful effects of excess “free” Cu. Figure 2 is a schematic diagram of the Cu homeostasis system in mammals. Cu+ enters cells via the 23 KDa channel-like Cu transporter 1 (Ctr1) and is handed to pathway-specific chaperones such as antioxidant protein 1 (Atox1), Cu chaperone for superoxide dismutase (CCS), and cytochrome c oxidase assembly homolog (COX-17) that delivers it to various organelles for transfer to Cu-requiring enzymes. Afterwards, the P1B type ATPases ATP7A and ATP7B positioned in the trans-Golgi network secrete Cu. It is worth mentioning that the discovery of an existing protein homologue of Ctr1, namely Ctr2, presents another potential player in Cu homeostasis.24 Up to now the physiology and mechanism of Cu transport via Ctr2 has been poorly understood. In mammalian cells Ctr2 localizes to intracellular vesicular compartments including endosomes and lysosomes; however, when overexpressed via transfection with an epitope tag attached to the protein, Ctr2 has been localized at the plasma membrane, similarly to Ctr1.    	132	 Over several years, Cu transporters have been proposed to be involved in cellular import and export of Pt(II) chemotherapeutic agents, as well as in their resistance mechanisms.19, 25  In particular, expression of the human Cu transporter 1 (hCtr1) is thought to result in increased sensitivity to cisplatin, whereas expression of two Cu(I) proteins exporting ATPase, i.e. ATP7A and ATP7B, is believed to be involved in the resistance to cisplatin, either by sequestering drug away from its targets (ATP7A), or by exporting the drug from the cell (ATP7B).   Figure 2.  Schematic drawing of the major intracellular human Cu trafficking pathways.  The human hCtr1 (SLC31A1) is an evolutionarily conserved Cu influx transporter present in plants, yeast, and mammals, and the main Cu importer in mammalian cells. It is also a key player in the homeostatic regulation of intracellular Cu levels to ensure that nutritional delivery of Cu to enzymes, such as cytosolic Cu,Zn-superoxide dismutase (SOD).  hCtr1 is located in the plasma membrane and is constituted by three transmembrane helices, an extracellular N-terminal domain and a cytosolic C-terminal domain.26  Three hCtr1 molecules form a channel-like symmetric trimer, as revealed by electron microscopy.  hCtr1 contains two Met-rich motifs and two His-rich motifs on its extracellular N-terminus; both are thought to be essential for the function of the transporter.27 Interestingly, the Met-rich motifs located in the N-terminal domain and in the inner side of the channel pore are critical for the binding of Cu 28.   Studies (more than a decade old) revealed Ctr1 to be a significant pathway for the import of Pt cancer therapeutics into both yeast and mammalian cells. Thus, hCtr1 has been proven to play an essential role in the cytotoxic effects of Pt(II) drugs in cancer cells. For example, enhanced expression of the human Ctr1 gene in cancer cells normally resistant to cisplatin (including small cell lung cancers SCLC, and SR2 cells) led to an accumulation of cisplatin, carboplatin, and oxaliplatin, suggesting that hCtr1 can transport not only cisplatin but also carboplatin and oxaliplatin, albeit at reduced rates.27  In the same study, it was also shown that, although oxaliplatin transport was increased in cisplatin-resistant cells, the enhanced oxaliplatin accumulation failed to sensitize SR2 cisplatin-resistant cell lines, demonstrating that other intracellular pathways contribute to resistance mechanisms.   Always at a cellular level, studies of the pharmacology of cisplatin in human ovarian carcinoma A2780 cells, molecularly engineered to express increased hCtr1, showed that the overexpression of the transporter led to an increase in Pt accumulation and a decreased cell growth rate, yet it had limited effect on the sensitivity to cisplatin and to the amount of Pt binding to DNA.29 This discrepancy suggests that much of the Cu and cisplatin conducted into the cell must be sequestered away from the target through which it triggers cytotoxicity. Similarly, Beretta et al. reported that, whereas transfecting expression human hCtr1 cDNA into cisplatin-resistant epidermoid carcinoma A431 cells conferred increased uptake of Cu, no changes in cisplatin uptake and cellular sensitivity to the drug were observed.30 In parallel, the absence of the transporter renders cells resistant to cisplatin and carboplatin as demonstrated by Holzer et al. in ingenierized murine embryonic fibroblasts (Ctr1-/-).31 Moreover, these last results provide strong evidence that, at COX17Atox1Ctr1TGNmitochondrianucleusSOD1CuCu?ATP7A/B   	 133	concentrations < 1 µM, Ctr1 mediates cellular accumulation of all three clinically used Pt-containing drugs currently used in patients, but oxaliplatin differs from cisplatin and carboplatin in that its dependence on Ctr1 diminishes at higher concentrations. Overall, the idea that these drugs may have different influx transporters is consistent with their different spectra of action against various types of human cancer.   It must be mentioned that the interplay between Pt drugs, hCtr1 and Cu is more complicated than expected. As an example, another study by Holzer, Howell et al. showed that cisplatin rapidly triggers loss of endogenously and exogenously expressed hCtr1 in both human ovarian cancer cell lines A2780 and 2008, and that this effect has functional consequences for the uptake of Cu.32 Moreover, the same authors provided evidence that cisplatin-induced loss of hCtr1 in 2008 cells involves internalization from the plasma membrane by macropinocytosis followed by proteasomal degradation.33   An original study by Chen et al.34 using transfected cells displaying elevated levels of GSH, showed that such cells exhibited marked sensitivity to cisplatin due to up-regulation of hCtr1. Following these intriguing results, in 2012 Liang, Kuo et al demonstrated in multiple cell models that expression of hCtr1 in cisplatin resistant variants can be preferentially up-regulated by Cu-lowering agents (chelators) as compared with those in their drug-sensitive counterparts, providing greater sensitivity to the killing by Pt drugs.35 The parallel with the previous study by Chen34 lies in the fact that GSH is also an abundant physiologic Cu chelator. In that same study, enhanced cisplatin efficacy by a Cu-lowering agent was also observed in animal tumor xenografts bearing cisplatin resistant cells. Furthermore, analysis of a public dataset concluded35 that ovarian cancer patients with elevated expression levels of hCtr1 in their tumors had more favourable treatment outcomes after Pt-drug treatment than did those with low hCtr1 levels. Together, these findings provide a mechanistic basis for overcoming cisplatin resistance using Cu chelation strategy. Interestingly, the same authors conducted a pilot clinical study using the Cu-lowering agent trientine in combination with carboplatin in five Pt-resistant high-grade epithelial ovarian cancer patients.36 Encouraging results were obtained showing preliminary clinical evidence that the role of decreasing Cu levels in reversing Pt resistance merits additional clinical investigation.  Interestingly, patients with high levels of Ctr1 in their tumor appear to respond better to drug treatment.35 Notably, 22 single nucleotide polymorphisms (SNP) of Ctr1 have been identified by the screening of 282 non-small-cell lung carcinoma (NSCLC) Chinese patients.37 In particular, genetic polymorphisms of Ctr1 at rs7851395 and rs12686377 were associated with Pt resistance in NSCLC patients. Thus, these findings corroborate the idea that Ctr1 plays an essential role in Pt resistance and that it could be considered a predictive marker for the pretreatment evaluation of NSCLC patients.  Very recently, Kim et al.38 compared tumor Ctr1 expression with intratumoral Pt concentration in clinical specimens. In detail, these authors have hypothesized that a defect in tumor Ctr1 expression is associated with reduced tissue Pt accumulation and tumor response in NSCLC following Pt-based chemotherapy. The study showed that NSCLC patients with undetectable Ctr1 expression in their tumors had reduced intratumoral Pt concentration and tumor response compared to patients with any level of Ctr1 expression. Unfortunately, this study enrolled a limited number of patients and independent validation with a prospective clinical trial would be necessary.38  Concerning side-effects of Pt drugs, renal tubular damage is recognized as a major pathogenic factor in cisplatin nephrotoxicity. In fact, cisplatin is accumulated in renal tubular cells at high concentrations, leading to tubular injury and cell death. Recent research has revealed multiple signalling pathways that are responsible for tubular cell injury and death during cisplatin nephrotoxicity. In 2009 Pabla et al. demonstrated that Ctr1 is mainly expressed in both proximal and distal tubular cells in mouse kidneys.39 Importantly, down-regulation of Ctr1 in human embryonic kidney (HEK293) by small interfering RNA or Cu pre-treatment resulted in decreased cisplatin uptake;39 however, cimetidine, a substrate of organic cation transporters (OCTs), also had partial inhibitory effects on cisplatin uptake in HEK293 cells. Notably, it was also shown that cimetidine could further reduce cisplatin uptake in Ctr1 knockdown HEK293 cells, and both apoptosis and necrosis induced by cisplatin were further reduced by cimetidine in Ctr1 knockdown HEK293 cells.39 According to these results, it could be hypothesized that not only Ctr1 but also a cimetidine-inhibitable transport system, probably an organic cation transporter, contribute to cisplatin transport in renal tubular cells, resulting in nephrotoxicity. Nevertheless, it must be considered that cimetidine may also have off-target effects.  Interestingly, cisplatin treatment of a cell line expressing hCtr1 revealed the time- and concentration-dependent appearance of a stable hCtr1 multimeric complex, consistent with a homotrimer, that was not observed following Cu treatment of these same cells.40 Mutagenesis studies identified two methionine-rich clusters in the extracellular amino-   	134	terminal region of hCtr1 that were required for stabilization of the hCtr1 multimer by cisplatin, suggesting that these sequences bind cisplatin, and, subsequently, form crosslinks between hCtr1 polypeptides 40.   At a molecular level, Natile and co-workers investigated the binding of Pt complexes to the Met-rich domain of hCtr1 by different techniques including mass spectrometry and NMR spectroscopy;41 according to their findings cisplatin appears to easily form adducts with the peptide domain in which all the original ligands of Pt are lost and replaced by the S-donor Met groups. Based on these observations, cisplatin would be actually sequestered by hCtr1 and not transported, while a possible transport system could actually be an endocytotic process, incorporating a portion of the extracellular milieu (containing non-degraded cisplatin) into vesicles, which are subsequently delivered to subcellular compartments. These latter results might be in accordance with a more recent paper reporting on the fact that overexpression of hCtr1 in the human embryonic kidney HEK293 cell line did not result in increased sensitivity to cisplatin.42  Other studies based on MS and NMR spectroscopy characterized the binding of Pt compounds with synthetic peptides corresponding to hCtr1 Met-motifs at a molecular level, in some cases highlighting the differences among the various Pt drugs.43, 44 Finally, recent studies by Wang et al. based on NMR spectroscopy and electrospray ionization mass spectrometry (ESI-MS) show that a maximum of two Pt atoms is bound to each monomer unit of hCtr1 for cisplatin, carboplatin and nedaplatin.45  Once Pt enters the cells, although many different systems appear implicated in cisplatin trafficking, mounting evidence suggests a linkage between cisplatin resistance and the human Cu homeostatic proteins Atox1 and ATP7A or ATP7B.46 The Cu chaperone Atox1 binds Cu(I) at a conserved CXXC motif and delivers it to the N-terminal metal binding domains (MBDs) of ATP7B and ATP7A, which are Cu(I) specific P1B-type ATPases. Each human Cu(I) ATPase has six MBDs, which also bind Cu(I) with CXXC motifs and resemble Atox1 in the overall structure.47 The structure of a stoichiometric cisplatin-Atox1 adduct (Pt-Atox1) was determined at 1.6 A ̊ resolution showing a Pt(II) ion coordinated to Cys12 and Cys15 from the CXXC motif.48 The geometry is square planar with the two cysteine ligands oriented trans to one another. The remaining ligands are provided by the backbone amide nitrogen of Cys12 and a 2-carboxyethylphosphane (TCEP) molecule with a TCEP(P)-Pt distance of 2.48 A ̊. In the same paper the structure of a dimeric cisplatin adduct Pt-(Atox1)2 was also reported at 2.14 A ̊ resolution. Overall, the two structures support the idea that the cisplatin interaction with Cu(I) binding motifs may lead to unfavourable therapeutic outcomes, not only due to unproductive cisplatin trafficking, but perhaps also as a result of aberrant Cu(I) transport in cisplatin resistant tumours.  Several studies support the hypothesis that both ATP7A and ATP7B are involved in the resistance to cisplatin, either by sequestering drug away from its targets (ATP7A), or by exporting the drug from the cell (ATP7B). In eukaryotes, Cu(I) ATPases both efflux excess Cu and shuttle Cu to the secretory pathway for incorporation into enzymes.47 Mutations in the human Cu(I) P1B ATPases, ATP7A and ATP7B, lead to the Cu metabolic disorders Menkes syndrome and Wilson disease, respectively.49 The Cu(I) ATPases consist of eight transmembrane helices, an ATP binding domain that comprises a nucleotide binding domain (N domain) and a phosphorylation domain (P domain) containing an invariant DKTGT sequence that becomes phosphorylated at the aspartate residue during the ATP hydrolysis cycle. An A-domain, a key link in coupling nucleotide hydrolysis to ion transport, is also present. ATP7A is expressed in many tissues, while ATP7B is mainly present in liver and brain. The concept that Cu exporters may mediate cisplatin resistance was introduced by Komatsu et al.,50 reporting cisplatin resistance in prostate carcinoma cells overexpressing ATP7B. Afterwards, several other studies, in cancer cells resistant to cisplatin, demonstrated that these cell lines overexpress at least one of the two efflux Cu transporters.51-54 Interestingly, ATP7B siRNA incorporated into the neutral nanoliposome 1,2-dioleoyl-sn-glycero-3-phosphatidylcholine was highly effective in reducing tumor growth in combination with cisplatin in two orthotopic mouse models of ovarian cancer (70-88% reduction in both models compared with controls).52 This reduction in tumor growth was accompanied by reduced proliferation, increased tumor cell apoptosis, and reduced angiogenesis.52  In vivo studies of ATP7B expression in nine human NSCLC xenografts using real-time polymerase chain reaction (PCR) and immunohistochemistry, showed that ATP7B mRNA expression was significantly correlated with cisplatin sensitivity.55 Moreover, ATP7B mRNA and protein expression levels in the cisplatin-resistant xenografts were significantly higher than those in the sensitive.55 These results suggest that ATP7B is a cisplatin-resistance marker in human NSCLC xenografts in vivo. Notably, a clinical study showed that ATP7B mRNA and protein expression in colorectal tumors is associated with clinical outcomes to oxaliplatin combination therapy with 5-fluorouracil.56  Using genomic analysis, it was found that ATP11B gene expression was substantially increased in cisplatin resistant cells.57 Moreover, ATP11B enhanced cisplatin efflux and ATP11B silencing restored sensitivity of ovarian    	 135	cancer cells to cisplatin. ATP11B is included in the sub-family 4 of P-type ATPase. These proteins are thought to translocate phospholipids, rather than cations, from the outer to the inner leaflet of membrane bilayers. One hypothesis is that this type of ATPase may be involved in the vesicular transport of cisplatin.  Finally, at a molecular level, it has been demonstrated that, similar to Cu, Pt binds the CXXC motives of the cytosolic N-terminal binding domain of ATP7B,58 and that such interaction mediates cancer cell resistance to cisplatin. Recently, it has been shown by solid supported membrane technique (SSM) that Pt drugs can activate Cu-ATPases in microsome samples and undergo ATP-dependent translocation in a fashion similar to Cu.59   3.1.2 Organic cation transporters (OCTs) and toxin extrusion proteins (MATEs)   The organic cation transporters have been assigned to the solute carrier SLC22A family consisting of three sub-categories based on the charge of the transporter: the electrogenic transporter (OCT1-3), electroneutral organic cation/carnitine transporter (OCTN1-3) and the organic anion transporter (OATs, and urate transporters, URAT-1).15 Each transporter of the SLC22A family consist of 12 α-helical transmembrane domains (TMDs), a large glycosylated extracellular loop between TMDs 1 and 2, and a large intracellular loop between TMDs 6 and 7 with consensus sequences for phosphorylation. Most endogenous or exogenous substrates for OCTs are charged positively at physiological pH 7.4, and the electrochemical gradient is the crucial force for the uptake of the cationic substrates. The transport of organic cations is also independent of Na+ and reversible with respect to direction. A disadvantage of OCTs is that they are polyspecific, meaning that through their large binding domain, which contains partially overlapping interaction domains, a wide catalogue of substrates can be transported.  Many transporters of the SLC22A family are located in secretory organs like liver and kidney, as well as intestine; therefore, they play a pivotal role in drug adsorption and excretion, and different OCTs show species- and tissue-specific distributions. For example, the human OCT1 is highly expressed in the sinusoidal membrane of the liver and in jejunum. Instead, human OCT2 is mainly expressed in the basolaterial side of renal proximal tubule cells, and in the dopaminergic brain regions. For the correct interpretation of translational studies, it is important to mention that in rodents, both OCT1 and OCT2 show a high renal expression in the basolateral membrane of proximal tubule cells, with higher OCT2 expression in male animals. hOCT3 shows a much broader tissue distribution, including skeletal muscle, heart, brain, and placenta, but the distribution in the membrane and physiological role of OCT3 are not yet clearly understood.   Investigating the mechanisms of cisplatin nephrotoxicity has evinced the role of organic cation transporters (OCTs) in cisplatin transport. Various studies demonstrating that cisplatin can be transported by OCTs in cells were based on competition experiments with other established OCTs substrates such as tetraethylammonium (TEA) and cimetidine among others. Ciarimboli has recently extensively reviewed these studies and the reader is referred to his review paper for details.60, 61 As a representative example, interaction of cisplatin with hOCT2 in kidney or hOCT1 in liver was investigated with the fluorescent cation 4-[4-(dimethyl-amino)styril]-methylpyridinium (ASP) in stably transfected HEK293 cells and for the first time in tissues physiologically expressing these transporters, human proximal tubules, and human hepatocyte couplets.62 Cisplatin inhibited ASP transport in hOCT2-HEK293 but not in hOCT1-HEK293. In human proximal tubules the drug competed with basolateral organic cation transport, whereas it had no effect in tubules from a diabetic kidney or in hepatocytes. In hOCT2-HEK293 cells, 15 h incubation with cisplatin induced apoptosis, which was completely suppressed by simultaneous incubation with the hOCT2 substrate cimetidine.62 These findings support the idea of the interaction of cisplatin with hOCT2 in renal proximal tubules, but not with hOCT1, explaining its organ-specific toxicity.  More recently, the functional effects of cisplatin treatment on kidney (24 h excretion of glucose, water, and protein) and hearing (auditory brainstem response) were studied in wild-type and OCT1/2 double-knockout mice.63 No sign of ototoxicity and only mild nephrotoxicity were observed after cisplatin treatment of knockout mice. Co-medication of wild-type mice with cisplatin and the organic cation cimetidine protected from ototoxicity and partly from nephrotoxicity.63 Moreover, it should be noted that in rats treatment with both cisplatin and cimetidine did not interfere with the antitumoral activity of the Pt drug.64 Based on these studies, among others, hOCT2 has been proposed as target for protective therapeutic interventions accompanying cisplatin treatment.  Concerning other anticancer Pt drugs, Yonezawa et al. demonstrated that also oxaliplatin’s toxicity and uptake are enhanced by hOCT2 expression and weakly by hOCT3 in HEK293 cells,65 while no effect was observed in the case of    	136	carboplatin and nedaplatin. Oxaliplatin, however, showed almost no influence on the TEA uptakes in the HEK293 cells expressing hOCT1, hOCT2, and hOCT3.65 Recently, hOCT3 appeared to be also involved in the transport of cisplatin because cisplatin-sensitive cervical adenocarcinoma KB-3-1 cells express much higher level of hOCT3 than their Pt(II) resistant variants;66 however, studies on hOCT3-overexpressing HEK293 cells showed no effect on cisplatin accumulation with respect to wild-type cells.65 Therefore, these results point to differences in Pt accumulation due to the selected cell type.  Concerning hOCT3, selective induction of hOCT3 mRNA expression in colon cancer and colorectal cancer-derived cell lines has been reported.67 Interestingly, in this study the cytotoxicity and accumulation of Pt caused by the treatment with oxaliplatin, but not cisplatin, depended on the expression of hOCT3 mRNA. Thus, the uptake of oxaliplatin into the cancer cells via hOCT3 was suggested to be an important mechanism for its cytotoxicity, and the expression of hOCT3 in cancers was proposed to become a marker for including oxaliplatin in cancer chemotherapy.  Beside the OCTs, also the multidrug and toxin extrusion proteins (MATEs) are part of organic cation homeostasis, and belong to the SLC47 family. Specifically, MATEs act as H+/organic cation antiporters, transporting protons from the extracellular side to the cytoplasm while organic cations are exported. Two isoforms are known, SLC47A1 (MATE1) and SLC47A2 (MATE2-K). MATE1 is primarily expressed in the liver and kidney, while MATE2-K exhibits a kidney-specific expression.68 In the liver, MATE1 is localized on the canalicular membrane of hepatocytes and appears to form a functional unit with the basolaterally expressed OCT1 to mediate the biliary excretion of cationic drugs and their metabolites across the hepatocytes. In the kidney, MATE1 is highly expressed on the luminal membrane of the proximal tubular cells and is thought to play a key role in the excretion of organic cations. In the proximal tubule epithelium MATE1 cooperates with the basolaterally expressed OCT2 in the renal secretion of organic cations. Substrates for MATE1 and MATE2-K are typical organic cations, TEA, 1-methyl-4-phenylpyridinium (MPP), metformin, cimetidine, procainamide among others. Some compounds were reported to be specific inhibitors of MATE, although OCT and MATE are common in substrate specificity.  The few reports dealing with Pt drug accumulation by MATEs, have been well summarized by Ciarimboli.69 As a representative example of the knowledge in this area we decided to discuss the previously mentioned paper by Yonezawa et al.65 In this study, when HEK293 cells, transiently expressing hMATE1 or hMATE2-K, were treated with 50 to 1000 μM cisplatin for 2 h, the expression of the transporters did not affect cisplatin-induced cytotoxicity. In addition, the transporter activities were confirmed by the uptake of [14C]TEA; nevertheless, the accumulation of cisplatin was enhanced by hMATE1 more than hMATE2-K. Conversely, the accumulation of oxaliplatin was enhanced by hMATE2-K more than hMATE1.65 Since oxaliplatin is only poorly nephrotoxic, the obtained results suggest that the basolateral hOCT2 is the influx transporter responsible of oxaliplatin-induced toxicity, while the apical hMATE1 and hMATE2-K are efflux transporters as a means to protect cells. Overall, transcellular transport and cellular toxicity of oxaliplatin should be further examined to validate such a hypothesis.  Yokoo et al.70 reported in vivo studies in rats where higher accumulation of cisplatin in rat kidney tissues was observed in comparison to that of either oxaliplatin or carboplatin. As expected, such higher accumulation of cisplatin led also to an increase in nephrotoxicity, which was proven by overexpression of biomarkers of kidney injury like osteopontin in kidney slices. Moreover, in vitro studies showed that rat MATE1 as well as human MATE1 and MATE2-K, stimulated the H+-gradient-dependent antiport of oxaliplatin, but not of cisplatin, carboplatin and nedaplatin.70   Finally, a recent study by Li et al.71 reported the effects of coadministering cisplatin with the antiemetic 5-hydroxytryptamine-2 (5-HT3) receptor antagonist ondansetron. The introduction of 5-HT-3 receptor antagonists has been a significant clinical advance in the prevention and treatment of chemotherapy-induced nausea and vomiting, particularly for patients receiving highly emetogenic cisplatin-based regimens. Interestingly, 5-HT3 receptor antagonists such as ondansetron can interact with OCTs and MATEs. Initially, the inhibitory potencies of ondansetron on metformin accumulation mediated by OCT2 and MATEs were determined in stable HEK293 cells expressing these transporters,71 showing that ondansetron is a potent MATE inhibitor and a mild OCT2 inhibitor. Furthermore, in vivo experiments showed that cisplatin caused much more severe nephrotoxicity in Mate1-/- mice in comparison to wild-type mice.71 Similarly, co-treatment with ondansentron and cisplatin in wild-type mice caused increased nephrotoxicity as evidenced by increased levels molecular biomarkers of kidney injury and by more severe pathohistological changes in kidney tissues.71    	 137	 Overall, these studies demonstrate that in humans the interplay between OCT2 and MATE may affect the net renal secretion of shared drug substrates, including Pt drugs, but further investigation will be necessary to elucidate fully the complex pathways of interaction. 3.2 Experimental anticancer metal compounds 3.2.1 Ruthenium complexes  Concerning the most studied coordination Ru(III) compounds, the previously mentioned KP1019 and NKP-1339 are administered intravenously and, therefore, their interactions with serum proteins are of great relevance. In fact, several studies have shown strong affinities for both compounds to proteins in the bloodstream, particularly serum albumin and transferrin.3 Accordingly, it has been suggested that these proteins act not only as ruthenium carriers and delivery systems, but are also essential for tumor targeting. While binding to albumin may contribute to the targeted delivery of ruthenium compounds to cancer tissues due to the phenomenon known as Enhanced Permeability and Retention (EPR) effect72 (due to the combination of leaky blood capillaries and lack of lymphatic drainage in tumors), binding to transferrin may constitute an “active” targeting route. In fact, the iron transport protein transferrin (Tf) is crucial in tumor development since highly proliferative tumors have higher demand for iron than normal tissues, resulting in the overexpression of the transferrin receptor (CD71).   Numerous studies73 describing the reactivity of KP1019 and NKP-1339 with Tf, as well as the accumulation of the compounds in cancer cells, support the hypothesis that selective delivery of these Ru(III) compounds occurs into the malignant tissue via Tf followed by cellular uptake via Tf receptors. The receptor-mediated incorporation of Tf results in the formation of endosomes having low pH (ca. 5.5) with respect to the physiological one, which is supposed to trigger the release of the ruthenium compounds inside the cells.74 Unfortunately, to the best of our knowledge, no actual validation of such uptake pathway has been reported so far for ruthenium compounds.   In 2005 Heffeter, Keppler et al. investigated whether the ABC family of drug transporters may lead to resistance against KP1019-induced cytotoxicity in vitro.3, 75 Thus, KP1019 was tested against a panel of chemosensitive cell lines and their chemoresistant sublines expressing defined resistance mechanisms; the results showed that the cytotoxic effects of KP1019 are not substantially hampered by overexpression of the drug resistance proteins multidrug resistance-related protein 1 (MDR1), breast cancer resistance protein (BCRP), and lung resistance protein (LRP) or the transferrin receptor, and only marginally by the cellular p53 status. In contrast, P-glycoprotein overexpression reduced KP1019 activity weakly but significantly (up to 2-fold). P-glycoprotein (P-gp)-related resistance was based on reduced intracellular KP1019 accumulation and was reversible by known P-glycoprotein modulators.  To analyze whether KP1019 directly interacts with P-gp, the impact on the P-glycoprotein ATPase activity was measured in the presence of ouabain, EGTA, and sodium azide to block the membrane-bound Na+/K+, Ca2+, and mitochondrial ATPases.75 Interestingly, KP1019 dose-dependently inhibited ATPase activity of P-gp. Furthermore, it potently blocked P-gp-mediated rhodamine 123 efflux under serum-free conditions (EC50, ∼8 μM), however, with reduced activity at increased serum concentrations (EC50 at 10% serum, ∼35 μM).  Many P-gp substrates also act as P-gp modulators and competitively inhibit the efflux of other substrate drugs.76 To test whether KP1019 was able to modulate P-gp-mediated resistance, the compound was administered to P-gp-overexpressing cells together with the two well characterized P-gp substrates daunomycin and etoposide, as well as cisplatin, which is not transported by P-glycoprotein.75 Resistance of P-glycoprotein-overexpressing KBC-1 cells against daunomycin but not cisplatin was slightly but significantly reduced when KP1019 was added at low, nontoxic concentrations.  Following the assumption that Ru(II) species may be the active ones following Ru(III) prodrug(s) treatment, Ru(II) complexes were also synthesized and investigated for their anticancer properties. Within this strategic framework, organometallic Ru(II)-arene compounds have been developed, including those containing phosphine, amine, and sulfoxide as co-ligands. These compounds show promising in vitro and/or in vivo antitumor activities.77, 78 Interestingly, while monofunctional complexes of the type [η6-arene)Ru(en)X]+ (en = ethylenediamine or derivatives, X = halide) (Figure 3) exhibit high cytotoxicity in vitro comparable to that of cisplatin, which can be modulated by the arene ligand,79 the bi-functional complexes of the general formula [η6-arene)Ru(PTA)X2] (PTA = 1,3,5-triaza-7-phosphaadamantane, X = halide) (RAPTA, Figure 3) showed antimetastatic properties and generally low toxicity as reported for NAMI-A.77 The pharmacological properties of mono-functional Ru(II) complexes have been mainly attributed to their reactivity with nucleic acids leading to DNA damage and cell death, even if a different DNA mode of    	138	binding is observed compared to cisplatin. Instead, the RAPTA compounds appear to work on molecular targets other than DNA, implying a biochemical mode of action profoundly different from that of classical Pt anticancer drugs. Indeed, it is likely that the mechanism of action of the RAPTA complexes may involve interactions with critical intracellular or even extracellular proteins.77, 80   Figure 3.  Organometallic Ru(II) arene complexes.  Concerning possible transport mechanisms, Sadler et al. investigated two iminopyridine ruthenium(II) arene complexes, which differ in their halide ligands, Ru( η6-p-cymene)(N,N-dimethyl-N’-[(E)-pyridine-2-ylmethylidene]benzene-1,4-diamine)X]PF6 (X= Cl, I, Figure 3).81  Possible pathways for the accumulation of these two organometallics were studied in human ovarian cancer A2780 cells in comparison to cisplatin.  Cells were co-incubated with the compounds and different concentrations of one of the following substances: a) verapamil (competitor for efflux via P-gp), b) oubain (inhibition of Na+/K+ pump), c) CuCl2 (competitor for transport via hCtr1), d) antimycin A (ATP depletion), e) amphotericin B (membrane disruption and model for protein-mediated transport) and f) methyl ß-cyclodextrin (caveolae endocytosis pathway); the amount of metal (Ru/Pt) uptake was determined by ICP-MS. Interestingly, by changing from a chloro to an iodo ligand, the mechanism of uptake varied from being active to mainly passive, respectively. Nevertheless, competition experiments with Cu2+ (200 µM) indicate that, while Pt accumulation from cisplatin treatment is reduced by ca. 40%, accumulation of Ru is only reduced of 26% for the chloro complex.81 Instead, Ru uptake for the iodo derivative is reduced to a third of its original value. The experiments using the cardiac glycoside oubain suggest that the membrane potential of the cell and the corresponding electrochemical gradient are key determinants of Ru compounds uptake. The role of protein-mediated transport in the cellular accumulation of Pt and Ru drugs was also investigated co-incubating cells with variable concentrations of amphotericin B, which forms pores in the cellular membrane. These pores, permeable to water and non-electrolytes, may give rise to increased drug influx and therefore higher cellular accumulation. In this case, the obtained results showed no effect on the uptake of the chloro derivative, but had a marked influence on the iodo analogue, in accordance with the results of the temperature-dependent uptake studies, according to which passive diffusion of this complex through the cell membrane is involved. Finally, endocytosis pathways appear not to be involved in the uptake of Ru complexes as shown by co-incubation experiments with methyl ß-cyclodextrin.81  Concerning efflux mechanisms, using verapamil it was possible to impair the efflux of both ruthenium complexes most likely acting on a P-gp dependent efflux pathway.81 In fact, verapamil, an L-type calcium channel blocker, effectively abrogates P-gp mediated active efflux of anticancer drugs in ovarian cancer cells by competitive inhibition of drug transport, and is capable of reversing multi-drug resistance.81 Co-treatment with antimycin A, which can deplete ATP levels and therefore affects the ATP-dependent efflux pump, increased the accumulation of the chloro complex (consistent with verapamil results), but not that of the iodo derivative.81  Overall, these results evidence a highly complex network of accumulation pathways for Ru compounds which are dependent on various determinants including the cell type investigated, the type of ligand set stabilizing the metal centre, the oxidation state of the metal, the possible effects of the compounds on the transporters distribution and expression, as well as the metal complex speciation pathways. For example, reactions with glutathione may lead to RuNNNXX = Cl, IRAPTA-TPNNNRuClClClRuH2NNH2[(h6-biphenyl)Ru(en)Cl]+   	 139	thiolate as well as sulfenate and sulfinate derivatives, which may be recognized by different efflux transporters, as in the case of products from reaction of cisplatin with GSH.   3.2.2 Gold complexes  As mentioned above, gold compounds have attracted increased attention as a source of novel cytotoxic molecules with potential uses in cancer treatment.5, 82, 83 Indeed, both gold(I) and gold(III) complexes have been widely investigated and were found to induce important anticancer effects in vitro and in vivo. Among the various families of gold-based coordination compounds the following were extensively studied for their biological effects: gold(I)  phosphine  complexes,  gold(III)  porphyrins, gold(III)  dithiocarbamates, as well as gold(III) complexes with N-donor ligands. Representative structures for each family are presented in Figure 4.  It is worth mentioning that, in spite of their promising anticancer effects, the risk in developing gold compounds for biological applications is that they may be characterized by a remarkable oxidizing character (particularly Au(III)), especially within the fairly reducing intracellular milieu. Therefore, in order to guarantee more controlled chemical speciation in an aqueous environment, different types of organometallic gold complexes have been synthesized in which the presence of a direct carbon–gold bond greatly stabilizes the gold(I)/(III) redox couple.84 Thus, a variety of cyclometallated gold(III) complexes of nitrogen donor ligands have been synthesized, featuring both bidentate C,N- and terdentate C,N,N-, C,N,C- and N,C,N-donor ligands, with either five- or six-membered C,N rings (Fig. 4).84 Similarly, organometallic gold(I)/gold(III) complexes with N-heterocyclic carbene (NHC) ligands have also been explored as cytotoxic agents (Fig. 4).84, 85 In general, both organometallic gold(I) and gold(III) centers have increased stability with respect to classical gold-based coordination complexes and are extremely suitable to design gold compounds still acting as pro-drugs, but in which the redox properties and ligand exchange reactions can be modulated to achieve selective activation in diseased cells.    Figure 4.  Examples of cytotoxic coordination and organometallic gold(I) and gold(III) compounds.   Concerning Au(I) phosphine compounds which have shown early promise as anticancer drugs, they can be divided into two distinct classes based on coordination chemistry and propensity to undergo ligand exchange reactions with biological thiols and selenols. These are (i) neutral, linear, two-coordinate complexes such as auranofin; and (ii) lipophilic, cationic, bis-chelated tetrahedrally four-coordinate Au(I) diphosphine complexes such as [Au(dppe)2]+ (dppe=1,2- bis(diphenylphosphino)ethane). Although evidence suggests that the mechanism of antitumor activity of the two classes is different, mitochondria have been implicated as targets in both cases.86 Notably, among the members of the latter family, a bis-chelated Au(I) complex of the water-soluble bidentate pyridylphosphine ligand 1,3-bis(di-2-NAuNNAuNO2+NNAuNNOO NNNNOOBF4-N-heterocyclic carbenes+NAuClClNNNAu Cl2 Cl-N-donor ligands2+OH3C(O)COH3C(O)CO OC(O)CH3SOC(O)CH3Au PEt3phosphanes(auranofin) cyclometallatedNNNNAu+porphyrinsH3CH2CO CH2ONCH3CSSAuBrBrdithiocarbammates   	140	pyridylphosphino)propane (d2pypp), namely [Au(d2pypp)2]Cl (Figure 5),87 was designed with  a lipophilicity (log P = -0.46) in the optimal range derived from predictive models for the selective accumulation of the so-called DLCs (Delocalized Lipophilic Cations) in cancer cells based on lipophilicity.88   It is worth mentioning that DLCs have been explored as an approach to cancer chemotherapy that exploits their selective accumulation in mitochondria of cancer cells as a consequence of the elevated transmembrane mitochondrial potential Δψm.89 In fact, DLCs can pass easily through the lipid bilayer and their positive charge then directs them to the mitochondria where they accumulate at significantly higher concentrations than in the cytoplasm, owing to the large Δψm generated by the respiratory chain.90 While DLCs share a common mechanism for mitochondrial accumulation, their structures are diverse and consequently their mechanism of antitumor action and mitochondrial targets may vary.     Figure 5.  Examples of delocalized lipophilic cations (DLCs) including a Au(I) complex.  Noteworthy, in addition to the increased Δψm, some cancer cells have been found to have higher plasma membrane potentials (Δψp), which further contributes to the increased uptake of DLCs by cancer cells.91 Since all cells generate an electrical gradient, which is negative on the inside of the cell, Δψp could act as an attractive force for intracellular accumulation of agents of this nature. Notably, preferential accumulation and toxicity of DLCs has been demonstrated in a number of different carcinoma cell lines as compared to normal epithelial cell types. As an example, Chen and coworkers studied more than 200 epithelial-derived cell lines and found that carcinoma cells consistently had a higher level of uptake and retention of Rhodamine-123 (Rh-123) than normal human epithelial cells.92 Taken together, to date there is direct evidence supporting a link between the retention of positively charged compounds in tumor cells and increased Δψp,93 while the role of Δψm in this phenomenon remains to be further investigated.94 Nevertheless, recently Ott and coworkers evaluated the cellular uptake as well as the biodistribution of a series of Au(I) NHC complexes by atomic absorption spectroscopy.95 According to their results, a marked mitochondrial accumulation of compound triphenylphosphine-[1,3-diethylbenzimidazol-2-ylidene]gold(I) could be related to its high cellular uptake and higher lipophilic cationic character within the tested derivatives.95  Apart from the specific case of DLC-type Au(I) complexes, the mechanisms of uptake and accumulation of gold compounds in cells are, in general, poorly established. Since the interaction with thiols is an important parameter in the biochemistry of gold-based drugs, a “thiol shuttle” model involving binding of gold compounds to the surface exposed Cys-34 of serum albumin has been proposed. According to this model, cellular association, intracellular distribution, and efflux of gold complexes via sequential thiol exchange reactions, also involving reversible binding to serum albumin, may take place.96   Recently, neutral heterocyclic Au(I) NHC complexes of the type chloro-[1,3-dimethyl-4,5-diarylimidazol-2-ylidene]gold(I) 1-2 and chloro-[1,3-dibenzylimidazol-2-ylidene]gold(I) 3 reported in Figure 6, modeled on the vascular disrupting anticancer drug combrestatin A-4, were studied for their biological properties in cancer cells.97 The compounds were cytotoxic in the µM range and with distinct selectivity for certain cell lines. The contribution of the various routes of uptake of the test compounds 1-3 was assessed in 518A2 melanoma cells by pre-treating them with AuP PPPNNNNNNNNOH2N NH+ORhodamine 123O[Au(d2pypp)2]Cl   	 141	non-toxic concentrations of specific inhibitors or competitors of the individual uptake processes. Thus, cimetidine (inhibitor) and TEA (competitor) were used to assess possible transport via hOCT1-2, while CuCl2 (competitor) was employed to investigate hCtr1 related transport. Adding oubaine as inhibitor of the Na+/K+ pump could slow down endocytotic processes dependent on a sodium gradient. Subsequently, the cytotoxic effects were measured via MTT assays after 3 h incubation with the compounds, following 2 h pre-incubation with inhibitor/competitor. The results suggest that cellular uptake for all tested gold complexes occurs mainly via the OCT transporters, and for complex 2 also via hCtr1. In addition, complexes 2 and 3 were also internalized via the Na+/K+-dependent endocytosis. Interestingly, complex 1 did not show particular specificity for any of the investigated transport pathways. Unfortunately, these results were not complemented by measurements of the gold uptake in cell extracts, which may have ruled out other effects induced by the compounds’ treatment on the transporters expression and distribution.     Figure 6.  Au(I) NHC complexes chloride-[1,3-dimethyl-4,5-diarylimidazol-2-ylidene]gold(I) 1-2 and chloride-[1,3-dibenzylimidazol-2-ylidene]gold(I) 3. 3.2.3 Iridium complexes  Iridium compounds are another promising class of experimental metal-based cytotoxic agents.98 In a recently published paper from Novohradsky et al., the mechanisms of accumulation of new cytotoxic organometallic iridium(III) complexes were investigated in cancer cells.99 Specifically, a half-sandwich cyclometallated Ir(III) complex [(η5-Cp*)(Ir)( 7,8-benzoquinoline)Cl] bearing a C^N chelating ligand (Figure 7) was studied for its uptake in ovarian cancer A2780 cells in comparison to cisplatin. Beside temperature dependence experiments, co-incubation with different substrates such as a) ouabain (facilitated diffusion endocytosis pathway), b) 2-deoxy-D-glucose and oligomycin (ATP depletion), c) efflux inhibitors such as verapamil (P-gp inhibitor), reversan (MRP1 inhibitor) and buthionine sulfoximine (inhibition of GSH synthesis), d) CuCl2, as well as e) methyl-ß-cyclodextrin (inhibitor of endycytosis pathway) allowed some conclusions.  Overall, passive diffusion seems to be partly involved in the Ir compound uptake, as evinced by the results of the temperature dependence experiments and the blockage of Na+/K+ ATPases. In fact, co-incubation with ouabain, leads to a decrease in Ir intracellular accumulation. Moreover, the competition experiments with CuCl2 suggest the Ctr1 pathway may also be involved in the compound’s uptake, although to a lesser extent. Concerning the possible involvement of efflux systems in the accumulation of the compound, it appeared that the possible efflux transporters P-gp and MRP1 may play a role, as may formation of GSH-Ir conjugates.  Not surprisingly, verapamil and reversan do not restore cisplatin sensitivity since the drug is not recognized by either P-gp100 or MRP1101. Moreover, it could be observed that the endocytotic pathway doesn’t play a role in Ir accumulation. NNAuClOCH3OCH3OCH3OCH31NNAuClOCH3OCH3H3COOCH33NNAuClOCH3OCH3OCH3OCH2CH32F   	142	 Figure 7.  Schematic drawing of the organometallic compound [(η5-Cp*)(Ir)( 7,8-benzoquinoline)Cl]. 3.3 Transporter-targeted anticancer metal compounds  Several studies report on the possibility for metal compounds to be derivatized with biomolecules for different applications.102 Thus, as a strategy to enhance compound uptake via specific peptide transporters, anchoring of metal centers to peptides has been attempted; Fregona et al. reported on gold(III) dithiocarbammate complexes with peptide-based ligands to achieve carrier-mediated delivery of the compounds in cancer cells via peptide transporters (PEPT).103 Two peptide transporters, PEPT1 and PEPT2, have been identified in mammals, which are present predominantly in epithelial cells of the small intestine, bile duct, mammary glands, lung, choroid plexus, and kidney but are also localized in other tissues (pancreas, liver, gastrointestinal tract) and, intriguingly, appear to be overexpressed in certain types of tumors. A unique feature of such transporters is their capability for sequence independent transport of most possible di- and tripeptides inside the cells. Moreover, they are stereoselective toward peptides containing L-enantiomers of amino acids.  Thus, compounds of the type [Au(III)Cl2(dpdtc)] (dpdtc = dipeptidedithiocarbamate, 5-6) reported in Figure 8 were designed which could both preserve the antitumor properties and reduce toxic and nephrotoxic side-effects of the previously reported gold(III) analogues lacking the peptide moiety, together with an enhanced bioavailability and tumor selectivity due to the dipeptide-mediated cellular internalization provided by PEPTs.104 The compounds showed promising antiproliferative effects in a panel of human cancer cells (PC3, DU145, 2008, C13, and L540), reporting IC50 values much lower than those of cisplatin. Remarkably, the gold compounds also showed no cross-resistance with cisplatin itself and proved to inhibit tumor cell proliferation by inducing either apoptosis or late apoptosis/necrosis, depending on the cell lines. More recently, the same compounds were proven effective in inhibiting tumor growth in breast cancer xenograft models, and proved to be potent inhibitors of the proteasome.105 Unfortunately, the mechanisms of uptake of the compounds via PEPT was not investigated and further studies are necessary to validate the proposed strategy for enhanced gold(III) peptidomimetics transport in tumors.      IrN Cl4BrAuBrSSNCH3NHOOOC(CH3)3HHH HBrAuBrSSNCH3NHOOOC(CH3)3HHH3C CH3OAcOAcOOAcSOAcAu PNNN567   	 143	Figure 8.  Au(III) dipeptidedithiocarbamato and Au(I)-thiosugar complexes.  Interestingly, in recent years, several examples of carbohydrate compounds have been developed for diverse medicinal applications, ranging from compounds with antibiotic, antiviral, or fungicidal activity and anticancer compounds.106 Within this frame, Au(I) complexes with thiosugar ligands, analogues of auranofin (Fig. 8, compound 7), were shown to have an increased uptake in cancer cells.107 It is possible that the 1-thio-β-D-glucose-2,3,4,6-tetraacetate ligand in the compound is acting as a true substrate for the glucose active-transport system (via GLUT1 transporters), enhancing the uptake of the metal compound itself. 4 Conclusions and Perspectives   Pt-based anti-cancer drugs are effective pharmaceuticals and are still among the most widely used agents against malignancies. In parallel to the preparation and screening of new metal complexes as potential anticancer agents, extensive efforts must be directed toward elucidating their mechanism of accumulation in cancerous and non-tumorigenic tissues. Most importantly, a systematic investigational approach should be developed in order to characterize metal compound uptake, efflux and biodistribution in cells. Knowledge of the involvement of metallodrugs transporters could be exploited to develop appropriate intervention schedules. Although in vitro models, as those applied in the studies mentioned above, appear to be more suitable and easy to handle for a first approach to the problem, they are characterized by several limitations, including the absence of an extracellular matrix component and of 3D cell-cell interactions, high variability of results depending on the protocols of culture used in each lab, biased results toward cytotoxic agents and poor correlation with clinical efficacy. Therefore, new pre-clinical methodologies should be explored making use of tissues samples, such as the so-called Precision Cut Tissue Slices (PCTS) technique,108 in which viable slices from explants, from healthy or cancerous tissues, can be considered as mini-models of organs in which cells of different types are present in their natural environment, especially concerning cell-cell and cell-matrix interactions.    Overall, future developments in metal-containing agents and chemotherapeutic regimens should focus on novel delivery mechanisms with emphasis on improving the uptake and proper distribution of the existing metallodrugs by e.g. liposomal formulation of cisplatin-like drugs or by binding the Pt compounds to physiological (carrier) proteins to enhance their uptake.   5 References  1. Wheate, N. J.; Walker, S.; Craig, G. E.; Oun, R. The status of platinum anticancer drugs in the clinic and in clinical trials. Dalton Transactions 2010, 39, 8113-8127. 2. Bergamo, A.; Sava, G. 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Email: a.casini@rug.nl; Fax: +31 50 363 3274 b Institut de Chimie Moléculaire de l’Université de Bourgogne, ICMUB UMR CNRS 6302, 9 avenue A. Savary, 21078 Dijon, France. c Medicinal Inorganic Chemistry Group, Department of Chemistry, University of British Columbia, 2036 Main Mall, Vancouver, BC V6/1Z1, Canada. d Università degli Studi di Sassari, Dipartimento di Chimica e Farmacia, Via Vienna, 2, Sassari, I-07100, Italy. E-mail: cinellu@uniss.it; Fax: +39 079 229559  Published in: Dalton Trans. 2015, 44, 11911-11918.         	 151 1. Abstract A series of novel (C^N) cyclometallated Au(III) complexes of general formula [Au(pyb-H)L1L2]n+ (pyb-H = C^N cyclometallated 2-benzylpyridine, L1 and L2 being chlorido, phosphane or glucosethiolato ligands, n = 0 or 1) have been synthesized and fully characterized using different techniques, including NMR, IR and Far-IR, mass spectrometry, as well as elemental analysis. The crystal structure of one compound has been solved using X-ray diffraction methods. All compounds were tested in vitro in five human cancer cell lines including lung, breast, colon and ovarian cancer cells. For comparison purposes, all compounds were also tested in a model of human healthy cells from embryonic kidney. Notably, all new compounds were more toxic than their cyclometallated precursor bearing two chlorido ligands, and the derivative bearing one phosphane ligand presented the most promising toxicity profile in our in vitro screening, displaying a p53 dependent activity in colorectal cancer HCT116 cells. Finally, for the first time C^N cyclometallated gold(III) complexes were shown to be potent inhibitors of the zinc finger protein PARP-1, involved in the mechanisms of cisplatin resistance.    			    																				 	 	152 2. Introduction  Metallodrugs play an important role in the treatment of several diseases. Among them, cisplatin [cis-diamminedichlorido platinum(II)], discovered accidentally,1 is widely used in every second chemotherapy due to its strong effects against ovarian, testicular and lung cancer. Unfortunately, anticancer therapy with cisplatin has a range of limitations, including resistance, limited spectrum of action and severe side effects, including nephrotoxicity. For this reason, since cisplatin’s discovery, numerous Pt-based anticancer agents have been developed to minimize such harmful side effects.2 However, to date, only two additional Pt(II) compounds have achieved international marketing approval, carboplatin and oxaliplatin, although not being deprived of toxic effects.3 In the effort to develop improved anticancer metallodrugs, several other metal complexes have been explored in vitro and in vivo, including compounds of Fe, Ru, Ti and Au.4-6    On the basis of these chemical considerations, the peculiar properties of gold have been exploited for several applications. To date, many complexes containing Au in +1 and +3 oxidation states have shown therapeutic properties against cancer or inflammatory diseases. When exploiting the reactivity of gold derivatives against cells, one should bear in mind also the metal noble character. Thus, the stabilization of the oxidation states +1 and +3 is of paramount importance to observe any kind of biological activity. Otherwise, the metal center can ultimately undergo reduction process, thus leading to the formation of Au(0).7   In this context, the development of anticancer gold organometallics of various families has identified promising candidates with anticancer properties, including Au(I) alkynyl complexes, Au(I/III) N-heterocyclic carbene (NHC) complexes and Au(III) cyclometallated complexes.8-12 The advantage of NHC and cyclometallated organometallic gold compounds is their relative stability with respect to other classical coordination complexes.  There is evidence that Au compounds preferentially target proteins in biological environments, such as the Se-enzyme thioredoxin reductase (TrxR), 13-16 Zn finger enzymes,17,18 as well as the membrane water and glycerol channels aquaglyceroporins.19-21   Notably, increasing interest is given to the family of cyclometallated gold(III) complexes, in which the Au3+ ions are highly stabilized in physiological conditions. Recently, we reviewed the variety of cyclometallated gold(III) complexes of different scaffolds, namely (C^N)Au (C^N^N)Au, and (C^N^C)Au complexes.8 For example, Fricker and coworkers reported the synthesis and biological activity of square-planar six-membered cycloaurated Au(III) compounds with a pyridinyl-phenyl linked backbone and two monodentate or one bidentate leaving group(s), which were able to inhibit the cysteine proteases cathepsins B and K in vitro.22 Structure/activity relationships were investigated by modifications to the pyridinyl-phenyl backbone, and leaving groups, demonstrating optimal activity with substitution at the 6 position of the pyridine ring. In this study, the importance of the leaving groups was also highlighted.22 Moreover, one of the derivatives containing thiosalicylate, as one of the ligands bound to the (C^N)Au scaffold, was tested in vivo against the HT29 human colon tumor xenograft model, where a modest decrease in tumor growth was observed compared to the untreated control tumor.22  Taking into account these promising studies, we report here the synthesis and characterization of a new series of four gold(III) cyclometallated compounds of the general formula [Au(pyb-H)L1L2]n+ (pyb-H = C^N cyclometallated 2-benzylpyridine, L1 and L2 being 1,3,5-triazaphosphaadamantane (PTA), thio-ß-D-glucose tetraacetate (GluS-)  or chlorido ligands; n = 0, 1). Notably, the PTA ligand was chosen for its good water solubility, while the GluS- ligand, was selected with the aim of facilitating uptake into cancer cells modulating the lipophilic/hydrophilic character, as well as via possible interaction with GLUT1 transporters.23 Moreover, binding of Au(III) to thiolate should prevent ligand exchange reactions with biological nucleophiles leading to inactivation of the compound.  The four new gold(III) compounds were obtained in good yields and fully characterized by 1H-NMR, 13C-NMR, 31P-NMR and far-IR, as well as X-ray crystallography for compound [Au(pyb-H)(PTA)Cl][PF6] (2-PF6). All complexes were tested vs. a panel of human cancer cell lines, as well as in non-tumorigenic human embryonic kidney cells HEK-293T in vitro. Since Pt drugs have been argued to elicit apoptosis in certain cell lines via a p53-dependent pathway,24 the human colorectal carcinoma HCT116 p53+/+ cells overexpressing p53, and HCT116 p53-/- cells knock-out for this gene, were selected to compare the antiproliferative effects of the Au complexes with cisplatin. Finally, selected compounds were tested as inhibitors of the enzyme poly(adenosine diphosphate [ADP-ribose]) polymerase 1 (PARP-1).  It is worth mentioning that PARPs are Zn finger proteins playing a key role in DNA repair by detecting DNA strand breaks 	 153 and catalyzing poly(ADP-ribosylation).25,26 Specifically, PARP-1 is involved in cisplatin resistance mechanisms in cancer cells.24 3. Results and discussion 3.1. Synthesis and structural characterization The (C^N) cyclometallated precursor [Au(pyb-H)Cl2] (1) (pyb-H =-C^N cyclometallated 2-benzylpyridine) was synthesized as described by Cinellu et al. by reacting sodium tetrachloridoaurate with 2-benzylpyridine (pyb) in refluxing MeCN/H2O overnight.27 As previously reported, replacement of one chlorido ligand by triphenylphosphine can be achieved by reacting 1 with one equivalent of PPh3 in the presence of excess KPF6 or NaBF4.27 Following this method, we prepared the new phosphane derivative [Au(pyb-H)(PTA)Cl]PF6 (2-PF6) containing 1,3,5-triazaphosphaadamantane (PTA), which is not toxic to cancer cells and is known in general to improve water solubility. Thus, by reacting 1 with one equivalent of PTA in presence of KPF6 for 1.5 hours at room temperature, we obtained 2-PF6 in very good yield (Scheme 1). Complexation of the phosphane ligand as well as the isomeric purity of 2-PF6 were assessed by 31P NMR showing the PF6- anion heptuplet signal centred at -144.4 ppm and a singlet of the coordinated phosphorous at -14.2 ppm, downfield shifted by 86 ppm with respect to the free phosphane ligand.  Of the two possible geometrical isomers, trans-P-Au-N or trans-P-Au-C, the IR and 1H NMR spectra, taken together, support a trans-P-Au-N arrangement. Indeed, in the IR spectrum the Au-Cl stretching vibration is observed at values consistent with a chlorine trans to a carbon atom (see Experimental section) and in the 1H NMR spectrum the H6 proton (for numbering scheme, see Scheme 2) of the pyridine is strongly deshielded with respect to the free ligand (Dd = 0.44 ppm), as usually observed in related complexes and reflects the through-space influence of the adjacent chloride ligand.27 	154  Scheme 1: Synthesis of novel (C^N)gold(III) cyclometallated compounds.    X-ray-quality crystals of 2-PF6 were obtained by slow diffusion of diethyl ether into an acetone solution and the structure in the solid state was solved by X-ray diffraction analysis. The solid state molecular structure of the cation is depicted in Fig. 1 with principal bond lengths and angles reported in Table 1; the corresponding bond parameters of the analogous triphenylphosphine complex [Au(pyb-H)(PPh3)Cl][BF4]28 (5-BF4) are also reported for comparison. The compound crystallizes in the P21/n monoclinic space group and the asymmetric unit contains two independent cations, two independent PF6- anions and one acetone molecule (Fig. S1 in the Supplementary material). The conformations of the two cations are similar and simply differ by a rotation of the PTA groups by about 19°, thus for simplicity reasons, the following description will only concern one of these cations (depicted in Fig. 1). The gold atom displays an almost regular square-planar coordination, with a slight distortion in the N-Au-C angle: 87.39(7)° (Fig. 1). The bite of the six-membered cyclometallated ring, the bond lengths and angles involving the gold atom are comparable to those observed in 5-BF4 by Fuchita (Table 1);28 small differences - e.g. in the Au-N bond distance - may be attributed to the different electronic and steric properties of the two phosphane ligands. Moreover, the six-membered metallacycle is in a boat-like conformation (Fig. 1B) with atoms C1, C6, C8 and N1 essentially coplanar within the estimated standard deviation. This best plane forms dihedral angles with planes C6-C7-C8 and N1-Au1-C1 of 48.75(16) and 39.34(9)°, which is in line with previous observations made by Fuchita et al. for 5-BF4.28 Notably, the PTA ligand is trans to the nitrogen of the cyclometallated ligand, as previously suggested by the FIR and 1H NMR spectra.  NAuClClKPF6, PTAAcetone, r.t.1,5 hNAuClNPNNPF6 NAuClNa2CO3, DCM, r.t., 1.5 h2-PF6    98 %3     96 %1 eq.,2 eq., Na2CO3, DCM, r.t., overnight4       74 %1NAuOAcS OOAcAcOOAcOAcOOAcSOAcOAcOAcOOAcSHOAcOAcOAcOOAcSHOAcOAcOAcOOAc SOAcOAc	 155          Figure 1 - ORTEP view of the cation in complex 2-PF6; top (A) and side (B) views.     Table  1: Selected bond distances (Ä) and angles (°) with estimated standard deviations in parentheses for compound 2-PF6 and 5-BF428 Distances (Å) and angles(°) 2-PF6 (for both independent molecules) 5-BF4 Au-C Au-N  Au-P  Au-Cl C-Au-N  C-Au-P  C-Au-Cl  N-Au-P  N-Au-Cl  P-Au-Cl 2.040(2) 2.042(2) 2.0961(18)  2.0998(18) 2.2709(6)  2.2840(6) 2.3696(5)  2.3612(5) 87.39(7)  87.26(8) 93.44(6)  91.92(6) 178.35(6) 177.56(6) 175.43(5) 177.43(5) 91.25(5)  90.31(5) 87.985(19) 90.512(19) 2.03(1) 2.079(10) 2.311(3) 2.362(3) 85.8(4) 94.8(3) 175.2(3) 176.4(3) 89.4(3) 89.9(1)  Complex 1 was also treated with one or two equivalents of thio-β-D-glucose tetraacetate (GluSH) and sodium carbonate in dichloromethane for 1.5 hours to give compound 3, or overnight to give compound 4 in good yields. The non-planarity and the conformational stability of the cyclometallated Au(III) scaffold give rise to a planar chirality which upon coordination with optically pure tetraacetylated β-D-glucosethiolate leads to the formation of a mixture of diastereoisomers. In both the 1H and 13C NMR spectra, 3 presents doubled 	156 signals in a 50/50 ratio for protons and carbons close to the metal centre due to the equimolar presence of the two possible diastereoisomers. The largest split is observed for H6’ with a difference of 0.25 ppm between the two diastereoisomers, while the signals of H6’ are separated by only 0.03 ppm, thus suggesting close proximity of H6’ to the chiral centres.  The 1H NMR spectrum of 4 was quite complex, showing two signals for each proton of the cyclometallated ligand in a ratio 1/0.3. These signals correspond neither to the precursor 1 nor to the mono-sugar 3. However, the structure of the compound was further assessed by the association of the ESI-MS spectra showing the peak of adduct [4+Na]+ and the elemental analysis corresponding to 4.H2O.  In the absence of a crystal structure determination for complex 3, to discriminate between the two possible isomers, i.e. thio-glucose in trans position to the carbon or to the nitrogen atom, (Figure S2), its far-IR (FIR) spectrum was compared to those of the other complexes, whose structures were known (2-PF6 and 5-BF4) or unambiguous (1 and 4). Indeed, in this range of energy we can observe the stretching bands of Au-Cl and Au-S bonds. The main FIR bands are shown in Table S2 in the Supplementary material.  The dichlorido complex 1 displays a medium band at  358 cm-1 and a strong one at 287 cm-1, corresponding to the stretching of the Au-Cl bonds in trans position to the nitrogen and carbon atoms, respectively. In the case of 2-PF6, the FIR spectrum shows an intense band at 310 cm-1 close to the value (305 cm-1) observed for the analogous complex with PPh3, consistent with a chlorine trans to the carbon atom of the phenyl substituent.  Compound 3 presents two bands: a broad one at 372 cm-1 corresponding to an Au-S bond trans to a nitrogen atom, and a strong one at 295 cm-1 corresponding to the Au-Cl bond in trans position to the carbon atom. As expected, the FIR spectra of 4 did not show any Au-Cl stretching band, but a broad double band with peaks at 375 and 369 cm-1, consistent, respectively, with a sulfur trans to a nitrogen and to a carbon atom.  3.2. Antiproliferative activity Compounds 1, 2-PF6, 3 and 4 were screened for their antiproliferative properties in vitro in a panel of human cancer cell lines including ovarian adenocarcinoma (A2780), mammary carcinoma (MCF-7), lung carcinoma (A549) and colon carcinoma overexpressing p53 (HCT116 p53+/+) or p53 knock-out (HCT116 p53-/-), as well as on healthy human embryonic kidney cells (HEK-293T). The IC50 was determined after 72 hours of incubation with different concentrations of compounds using the classical MTT test. The results are summarized in Table 2.  In general, the new cyclometallated complexes 2-PF6, 3 and 4 were more toxic than their precursor 1, containing two chlorido ligands, in all cell lines with the exception of the A549 cell line in which most of the gold complexes appeared to be poorly toxic. The phosphane-containing complex 2-PF6 presents 	 157 the most interesting toxicity profile, comparable to that of cisplatin in A2780 cells (IC50 = 2.7 ± 0.2 and 1.9 ± 0.6 µM, respectively). Furthermore, complex 2-PF6 is twice as toxic as cisplatin against HCT116+/+ cells (IC50 = 2.1 ± 0.7 and 5.3 ± 0.2 µM) and poorly effective on the HCT116 p53-/- (IC50 = 14.0 ± 1.1 µM). The latter result suggests similar dependence on p53 pathways for compound 2-PF6 as for cisplatin. In terms of selectivity, 2-PF6 is also ca. 3-fold less toxic on the HEK-293T cells compared to the HCT116 p53+/+. Compounds 3 and 4 showed overall moderate antiproliferative properties, and their inactivity rules out the idea that the tetra-acetylated β-D-glucose-1-thiolato ligand may enhance the uptake of the compounds, for example through GLUT-1 transporters. Indeed, our previously reported studies on Au(I) NHC complexes with similar thio-sugar ligands also showed scarce cytotoxic effects most likely due to poor gold uptake.29   Table  2: Antiproliferative effects of compounds 1-4 (IC50 values) compared to cisplatin in different human cancer cell lines after 72 h incubation.   IC50 (µM)a Comp. A2780 HCT116 p53+/+ HCT116 p53-/- MCF7 A549 HEK-293T 1 36.1  ± 7.8 25.5  ± 6.6 21.1  ± 3.1 25.5  ± 4.7 54.4  ± 0.3 21.0  ± 5.1 2-PF6 2.7  ± 0.2 2.1  ± 0.7 14.0  ± 1.1 15.6  ± 4.6 40.5  ± 5.0 7.1  ± 0.8 3 15.7  ± 7.4 9.7  ± 4.8 18.4  ± 1.1 19.7  ± 3.8 40.0  ± 0.7 11.7  ± 6.1 4 17.4  ± 4.5 10.5  ± 2.0 18.5  ± 0.6 15.1  ± 3.9 18.2  ± 1.2 12.9  ± 3.1 cisplatin 1.9  ± 0.6 5.3  ± 0.2 22.9  ± 2.3 20.0  ± 3.0 12.06  ± 0.8 8.6  ± 1.3 a The reported values are the mean ± SD of at least three determinations.  	158 3.3. PARP-1 inhibition In order to further investigate the mechanisms of action of our organometallic compounds, and inspired by our recent results that indicate that some cytotoxic gold(III) compounds are efficient inhibitors of the zinc-finger protein PARP-1,17 we tested complexes 1 and 2-PF6 on the purified human enzyme as described in the experimental section. Remarkably, potent PARP-1 inhibition was indeed observed with both compounds: 1 with IC50  = 1.30 ± 0.40 nM, and 2-PF6 with IC50  = 1.87 ± 0.20 nM, respectively. Notably, these values are in the same range of those previously observed for Au(III) complexes with N-donor ligands.17  4. Conclusions The potential of organometallic compounds for biological applications, including as anticancer agents, have been demonstrated by numerous studies.30 Here, we report the synthesis and characterization of a new series of (C^N) cyclometallated gold(III) complexes bearing different ancillary ligands selected to confer different reactivity and biological properties to the resulting compounds, such as PTA for increased water-solubility, and thio-sugar moieties to influence uptake and reduce exchange with biological nucleophiles. The X-ray structure of compound 2-PF6 was solved and revealed the typical square-planar geometry of the gold(III) cation, as well as the coordination of the phosphane ligand trans to the nitrogen. Compound 1, which is the precursor of the series, was poorly cytotoxic on all cell lines, while the phosphane-containing compound 2-PF6 shows the most promising results against the HCT116 cancer cell line overexpressing p53.  Interestingly, compounds 1 and 2-PF6 inhibited the zinc-finger enzyme PARP-1 in nM concentrations, suggesting the possible design of selective inhibitors and the use of organometallic gold compounds in combination therapies with other anticancer drugs. PARP inhibitors are currently highly investigated for their selective cytotoxic properties and can be considered as DDR inhibitors,31 which can be used in combination with classical DNA damaging agents for optimizing the therapeutic outcome.  Overall, our study shows the potential for improvement of the biological properties of organometallic gold-based compounds by tuning their coordination environment. Further studies are ongoing to evaluate the mechanisms of transport and possible targets for this new series of gold compounds, including interactions with nucleic acids.   5. Experimental section General Remarks All reactions were carried out under purified argon using Schlenk techniques. Solvents were dried and distilled under argon before use. The precursor [Au(pyb-H)Cl2] was 	 159 synthesized according to a literature procedure.26 All other reagents were commercially available and used as received. All the physico-chemical analyses were performed at the “Plateforme d’Analyses Chimiques et de Synthèse Moléculaire de l’Université de Bourgogne”. The identity and purity (≥ 95%) of the complexes were unambiguously established using high-resolution mass spectrometry and NMR. The exact mass of the synthesized complexes was obtained on a Thermo LTQ Orbitrap XL. 1H- (300.13, 500.13 or 600.23 MHz), 13C- (125.77 or 150.90 MHz) and 31P- (121.49, 202.45 or 242.94 MHz) NMR spectra were recorded on Bruker 300 Avance III, 500 Avance III or 600 Avance II spectrometers, respectively. Chemical shifts are quoted in ppm (δ) relative to TMS (1H and 13C) using the residual protonated solvent (1H) or the deuterated solvent (13C) as internal standards. 85% H3PO4 (31P) was used as an external standard. Infrared spectra were recorded on a Bruker Vector 22 FT-IR spectrophotometer (Golden Gate ATR) and far infrared spectra were recorded on a Bruker Vertex 70v FT-IR spectrophotometer (Diamant A225 ATR). X-ray diffraction data for 2 were collected on a Bruker Nonius Kappa CCD APEX II at 115 K.    Synthesis of compounds Synthesis of cyclometallated gold(III) complexes based on 2-benzylpyridine ligand (pyb)       Scheme 2: 1H and 13C labelling used for the attribution of the NMR signals of the cyclometallated ligand.  [Au(pyb-H’)Cl2] (1) A round-bottom flask was charged with NaAuCl4.2H2O (1 eq, 1.19 g, 3.00 mmol) dissolved in distilled water (60 mL). Benzylpyridine (1 eq, 0.482 mL, 3.00 mmol) was added at room temperature and a yellow precipitate was formed. The reaction mixture was refluxed overnight until the yellow precipitate turned white. After filtration, the white solid was washed with methanol, and re-crystallised from a dichloromethane/Et2O mixture to give the pure product (912 mg, 70 % yield). 1H NMR (acetone-d6, 500.13 MHz, 298 K): 4.39 (d, 1H, JAB = 15.6 Hz, CHAHB), 4.66 (d, 1H, JAB = 15.6 Hz, CHAHB), 7.18 (m, 1H, JH-H = 7.6 Hz, H4’), 7.30 (m, 1H, JH-H = 7.6 Hz, H5’), 7.28 (d, 1H, JH-H = 7.6 Hz, H3’), 7.49 (d, 1H, JH-H = 8.0 Hz, H6’), 7.74 (m, 1H, JH-H = 6.8 Hz, H5), 8.05 (m, 1H, JH-H = 7.6 Hz, H3), 8.31 (m, 1H, JH-H = 7.6 Hz, H4), 9.30 (d, 1H, JH-H = 6.4 Hz, H6). Assignments based on 2D-COSY spectra. IR (υmax, cm-1): 3050, 1609, 1564, 1482, 1435, 1024, 829, 747, 358, 287. 	160  	[Au(pyb-H)(PTA)Cl](PF6) (2-PF6) A round-bottom flask was charged with 1 (1 eq, 50 mg, 0.115 mmol) and KPF6 (5 eq, 106 mg, 0.573 mmol) in suspension in acetone (5 mL). PTA (1 eq, 18 mg, 0.115 mmol) was added to the mixture at room temperature leading to the solubilisation of the starting Au complex. The reaction mixture was maintained at room temperature for 1.5 h. After partial removal of the solvent, dichloromethane was added and the solution was filtered through Celite® and the solvents evaporated to dryness. The pure product was obtained after recrystallization from a dichloromethane/pentane mixture (80.3 mg, 98 % yield). 1H NMR (acetone-d6, 500.13 MHz, 298 K): 4.40 (d, 1H, JAB = 15.6 Hz, CHAHB-pyb), 4.61 (2 d, 4H, JAB = 15.6 Hz, JAB = 13.2 Hz, CHAHB-pyb, 3 N-CHAHB-N), 4.81 (d, 3H, JAB = 13.2 Hz, 3 N-CHAHB-N),  4.96 (d, 6H,  3JP-H = 3.6 Hz, N-CH2-P), 7.22 (m, 1H, JH-H = 7.4 Hz, 1.0 Hz, H5’), 7.32 (m, 1H, JH-H = 7.6 Hz, H4’), 7.48 (dd, 1H, JH-H = 7.6 Hz, 1.0 Hz, H3’), 7.77 (m, 1H, JH-H = 7.0 Hz, 1.0 Hz, H5), 7.86 (ddd, 1H, JH-H = 7.5 Hz, 1.0 Hz, 4JP-H = 3.5 Hz, H6’), 8.02 (d, 1H, JH-H = 7.6 Hz, H3), 8.26 (m, 1H, JH-H = 7.8 Hz, 1.0 Hz, H4),  8.99 (m, 1H, H6). 13C{1H} NMR (acetone-d6, 125.77 MHz, 300 K): 47.5 (s, CH2-pyb), 53.6 (d, 1JP-C = 17.6 Hz, P-CH2), 73.1 (d, 3JP-C = 10.1 Hz, N-CH2-N), 125.7 (d, 4JP-C = 3.8 Hz, C5), 127.5 (d, 4JP-C = 3.8 Hz, C3), 129.5 (d, 4JP-C = 3.8 Hz, C5’), 129.8 (s, C4’), 131.1 (s, C3’), 134.6 (d, 3JP-C = 6.3 Hz, C6’), 136.3 (s, Cipso), 142.7 (d, 2JP-C = 3.8 Hz, C-Au), 144.2 (s, C4), 151.4 (s, C6), 156.9 (s, Cipso). 31P{1H} NMR (acetone-d6, 202.45 MHz, 300 K): -16.6 (s, PTA), -144.2 (h, 1JP-F = 711 Hz,  PF6). ESI-MS (MeCN-MeOH), positive mode exact mass for [C18H22N4O3PAuCl]+ (557.09306): measured m/z 557.09246 [M-PF6]+. IR (υmax, cm-1): 2935, 1611, 1565, 1446, 1413, 1283, 1243, 823, 775, 737, 310, 227. Anal. Calc. for C18H22N4O3P2F6AuCl.CH2Cl2: C, 28.97, H, 3.07, N, 7.11 %. Found: C, 28.96, H, 2.02, N, 7.26 %.  [(Au(pyb-H)(GluS)Cl] (3) A round-bottom flask was charged with 1 (1 eq., 50 mg, 0.115 mmol), thio-β-D-glucose tetraacetate (GluSH) (1 eq., 42 mg., 0.115 mmol) and Na2CO3 (2 eq., 24 mg, 0.230 mmol) in suspension in dichloromethane (5 mL). The reaction was maintained at room temperature for around 1.5 h (until the solution turned yellow). The solution was filtered through Celite® and concentrated under reduced pressure. Upon addition of pentane a yellow precipitate was formed that was filtered off and dried under vacuum to give compound 3 as a 1:1 mixture of diastereomers (83.9 mg, 96 % yield). 1H NMR (acetone-d6, 500.13 MHz, 298 K): 1.89/1.94 (2 s, 3H, CH3), 1.95 (s, 3H, CH3), 1.98/1.99 (2 s, 3H, CH3), 2.07/2.09 (2 s, 3H, CH3), 3.74 (m, 1H, CH), 4.02/4.13 (2 dd, 1H, JH-H = 12.0 Hz, 2.0 Hz, CH2-sugar), 4.22/4.26 (2 dd, 1H, JH-H = 12.0 Hz, 5.5 Hz, CH2-sugar), 4.33 (d, 1 H, JAB = 14.5 Hz, CHAHB-pyb), 4.52 (d, 1 H, JAB = 14.5 Hz, CHAHB-pyb),  4.99-5.09 (m, 2H, 2 CH), 5.17-5.25 (2 t, 1H, JH-H = 9.5 Hz, CH), 5.45/5.64 (2 d, 1H, JH-H = 9.5 Hz, CH), 7.08-7.12 (2 m, 1H, JH-H = 5.5 Hz, H5’), 7.16-7.20 (m, 1H, H4’), 7.29-7.31 (2 d, 1H, JH-H = 5.5 Hz, H3’), 7.33/7.58 (2 d, 1H, JH-H = 5.5 Hz, H6’), 7.70-7.73 (m, 1H, H5), 7.98 (broad d, 1H, JH-H = 8.0 Hz, H3), 8.22 (m, 1H, JH-H = 8.0 Hz, H4), 9.17/9.20 (2 d, 1H, JH-H = 5.5 Hz, H6). 13C{1H} NMR 	 161 (acetone-d6, 125.77 MHz, 300 K): 20.6-21.0 (CH3), 47.9 and 48.0 (CH2-pyb), 63.0 (CH2-sugar), 69.7/69.8, 73.0/73.3, 75.4, 76.2/76.4 and 83.0/83.1 (CH-sugar), 125.3 (C5), 127.1 (C3), 128.7 (C4’/5’), 128.8 (C4’/5’), 129.9/130.0 (C3’), 131.9-133.1 (C6’), 134.0/134.1 (C-Au), 143.1/143.2 (C4), 144.5/144.6 (Cipso), 152.0/152.1 (C6), 156.9/157.1 (Cipso), 169.8-170.9 (C=O). ESI-MS (DMSO-MeOH), positive mode exact mass for [C26H29NO9SAuClNa]+ (786.08093): measured m/z 786.07946 [M+Na]+. IR (nmax, cm-1): 1743, 1612, 1569, 1435, 1367, 1219, 1029, 912, 753, 376, 295, 221. Anal. Calc. for C26H29NO9SAuCl: C, 40.87, H, 3.83, N, 1.83, S, 4.20 %. Found: C, 40.58, H, 4.18, N, 1.85, S, 3.73 %.      [Au(pyb-H)(GluS)2] (4) A round-bottom flask was charged with 1 (1 eq., 50 mg, 0.115 mmol), thio-β-D-glucose tetraacetate (2 eq., 84 mg, 0.230 mmol,) and Na2CO3 (5 eq., 61 mg, 0.575 mmol) in suspension in dichloromethane (10 mL). The reaction mixture was maintained at room temperature overnight until the solution turned yellow. The solution was filtered through Celite® and concentrated under reduced pressure. Upon addition of pentane a yellow precipitate was formed which was filtered off and dried under vacuum to give the analytical sample (93.5 mg, 74 % yield). 1H NMR (acetone-d6, 300 K, 500.13 MHz): 1.91 (s, 4H, CH3), 1.93-1.95 (m, 8H, CH3), 2.00 (s, 6H, CH3), 2.09 (s, 3H, CH3), 2.11 (s, 3H, CH3), 3.24-3.28 (m, 0.7H, CH), 3.51-3.56 (m, 0.3 H, CH), 3.76-3.82 (m, 1.7H, CH), 4.02 (dd, 1H, JH-H = 8.0 Hz, 4.0 Hz, CH), 4.06 (m, 0.3H, CH), 4.12 (dd, 1H, JH-H = 8.0 Hz, 2.5 Hz, CH), 4.20-4.32 (m, 2.5H, CH-sugar + CHAHB-pyb), 4.43 (d, 1H, JAB = 14.5 Hz, CHAHB-pyb), 4.50 (d, 0.7H, JH-H = 9.5 Hz, CH), 4.79 (t, 0.7H, JH-H = 9.8 Hz, CHCH2), 4.93 (t, 0.7H, JH-H = 9.8 Hz, CHCH2), 4.97-5.13 (m, 4H, CH), 5.18-5.24 (m, 1.7H, CH), 5.42 (d, 0.7H, JH-H = 10.5 Hz, CH), 7.09-7.15 (m, 2H, H-pyb), 7.30-7.33 (m, 1 H, H-pyb), 7.53 (dd, 0.3H, JH-H = 7.0 Hz, 1.5 Hz, H6’), 7.62 (dd, 0.7H, JH-H = 7.0 Hz, 1.5 Hz, H6’), 7.67-7.72 (m, 1H, H-pyb), 7.96 (d, 1H, JH-H = 7.5 Hz, H-pyb), 8.19-8.24 (m, 1H, H-pyb), 9.39 (dd, 0.7H, JH-H = 6.0 Hz, 1.0 Hz, H6), 9.45 (d, 0.3H, JH-H = 6.0 Hz, H6). 13C{1H} NMR (acetone-d6, 300 K, 125.77 MHz): major isomer: 20.6, 20.7, 20.8, 21.0 and 21.5 (CH3), 48.8 (CH2-pyb), 62.7 and 62.8 (CH2-sugar), 69.4, 69.5, 72.8, 74.7, 75.6, 75.7, 75.9, 76.1, 82.4 and 83.6 (CH-sugar), 125.0, 127.1, 128.0, 128.6, 129.4 and 132.5 (CH-pyb), 135.4 (C-Au), 142.9 and 153.1 (CH-pyb) 154.8 and 157.7 (Cipso-pyb), 169.7, 169.8, 170.0, 170.1, 170.2, 170.2, 170.7 and 170.9 (C=O); minor isomer: 20.6, 20.7, 21.1 and 21.2 (CH3), 48.4 (CH2-pyb), 63.0, 63.2, 69.6, 70.0, 73.6, 75.0, 75.3, 76.4, 76.6, 83.8 and 84.8 (CH-sugar), 123.8, 125.4, 126.9, 127.8, 131.7 and 133.4 (CH-pyb), 134.5 (C-Au), 137.2 (CH-pyb), 150.4 (Cipso-pyb), 152.9 (CH-pyb), 157.9 (Cipso-pyb), 169.8, 169.9, 170.1 and 170.8 (C=O). (CH2Cl2/MeOH), positive mode exact mass for [C40H48NO18S2AuNa]+ (1114.18705): measured m/z 1114.18560 [M+Na]+. IR (nmax, cm-1): 1740, 1614, 1569, 1437, 1367, 1218, 1029, 754, 375, 369, 212. Anal. Calc. for C40H48AuNO18S2.H2O: C, 43.29, H, 4.54, N, 1.26, S, 5.78 %. Found: C, 43.11, H, 4.72, N, 1.30, S, 4.78 %. 	 	162 X-ray crystallography Crystals of 2-PF6 were obtained by slow diffusion of diethyl ether into a concentrated solution of 2-PF6 in acetone. Intensity data were collected on a Bruker Kappa CCD APEX II at 115 K. The structure was solved by direct methods (SIR92)32 and refined with full-matrix least-squares methods based on F2 (Shelx 97)33 with the aid of the Olex2 program.34 All non-hydrogen atoms were refined with anisotropic thermal parameters. Hydrogen atoms were included in their calculated positions and refined with a riding model. Crystallographic data are reported in Table S1 (Supplementary Information Available).   Cell viability assay The human breast cancer cell line MCF7, human lung cancer cell line A549 and human ovarian cancer cell line A2780 (obtained from the European Centre of Cell Cultures ECACC, Salisbury, UK). Human colon cancer cell lines HCT116 p53+/+ and HCT116 p53-/- were a kind gift from Dr. Götz Hartleben (ERIBA, Groningen, NL), while non-tumoral human embryonic kidney cells HEK-293T were kindly provided by Dr. Maria Pia Rigobello (CNR, Padova, Italy). Cells were cultured in DMEM (Dulbecco’s Modified Eagle Medium) or RPMI containing GlutaMax, supplemented with 10 % FBS and 1 % penicillin/streptomycin (all from Invitrogen), at 37°C in a humidified atmosphere of 95 % of air and 5 % CO2 (Heraeus, Germany).  For evaluation of growth inhibition, cells were seeded in 96-well plates (Costar, Integra Biosciences, Cambridge, MA) at a concentration of 10000 cells/well (A2780, MCF-7, HEK-293T) or 6000 cells/well (HCT116, A549) and grown for 24 h in complete medium. Solutions of the gold compounds were prepared by diluting a freshly prepared stock solution (10-2 M in DMSO) of the corresponding compound in aqueous media (RPMI or DMEM for the A2780 or A549, MCF-7, HCT116 p53+/+ and HEK-293T, respectively). Stability in DMSO was checked by NMR, and the compounds resulted to be stable over several hours. Cisplatin was purchased from Sigma-Aldrich and stock solutions were prepared in water. Afterwards, the intermediate dilutions of the compounds in cell culture medium were added to the wells (200 µL) to obtain a final concentration ranging from 0 to 50 µM, and the cells were incubated for 72 h. Afterwards, 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazoliumbromide (MTT) was added to the cells at a final concentration of 0.5 mg ml-1 and incubated for 2 h, then the culture medium was removed and the violet formazan (artificial chromogenic precipitate of the reduction of tetrazolium salts by dehydrogenases and reductases) dissolved in DMSO. The optical density of each well (96-well plates) was quantified three times in quadruplicates at 550 nm using a multi-well plate reader, and the percentage of surviving cells was calculated from the ratio of absorbance of treated to untreated cells. The IC50 value was calculated as the concentration reducing the proliferation of the cells by 50 % and it is presented as a mean (± SE) of at least three independent experiments.   	 163 PARP-1 activity determinations PARP-1 activity was determined using Trevigen’s HT Universal Colorimetric PARP Assay. This assay measures the incorporation of biotinylated poly(ADP-ribose) onto histone proteins in a 96 microtiter strip well format. Recombinant human PARP-1 (high specific activity, purified from E.coli containing recombinant plasmid harboring the human PARP gene, supplied with the assay kit) was used as the enzyme source. 3-Aminobenzamide (3-AB), provided in the kit, was used as control inhibitor. Two controls were always performed in parallel: a positive activity control for PARP-1 without inhibitors, that provided the 100% activity reference point, and a negative control, without PARP-1 to determine background absorbance. The final reaction mixture (50  µL) was treated with TACS-Sapphire, a horseradish peroxidase colorimetric substrate, and incubated in the dark for 30 min. Absorbance was read at 630 nm after 30 min. The data correspond to the mean of at least three experiments performed in triplicate ± SD.  6. References  1. B. Rosenberg, L. Vancamp and T. Krigas, Nature, 1965, 205, 698-699. 2. K. D. Mjos and C. Orvig, Chem. Rev., 2014, 114, 4540-4563. 3. N. J. Wheate, S. Walker, G. E. Craig and R. Oun, Dalton Trans., 2010, 39, 8113-8127. 4. G. Boscutti, L. Marchio, L. Ronconi and D. Fregona, Chemistry Eur. J., 2013, 19, 13428-13436. 5. I. Ott, Coord. Chem. Rev., 2009, 253, 1670-1681. 6. G. Gasser, I. Ott and N. Metzler-Nolte, J Med Chem., 2011, 54, 3-25. 7. E. A. 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Oehninger, Y. Geldmacher, H. Alborzinia, S. Wolfl, W. S. Sheldrick and I. Ott, ChemMedChem, 2014, 9, 1794-1800. 17. F. Mendes, M. Groessl, A. A. Nazarov, Y. O. Tsybin, G. Sava, I. Santos, P. J. Dyson and A. Casini, J. Med. Chem., 2011, 54, 2196-2206. 18. M. Serratrice, F. Edafe, F. Mendes, R. Scopelliti, S. M. Zakeeruddin, M. Grätzel, I. Santos, M. A. Cinellu and A. Casini, Dalton Trans., 2012, 41, 3287-3293. 19. A. De Almeida, G. Soveral and A. Casini, Med. Chem. Commun., 2014, 5, 1444-1453. 20. A. P. Martins, A. Ciancetta, A. deAlmeida, A. Marrone, N. Re, G. Soveral and A. Casini, ChemMedChem, 2013, 8, 1086-1092. 21. A. P. Martins, A. Marrone, A. Ciancetta, A. G. Cobo, M. Echevarría, T. F. Moura, N. Re, A. Casini and G. Soveral, PlosONE, 2012, 7. 22. Y. Zhu, B. R. Cameron, R. Mosi, V. Anastassov, J. Cox, L. Qin, Z. Santucci, M. Metz, R. T. Skerlj and S. P. Fricker, J. Inorg. Biochem., 2011, 105, 754-762. 23. E. Vergara, E. Cerrada, C. Clavel, A. Casini and M. Laguna, Dalton Trans., 2011, 40, 10927-10935. 24. Z. H. Siddik, Oncogene, 2003, 22, 7265-7279. 25. V. Schreiber, F. Dantzer, J. C. Ame and G. de Murcia, Nat. Rev. Mol. Cell. Bio., 2006, 7, 517-528. 26. A. I. Anzellotti and N. P. Farrell, Chem. Soc. Rev., 2008, 37, 1629-1651. 27. M. A. Cinellu, A. Zucca, S. Stoccoro, G. Minghetti, M. Manassero and M. Sansoni, J. Chem. Soc. Dalton. Trans., 1996, 4217-4225. 28. Y. Fuchita, H. Ieda, Y. Tsunemune, J. Kinoshita-Nagaoka and H. Kawano, J. Chem. Soc. Dalton Trans., 1998, 791-796. 29. B. Bertrand, A. de Almeida, E. P. M. van der Burgt, M. Picquet, A. Citta, A. Folda, M. P. Rigobello, P. Le Gendre, E. Bodio and A. Casini, Eur. J. Inorg. Chem., 2014, 2014, 4410-4410. 30. M. A. Cinellu, I. Ott and A. Casini, in Bioorganometallic Chemistry, Wiley-VCH Verlag GmbH & Co. KGaA, 2014, pp. 117-140. 31. S. P. Jackson and J. Bartek, Nature, 2009, 461, 1071-1078. 32. A. Altomare, M. C. Burla, M. Camalli, G. L. Cascarano, C. Giacovazzo, A. Guagliardi, A. G. G. Moliterni, G. Polidori and R. Spagna, J. Appl. Crystallogr., 1999, 32, 115-119. 33. G. M. Sheldrick, Acta Crystallogr. A, 2008, 64, 112-122. 34. O. V. Dolomanov, L. J. Bourhis, R. J. Gildea, J. A. K. Howard and H. Puschmann, J. Appl. Cryst., 2009, 42, 339-341.  	 165  	   Supporting Information Chapter B2 Exploring the potential of gold(III) cyclometallated compounds as cytotoxic agents: variations on the C^N theme  							B. Bertrand,a,b S. Spreckelmeyer,a,c E. Bodio,b F. Cocco, d M. Picquet, b P. Richard,b  P. Le Gendre,b C. Orvig,c M. A. Cinellu,d,* and A. Casinia,* a Dept. Pharmacokinetics, Toxicology and Targeting, Research Institute of Pharmacy, University of Groningen, A. Deusinglaan 1, 9713 AV Groningen, The Netherlands. Email: a.casini@rug.nl; Fax: +31 50 363 3274 b Institut de Chimie Moléculaire de l’Université de Bourgogne, ICMUB UMR CNRS 6302, 9 avenue A. Savary, 21078 Dijon, France. c Medicinal Inorganic Chemistry Group, Department of Chemistry, University of British Columbia, 2036 Main Mall, Vancouver, BC V6/1Z1, Canada. d Università degli Studi di Sassari, Dipartimento di Chimica e Farmacia, Via Vienna, 2, Sassari, I-07100, Italy.       	166 Compound 2-PF6 X-ray diffraction  Figure S1 - The asymmetric unit of complex 2-PF6.    	 167 Table S1 - Crystal data and structure refinement for [Au(pyb-H)(pta)Cl]PF6. Identification code 2 Empirical formula C39H50Au2Cl2F12N8OP4 Formula weight 1463.58 Temperature/K 115 Crystal system monoclinic Space group P21/n a/Å 13.3224(9) b/Å 17.2070(13) c/Å 20.8871(17) α/° 90 β/° 101.168(3) γ/° 90 Volume/Å3 4697.5(6) Z 4 ρcalcg/cm3 2.069 μ/mm-1 6.581 F(000) 2832.0 Crystal size/mm3 0.5 × 0.2 × 0.12 Radiation MoKα (λ = 0.71073) 2Θ range for data collection/° 5.668 to 55.038 Index ranges -17 ≤ h ≤ 14, -22 ≤ k ≤ 22, -26 ≤ l ≤ 27 Reflections collected 105261 Independent reflections 10774 [Rint = 0.0327, Rsigma = 0.0162] Data/restraints/parameters 10774/0/615 Goodness-of-fit on F2 1.052 Final R indexes [I>=2σ (I)] R1 = 0.0155, wR2 = 0.0351 Final R indexes [all data] R1 = 0.0181, wR2 = 0.0358 Largest diff. peak/hole / e Å-3 0.93/-0.62    	168 Medium Infrared  Far Infrared   	 169 HRMS     	170 1H NMR   13C NMR  	 171 31P NMR   Compound 3 Medium Infrared  	172 Far Infrared  HRMS  	 173  1H NMR  	174  13C NMR   Compound 4 Medium Infrared  	 175 Far Infrared  HRMS  	176  1H NMR  	 177  13C NMR    	178 Miscellaneous information  Figure S2 – Schematic representation of the two possible isomers of the monosubstituted Au(III) compounds.        Table S2 - Far-IR absorption bands of the (2-benzylpyridine)-based cyclometallated Au(III) complexes (cm-1).  Compound ῡ(Au-Cl)   trans to N ῡ(Au-Cl)   trans to C ῡ(Au-S)  1 358 287 - 5-BF4 - 305a;  310b - 2-PF6 - 310 - 3 - 295 372 4 - - 375, 369 a M. Z. Cinellu, A.; Stoccoro, S.; Minghetti, G.; Manassero, M.; Sansoni, M., J. Chem. Soc. Dalton Trans., 1996, 4217-4225 b Fuchita : Y. Fuchita, H. Ieda, Y. Tsunemune, J. Kinoshita-Nagaoka and H. Kawano, Dalton Trans, 1998, 791-796  		 179	  Chapter B3 On the Toxicity and Transport Mechanisms of Cisplatin in Kidney Tissues in Comparison to a Gold-based Cytotoxic Agent  												Sarah Spreckelmeyer,a,b* Natalia Estrada-Ortiz,a* Gerian Prins,a Margot van der Zee,a Bente Gammelgaard,c Stefan Stürup,c Inge A. M. de Graaf,a Geny Groothuis,a Angela Casinia,d ,*  a Dept. Pharmacokinetics, Toxicology and Targeting, Groningen Research Institute of Pharmacy. University of Groningen. A. Deusinglaan 1, 9713AV Groningen, The Netherlands.  b Medicinal Inorganic Chemistry Group, Department of Chemistry, University of British Columbia, 2036 Main Mall, Vancouver, British Columbia, V6T 1Z1, Canada c Dept. of Pharmacy, University of Copenhagen, Universitetsparken 2, 2100 Copenhagen, Denmark.  d School of Chemistry. Cardiff University. Main Building, Park Place, CF103AT Cardiff, United Kingdom. Email: casinia@cardiff.ac.uk *Shared 1st authors   Published in: Metallomics, 2017, DOI: 10.1039/C7MT00271H      	180	1. Abstract Mechanisms of toxicity and cellular transport of anticancer metallodrugs, including platinum-based agents, have not yet been fully elucidated. Here, we studied the toxic effects and accumulation mechanisms of cisplatin in healthy rat kidneys ex vivo, using the Precision Cut Tissue Slices (PCTS) method. In addition, for the first time, we investigated the nephrotoxic effects of an experimental anticancer cyclometallated complex [Au(pyb-H)(PTA)Cl]PF6 (PTA = 1,3,5-triazaphosphaadamantane). The viability of the kidney slices after metallodrug treatment was evaluated by ATP content determination and histomorphology analysis. A concentration dependent decrease in viability of PCKS was observed after exposure to cisplatin or the Au(III) complex, which correlated with the increase in slice content of Pt and Au, respectively. Metal accumulation in kidney slices was analysed by ICP-MS. The involvement of OCTs and MATE transporters in the accumulation of both metal compounds in kidneys was evaluated co-incubating the tissues with cimitedine, inhibitor of OCT and MATE. Studies of mRNA expression of the markers KIM-1, villin, p53 and Bax showed that cisplatin damages proximal tubules, whereas the Au(III) complex preferentially affects the distal tubules. However, no effect of cimetidine on the toxicity or accumulation of cisplatin and the Au(III) complex was observed. The effect of temperature on metallodrug accumulation in kidneys suggests the involvement of a carrier-mediated uptake process, other than OCT2, for cisplatin; while carrier-mediated excretion was suggested in the cases of the Au(III) complex.               	 181	2. Introduction Cisplatin (cis-diamminedichloridoplatinum(II), Figure 1) is an antineoplastic drug used in the treatment of many solid tumours, including those of the head, neck, lung, and testis. Unfortunately, severe side effects following cisplatin treatment may occur, including ototoxicity and myelosuppression,1 with the main dose-limiting side effect being nephrotoxicity.2 The pathophysiological basis of cisplatin nephrotoxicity has been studied for the last four decades, and the emerging picture is that the exposure of tubular cells to cisplatin activates complex signalling pathways, leading to tubular cell injury and death via both apoptosis and necrosis.3 Studies in rats and mice suggest that the drug undergoes metabolic activation in the kidney to a more potent toxin, a process possibly involving glutathione and mediated by gluthathione-S-transferase.4 Activation of cisplatin to its highly reactive and toxic metabolites includes spontaneous intracellular aquation reactions, which involve the substitution of the chlorido ligands with water/hydroxide molecules.5,6 Previously reported studies using kidney slices,7 cultured renal epithelial cells8 and isolated perfused proximal tubule segments9 have provided evidence for basolateral carrier-mediated uptake of cisplatin. Moreover, it was found that cisplatin concentration within the kidneys exceeds the concentration in blood by at least five-fold, suggesting accumulation of the drug by renal parenchymal cells.7 At a molecular level, experimental evidence has led to the conclusion that cisplatin enters cells via two main pathways: (i) passive diffusion and (ii) facilitated uptake by a number of transport proteins,6,10 including copper transporters (CTR) and organic cation transporters (OCT).6,11 Pabla et al demonstrated that CTR1 is mainly expressed in both proximal and distal tubular cells in mouse kidneys, whereas cisplatin toxicity has been observed mainly in the proximal tubular cells.11 In the same study, it was shown that down-regulation of CTR1 in human embryonic kidney cells (HEK293), by small interfering RNA or copper (Cu(I)) pre-treatment, resulted in decreased cisplatin uptake.  Various studies, demonstrating that cisplatin can be transported by OCTs in cells, are based on competition experiments with other established OCTs substrates such as tetraethylammonium (TEA) and inhibitors such as cimetidine.12–14 OCTs belong to the solute carrier SLC22A family consisting of three sub-categories: the electrogenic transporter (OCT1-3), electroneutral organic cation/carnitine transporter (OCTN1-3) and the organic anion transporter (OATs and urate transporters, URAT-1).15 Many transporters of the SLC22A family are found in secretory organs such as the liver and the kidneys, as well as the intestine, where they play pivotal roles in drug adsorption and excretion.16 Moreover, different OCTs show species and tissue-specific distribution. For example, the human OCT1 is highly expressed in the sinusoidal membrane of the liver and in the apical membrane of the jejunum17 but not in the kidney. Instead, human OCT2 is mainly expressed in the basolateral side of renal proximal tubule cells, and in the dopaminergic brain regions.18 In order to correctly interpret translational studies, it is important to note that in rodents, both OCT1 and OCT2 show a high renal expression in the basolateral membrane of proximal tubule cells. 19–21 hOCT3 shows a much broader tissue distribution, including skeletal muscle, heart, brain, and placenta, but the distribution in the membrane and physiological role of OCT3 are not yet clearly understood.16 The interaction of cisplatin with hOCT2 in the kidney, or hOCT1 in the liver, was investigated with the fluorescent cation 4-[4-(dimethyl-amino)styril]-methylpyridinium (ASP) in stably transfected HEK293 cells overexpressing these transporters, and in cells physiologically expressing them, such as human proximal tubules and human hepatocyte couplets.22 Notably, cisplatin inhibited ASP transport in hOCT2-HEK293 but not in hOCT1-HEK293. Furthermore, incubation with cisplatin induced apoptosis in hOCT2-HEK293 cells; a process that was completely suppressed by simultaneous incubation with the hOCT2 inhibitor cimetidine. Moreover, in isolated human proximal tubules, cisplatin competed with basolateral organic cation transport, whereas it had no effect in human hepatocytes.22 Overall, these findings support the idea of the interaction of cisplatin with hOCT2 in renal proximal tubules, but not with hOCT1, possibly explaining its organ-specific toxicity. In 2010, the functional effects of cisplatin treatment on kidney and hearing were studied in vivo in wild-type and OCT1/2 double-knockout mice.23 No sign of ototoxicity and only mild nephrotoxicity were observed after cisplatin treatment of knockout mice, while cisplatin accumulation in the kidneys was reduced.23 Co-medication of wild-type mice with cisplatin and the organic cation cimetidine resulted in protection against 	182	ototoxicity and partly against nephrotoxicity.23 Moreover, experiments in rats showed that treatment with both cisplatin and cimetidine did not interfere with the antitumoral activity of the Pt drug.24 Based on these studies and others, hOCT2 has been proposed as a target for protective therapeutic interventions in cisplatin chemotherapy.  Furthermore, membrane transporters are also involved in carrier-mediated Pt efflux pathways, including the ATP-binding cassette (ABC) multidrug transporters6,25 and the multidrug and toxin extrusion proteins (MATEs).26–28 MATEs belong to the SLC47 family and are also part of organic cation homeostasis. Specifically, MATEs act as H+/organic cation antiporters, transporting protons from the extracellular side to the cytoplasm in concomitance with organic cations export to the lumen of the proximal tubule. Two isoforms are known, SLC47A1 (MATE1) and SLC47A2 (MATE2-K). Both in human and in rat, MATE1 is primarily expressed in the liver and kidney, while MATE2-K exhibits a kidney-specific expression at the brush border membrane of the tubular cells. 29 Several studies have confirmed cisplatin transport by MATEs.13,14 Overall, these studies suggest that, in humans, the interplay between OCT2 and MATE is responsible for the net renal secretion of cisplatin, and possibly also for the net accumulation of cisplatin in the tubular cells, but further investigation is essential to fully elucidate the complex pathways of cisplatin transport and related side-effects.30 Within this framework, the lack of conclusive information is at least partly due to the lack of suitable models to study transport mechanisms in renal tissues. In vitro models, generally 2D cell cultures, have been applied to study the mechanisms of action, metabolism and transport of metallodrugs.30 These 2D cultured cells usually are characterized by low level of differentiation and mostly consist of one cell type, thereby lacking interactions between the different cell types as in a tissue. Therefore, a model including all cell types in their natural environment is indispensable for studying complex, multicellular organ functions and the pharmacological and toxicological response to drugs, as well as for the identification of the transport mechanisms. The precision cut tissue slices (PCTS) is such a technique, where the original cell-cell and cell-matrix contacts stay unaltered and as such is a useful technique for drug testing ex vivo.31 In a PCTS model, the tissue can remain viable during culture with physiological expression and localization of enzymes and transporters. Thus, the PCTS system is uniquely suited to examine molecular responses to toxicant exposures and compare species differences, and is nowadays a FDA-approved technology. 32 Recently, our group has successfully used the PCTS technique to study the toxic effects of experimental anticancer organometallic compounds,33–36 aminoferrocene-containing pro-drugs,37 ruthenium-based kinase inhibitors,38 as well as supramolecular metallacages as possible drug delivery systems.39 Interestingly, nephrotoxic side effects induced by cisplatin were already investigated in human and rat kidney slices and characterized morphologically, as well as in terms of gene expression and functional changes, providing evidence for the mechanisms of apoptosis induction.40–42 Furthermore, the acute nephrosis of tubular epithelium induced by cisplatin in vivo was reproduced in both human and rat kidney slices ex vivo, while the glomerulus appeared unaffected even at high drug concentration (80 µM).40  However, further studies are necessary to evaluate possible transport mechanisms using the precision-cut kidney slices (PCKS) ex vivo model.  Here, we report on the toxicity and mechanisms of accumulation and transport of cisplatin studied in rat kidney using the PCKS technique. Tissues viability was assessed by three different methods, including ATP content, histomorphology, and mRNA determination of different biomarkers. Moreover, intracellular metal accumulation was determined by inductively coupled plasma mass spectrometry (ICP-MS). In addition, the involvement of carrier–mediated transport was investigated performing experiments in the presence of cimetidine, an inhibitor of both rat OCTs and MATE transporters, as well as varying the temperature of tissues incubation. Furthermore, we also studied the toxicity, accumulation and transport mechanisms of another metallodrug, the previously reported experimental cytotoxic cyclometallated (C^N) Au(III) complex [Au(pyb-H)(PTA)Cl]PF6 (PTA = 1,3,5-triazaphosphaadamantane, Figure 1) featuring a relatively stable Au(III) centre.43 Interestingly, this Au(III) complex showed promising antiproliferative effects against several cancer cell lines and inhibits the zinc-finger enzyme PARP-1 in nM concentrations.43 For most of these new generation experimental metal-based compounds with cytotoxicity towards cancer cells, the mechanisms leading to their pharmacological and toxicological profiles are still not fully elucidated and different biological targets and transport systems have been proposed which still need validation.30 	 183	  Figure 1. Structure of the anticancer metal complexes evaluated in this study. 3. Results and discussion 3.1. Toxicity evaluation 3.1.1. ATP content determination.  Initially, to determine the toxicity of the evaluated compounds, PCKS were incubated with different concentrations of cisplatin and Au(III) complex for 24 h. In addition, another set of kidney slices of the same rat were co-incubated with 100 µM cimetidine to assess its effect on the toxicity of cisplatin and Au(III) complex in the PCKS. It is hypothesized that this inhibitor for OCTs and MATE transporters might reduce the accumulation of cisplatin and thereby protects against toxicity. The viability of the kidney slices was determined by measuring the ATP/protein content. The obtained results are presented in Figure 2. Both compounds show concentration-dependent reduction of viability. The Au(III) complex showed a higher toxicity than cisplatin, with TC50 values of 4.3 ± 0.2 µM and 17 ± 2.0 µM, respectively (Table 1). Co-incubation of cisplatin or Au(III) complex with 100 µM cimetidine did not result in any significant change of toxicity in the rat kidney slices. These results are in contrast with previously reported studies in in vitro cellular models. 22,23 It should be noted that cimetidine inhibits not only the uptake transporter OCT2, which is supposed to reduce cellular accumulation, but also the active efflux transporter MATE, which may result in higher accumulation in the slices and thereby increasing toxicity.45 Thus, our toxicity evaluation results suggest that cisplatin and Au(III) complex accumulation is not dependent on rat OCTs or MATEs.  Table 1. TC50 values for PCKS treated with cisplatin or Au(III) complex, in the absence and presence of cimetidine, for 24 h. Compound TC50 No cimetidine + 100 µM cimetidine cisplatin 17.0 ± 2.0 14.7 ± 3.5 Au(III) complex 4.3 ± 0.2 4.4 ± 0.9     	184	 Figure 2. Viability of PCKS treated for 24 h with different concentrations of cisplatin (top) and of Au(III) complex (bottom), without cimetidine (black bars) and co-incubated with cimetidine (grey bars, indicated as: + CIM). The error bars show the standard deviation of at least three independent experiments.  3.1.2. Histomorphology The differential effects of cisplatin and Au(III) complex on PCKS viability, in the absence or presence of cimetidine, were further assessed by histomorphology. Specifically, Periodic Acid-Schiff staining (PAS) was used to evaluate slice integrity and particularly to visualize the basal membranes and epithelial brush border in the proximal tubule. After 24 h incubation, the untreated kidney slices show minor morphological changes, indicated by occasional pyknosis and swelling of tubular cells (Figure 3A). The kidney slices co-incubated with cimetidine (100 µM) showed similar characteristics of integrity as the untreated controls (Figure 3A and 3B). However, the cell swelling is more evident in the samples treated with cimetidine, including slight Bowman’s space dilatation (Figure 3B).  As expected, exposure of PCKS to cisplatin (Figure 3) and to the Au(III) complex (Figure 4) results in damage in several of the cellular structures in the cortex. In the case of cisplatin, at 5 µM concentration (Figure 3C-D) there is evidence of damage on the distal and proximal tubular cells as well as some dilatation of the Bowman’s space in the glomerulus, which seems more prominent in the samples treated with cimetidine. With the increase of the concentration of cisplatin to 10 and 25 µM (Figure 3E-H) the damage intensifies and disruption of the brush borders of the proximal tubules becomes more evident. Furthermore, cimetidine treatment did not reduce the kidney damage induced by cisplatin, and slices resulted to be equally affected by the metallodrug as those incubated without cimetidine.  The tubular damage found in our study for cisplatin is in line with the results of Vickers et al. in rat PCKS. 40 However, in our study we observed that cisplatin also affects the glomerulus structure. This difference in toxicity profiles might be caused by the different culture media used, especially the presence of serum in the 	 185	study of Vickers might be the cause of the difference between these results. In conclusion, our data shows tubular damage by cisplatin, which is not influenced by cimetidine, in line with the ATP viability data.  For the samples exposed to the Au(III) complex, the increase in drug concentration does not produce any major differences with respect to the lower tested concentration. In fact, in all cases, extensive damage is observed mainly in the distal tubular cells, with the structure of the brush border in the proximal tubule almost intact (Figure 4 C-H). No difference was observed in the absence (Fig. 4, left column) or presence (Fig. 4, right column) of cimetidine. While cisplatin generates more damage towards the glomeruli and the proximal tubular cells, the Au(III) complex displayed selective damage of the distal tubular cells, which was also previously described for anticancer Au(I) complexes.36 Overall, the presented histomorphology and ATP results show that there is no evidence of reduced toxicity in the slices exposed to either cisplatin or Au(III) complex co-incubated with cimetidine (Figures 3 and 4, right columns).    Figure 3. Morphology of rat precision cut kidney slices exposed to different concentrations of cisplatin for 24 h. Left column: absence of cimetidine; right column: co-incubation with cimetidine. A and B: 24 h control incubation; C and D: 5 µM; D and E: 10 µM; F and G: 25 µM h. PT: proximal tubule, DT: distal tubule, G: glomerulus, BSD: Bowman’s space dilatation, N: Necrotic areas. Scale bar indicates 50 µm. 	186	 Figure 4. Morphology of rat precision kidney slices exposed to different concentrations of Au(III) complex for 24 h. Left column: absence of cimetidine; right column: co-incubation with cimetidine. A and B: 24 h control incubation; C and D: 1 µM; E and F: 2.5 µM; G and H: 5 µM. PT: proximal tubule, DT: distal tubule, G: glomerulus, BSD: Bowman’s space dilatation, N: Necrotic areas. Scale bar indicates 50 µm. 3.1.3. Expression of kidney-injury molecule-1 (KIM-1), villin, p53 and Bax. The variation of the expression levels of different markers were evaluated in PCKS after incubation with the Pt(II) and the Au(III) metallodrugs. Specifically, kidney injury molecule-1 (KIM-1) and villin were chosen as proximal tubule damage specific biomarkers. An increase of KIM-1 fold expression is expected in the presence of tubular damage, 46,47 whereas decrease in the expression levels of villin is considered a sign of brush border damage.48–50 Additionally, p53 and Bax were studied to determine the role of apoptosis in the toxicity of PCKS upon exposure to cisplatin and to the Au(III) complex. In fact, it has been proposed that cisplatin induced nephrotoxicity consists of activation of multiple stress pathways, including p53 mediated responses and intrinsic and extrinsic apoptosis pathways (with Bax playing an important role in the intrinsic ones).51–54 Thus, KIM-1, villin, p53 and Bax mRNA expression was evaluated in PCKS after 3 h, 5 h and 24 h exposure with cisplatin and Au(III) complex at concentrations close to their TC25 and TC50. The obtained results are shown in Figure 5.   In the untreated control samples, the KIM-1 expression showed a tendency to increase over time compared with the 0 h controls reaching a peak after 24 h, ca. 100-fold increase, (Figure 5A); these results suggest the tubular cells are per se undergoing damage by the slicing and/or culturing process. Interestingly, KIM-1 expression decreased with increasing cisplatin concentration compared to the time-matched controls; even the lowest concentration of cisplatin and the shortest period of incubation (7.5 μM – 3 h) resulted in a 	 187	decreased KIM-1 expression compared to untreated controls. However, due to the high variation, this decrease reached significance only after 24 h exposure to 15 µM cisplatin. Interestingly, upon exposure to the Au(III) complex, no change in KIM-1 expression was observed. From these findings, it can be hypothesized that when slices are exposed to high concentrations of cisplatin, the cellular machinery of the proximal tubular cells is too damaged to be able to produce KIM-1 mRNA.  As shown by the morphological studies, the gold compound seems to target more specifically the distal tubular cells, with reduced damage to the proximal tubular cells, explaining the lack of effect on KIM-1. Concerning villin mRNA expression (Figure 5B), a reduction up to 40% was observed in the controls after incubation for 24 h, indicating some damage of the brush border of the proximal tubular cells. However, no dose-dependent effect of cisplatin was observed, while the Au(III) compound slightly reduced the villin expression, which only reached significance after 5 h at 2 µM.  The assessment of the expression patterns of p53 (Figure 5C) displayed a slight, but not significant increase of the p53 expression during incubation of the control slices. Conversely, upon treatment with cisplatin after 5 h at 15 µM, a decrease of p53 expression is observed, as well as 24 h at 7.5 and 15 µM compared to the untreated samples at each time point. These results differ from the findings in the previously mentioned study of Vickers et al,40 where the expression of p53 increased in rat PCKS after treatment with cisplatin at 20 and 40 µM for 24 and 48 h, respectively.40 Exposure to the Au(III) complex had no effect on p53 expression (Figure 5C, right panel).  Finally, no significant difference in Bax expression levels was observed during incubation of the control samples (Figure 5D). Bax expression increased slightly but significantly after 24 h exposure to the highest concentration of both compounds. These results indicate possible activation of the intrinsic apoptotic pathway depending mainly on the mitochondrial integrity. This finding is in line with previous reports that indicate the possibility of induction of apoptosis independent of p53 after treatment with cisplatin of human and mouse cancer cell lines.54–59  	188	 Figure 5. Fold change of KIM-1 (A), villin (B) p53 (C) and Bax (D) after exposure to cisplatin (left) and to the Au(III) complex (right) during 3 h, 5 h and 24 h. The untreated control (0 h) was set as 1 to calculate the relative fold induction (not shown). The error bars show the standard deviation of at least three independent experiments. Statistical significance was determined by repeated measures ANOVA and Bonferroni as post hoc test to compare the treated samples with the untreated controls for each time point (*: p<0.05, **: p<0.01, ***: p<0.001). 3.2. Uptake studies 3.2.1. Metal content determination by ICP-MS.  In order to assess the intracellular accumulation of the tested compounds and to evaluate the relation between toxicity and intracellular metal content, we determined the Pt and Au content of PCKS exposed to cisplatin and to the Au(III) complex by ICP-MS. Thus, PCKS were incubated for 24 h in the same conditions as for the ATP determination. The concentrations of cisplatin and Au(III) complex used were below or around their TC50. As can be seen in Figure 6, the Pt and Au contents increase as a function of the compounds’ initial concentration (up to 110.3 ng Pt per slice in the case of slices treated with 10 µM cisplatin, and up to 84 ng Au per slice treated with 5 µM of Au(III) complex). However, the obtained results did not show a significant difference between the samples treated with cimetidine or without it, which is in line with the viability and histomorphology studies above, suggesting that OCT and MATE are not involved in the transport of the compounds at the tested concentrations. Moreover, it is worth mentioning that the Au(III) complex appears to 	 189	be more efficiently accumulated into PCKS than cisplatin: the amount of Au and Pt after 24 h of incubation is approximately the same (~28 ng) for the samples treated with cisplatin at 3 µM concentration or Au(III) complex at 1 µM. Additionally, from the obtained results, it can be calculated that the accumulation of cisplatin and of the Au(III) complex results in approximately 20-fold and 30-fold increased concentration of the metals in the slices compared to the medium, respectively. This is in line with the reported accumulation of cisplatin in kidneys in vivo,40 indicating either high binding or metabolism in the cells, or the involvement of active uptake transporters.       Figure 6. Total metal content determined by ICP-MS in PCKS treated with cisplatin or with the Au(III) complex at different concentrations, without and with cimetidine (indicated as: + CIM), for 24 h. The error bars show the standard deviation of at least three independent experiments. 3.2.2. Effect of temperature on uptake in PCKS.  To evaluate if the uptake of cisplatin and of the Au(III) complex is by passive diffusion or carrier-mediated transport, PCKS were incubated with the selected compounds at three different concentrations at 4°C or 37 °C over a period of 60 min. Slices were collected after 0, 10, 30 and 60 min incubation with the metallodrugs, washed with ice-cold Krebs Henseleit buffer and their metal content was evaluated by ICP-MS to assess the effect of different temperatures on the uptake of the drugs.   Slices treated with cisplatin at 5 µM and 25 µM incubated at 4°C or 37°C showed an initial rapid uptake phase followed by a slower accumulation, indicating sequestration by excretion. No significant differences in the Pt content were seen between the two temperatures (Figure 7), suggesting that only passive uptake mechanisms play a role at these tested concentrations. However, slices treated with cisplatin at 100 µM showed significant differences at 30 and 60 min, with a lower Pt content in the slices incubated at 4°C compared to 37°C, indicating that carrier-mediated uptake mechanisms are implicated in cisplatin accumulation in cells at this high concentration. Apparently, both passive and carrier-mediated mechanisms are involved with cisplatin uptake at low concentrations, while carrier-mediated transport is only significantly involved in Pt accumulation at higher concentrations, indicating a low affinity for the transporter.6,26,30  	190	Figure 7. Pt content (ng per slice) in PCKS treated with cisplatin at 5 µM, 25 µM, 100 µM. Incubated at 37°C and 4°C and collected at three different time points (10, 30 and 60 min). t=0 value for the lowest concentration was not determined due to limitations in the amount of tissue, but is estimated to be 0.8 ng Pt/slice based on the values found for the two higher concentrations. The error bars show the standard deviation of at least three independent experiments. Remarkably, evaluation of the Au content in rat kidney slices  upon treatment with the Au(III) complex showed a significant higher Au accumulation at 4°C compared to 37°C at all concentrations and time points (Figure 8). A fast initial uptake rate is followed by a slower uptake rate, which is observed for all concentrations during 60 min. The higher Au content found in the slices incubated at 4°C could be due to the inhibition of active excretion mechanisms such as an efflux transporter, other than MATE, at this low temperature.  Figure 8. Au content (ng per slice) in PCKS treated with 1 at 2 µM (A), 5 µM (B), 10 µM (C). Incubated at 37°C and 4°C and collected at three different time points (10, 30 and 60 minutes). t=0 value for the lowest concentration was not determined due to limitations in the amount of tissue, but is estimated to be 0.5 ng Au/slice. The error bars show the standard deviation of at least three independent experiments. 4. Conclusions In the past decades, several studies have been carried out to elucidate the mechanisms of uptake and efflux of cisplatin on kidney cells, related to the nephrotoxic effects of this extensively used anticancer drug, and various in vitro assays were conducted. Nonetheless, such transport mechanisms are not yet fully understood.30 Moreover, the mechanisms leading to toxicity and accumulation of new generation anticancer gold complexes have not been fully elucidated. Even less is known on the transport of organometallic gold complexes in the kidney. Therefore, we investigated and compared the toxicity and the accumulation of cisplatin and a cytotoxic experimental organometallic Au(III) complex in rat kidney tissues using the PCKS technology. Additionally, we evaluated the involvement of rOCTs and rMATE transporters using their inhibitor cimetidine, in competition experiments. Furthermore, passive or active transport mechanisms were assessed by measuring metal uptake by ICP-MS in PCKS at 37°C or 4°C, respectively.  As expected, a concentration dependent decrease in viability of PCKS was observed after 24 h exposure to both compounds, which correlated with the increase in PCKS content of platinum and gold after treatment. The gold complex seems to be more toxic for the kidney slices than cisplatin based on the TC50’s being 4.3 ± 0.2 µM and 17 ± 2.0 µM respectively. Interestingly, the histomorphological changes after treatment suggest that the Au(III) compound exerts its toxicity towards different target cells than cisplatin, showing extensive damage of the distal tubular cells, whereas cisplatin is more toxic towards the proximal tubular cells. The latter results are in line with previously reported studies.36,40 However, at variance with other studies present in the literature using cell cultures or isolated human tubuli,22–24 in our ex vivo model no effect of co-incubation with cimetidine on the toxicity or accumulation of cisplatin and Au(III) complex was found. Based on these results we conclude that rOCTs and rMATE transporters do not play a prominent role in cisplatin or Au(III) complex accumulation in 	 191	the rat kidney slices at the tested cimetidine concentration. Alternatively, cimetidine may inhibit MATE transporters with higher efficacy than OCT, thereby reducing its renoprotective effect. Furthermore, KIM-1 and villin mRNA expression were studied as markers of proximal tubular damage. KIM-1 expression decreased with increasing cisplatin concentrations, whereas the Au(III) complex induced no change in KIM-1 expression at increasing concentrations. These findings are in agreement with the different localization of the damage in the tissue. In contrast, villin expression was not affected by cisplatin, but was slightly reduced by the Au(III) complex. It could be important to evaluate other markers of damage including markers of structures other than proximal tubular cells to assess kidney injury induced by metallodrugs in a more comprehensive way. Surprisingly, p53 overexpression was not induced in PCKS exposed to cisplatin as previously reported.40 However, as mentioned before, the culture conditions, and specifically the serum protein content of the medium, and the concentrations of cisplatin used were different and these circumstances can lead to substantial differences in the obtained results. Moreover, Bax mRNA expression increased over time in control slices indicating possible activation of intrinsic apoptotic pathways. Nevertheless, the increment is higher when PCKS were treated during 24 h with the highest concentration of cisplatin or gold compound as evidence of further stress compared to the controls.   To get insight into the specific toxic mechanisms activated after treatment with cisplatin or other metallodrugs, it is imperative to consider the species and tissue distinct gene expression profiles during incubation without and with drugs. Specifically, cisplatin is known to activate several stress pathways, but it is dependent on the concentration, cell type and culture conditions whether the cells die by apoptosis, necrosis or both.  Our ex vivo studies to evaluate the passive or active character of the transport of cisplatin and the Au(III) complex revealed that both passive and active processes might play a role. Moreover, the uptake of cisplatin is achieved by both passive and active mechanisms but this becomes evident only at higher concentrations, indicating a low affinity for the active transporters. On the other hand, the results obtained for the Au(III) complex suggested an important role of carrier-mediated excretion, shown by the increased Au content in the slices incubated at 4°C compared to 37°C. Further studies to explore the role of different transporters are needed to better understand the concentration dependent and organ-specific toxicity of our metallodrugs, which is valuable to design new experimental metallodrugs with reduced side effects in specific tissues. Certainly, the use of PCKS offers good opportunities to evaluate toxicity, uptake and accumulation of metallodrugs in different organs and species, and finally to get insight into the effect in human tissues derived from patients. However, optimization of the experimental set-up to reduce the damage of the PCKS induced by culturing is still necessary to exclude possible interference on the obtained results.  Furthermore, new advanced approaches, such as the CRISPR-Cas9 genome editing, should be applied to validate both transport and intracellular trafficking mechanisms for metallodrugs. This technology has been recently applied to individually knock out the human copper transporters CTR1 and CTR2 and the copper chaperones ATOX1 and CCS in cells, in vitro.60 The obtained results suggest that these proteins are not essential for the mechanism by which cisplatin enters human embryonic kidney cells (HEK293T) and ovarian carcinoma OVCAR8 cell lines and is transported to the nucleus, contradicting numerous previously reported studies in the field. Overall, new investigational efforts should be spent to elucidate the complex mechanism of toxicity of metallodrugs and the role of different transport pathways in tissues.  5. Experimental methods Materials Cisplatin and cimetidine were purchased from Sigma Aldrich, and the Au(III) complex was synthesized according to the protocol previously reported.43  PCKS 	192	Male Wistar rats (Charles River, France) of 250-300 g were housed under a 12 h dark/light cycle at constant humidity and temperature. Animals were permitted ad libitum access to tap water and standard lab chow. All experiments were approved by the committee for care and use of laboratory animals of the University of Groningen and were performed according to strict governmental and international guidelines. Kidneys were harvested (from rats anesthetized with isoflurane) and immediately placed in University of Wisconsin solution (UW, ViaSpan, 4⁰C) until further use. After removing fat, kidneys were cut in half lengthwise using a scalpel, and cortex cores of 5 mm diameter were made from each half perpendicular to the cut surface using disposable Biopsy Punches (KAI medical, Japan). PCKS were made as described by de Graaf et al.31,44 The cores were sliced with a Krumdieck tissue slicer (Alabama R&D, Munford, AL, USA) in ice-cold Krebs-Henseleit buffer, pH 7.4 saturated with carbogen (95% O2 and 5% CO2). Kidney slices weighing about 3 mg (~150 μm thickness), were incubated individually in 12-well plates (Greiner bio-one GmbH, Frickenhausen, Austria), at 37°C in culture medium, Williams’ medium E (WME, Gibco by Life Technologies, UK) with glutamax-1, supplemented with 25 mM D-glucose (Gibco) and ciprofloxacin HCl (10 µg/mL, Sigma-Aldrich, Steinheim, Germany) in an incubator (Panasonic biomedical) in an atmosphere of 80% O2 and 5% CO2 with shaking (90 times/min).  In order to remove debris and dead cells before the start of the experiments, PCKS were pre-incubated for 1 h in culture medium and then transferred to new plates containing fresh medium.  Evaluation of ATP content After PCKS pre-incubation, different concentrations of cisplatin and Au(III) complex were added to the wells and the slices were incubated for 10 min, 30 min, 60 min or 24 h. After the incubation time, slices were collected for ATP and protein determination, by snap freezing in 1 ml of ethanol (70% v/v) containing 2 mM EDTA with pH=10.9. After thawing, the slices were homogenized using a mini bead beater and centrifuged. The supernatant was used for the ATP essay and the pellet was dissolved in 5N NaOH for the protein essay. ATP was measured using the ATP Bioluminescence Assay kit CLS II (Roche, Mannheim, Germany) as described in the experimental section. The ATP content was corrected by the protein amount of each slice and expressed as pmol/μg protein. The protein content of the PCKS was determined by the Bio-Rad DC Protein Assay (Bio-Rad, Munich, Germany) using bovine serum albumin (BSA, Sigma-Aldrich, Steinheim, Germany) for the calibration curve.   Evaluation of involvement of rOCT2/rMATE drug transporter  To evaluate the involvement of rOCT2 as uptake transporter and rMATE as efflux transporter, the slices were first incubated for 30 min with a non-toxic concentration of 100 µM cimetidine. Afterwards, cisplatin or the Au(III) complex were added in the selected concentrations. Each condition was evaluated in triplicates after 24 h incubation. Control slices were taken directly after slicing, after pre-incubation and after 24 h incubation.  Histomorphology Kidney slices were fixated in 4% formalin for 24 h and stored in 70% ethanol at 4⁰C until processing for morphology studies. After dehydration, the slices were embedded in paraffin and 4 μm sections were made, which were mounted on glass slides and PAS staining was used for histopathological evaluation. Briefly, the glass slides were deparaffinised, washed with distilled water, followed by treatment with a 1% aqueous solution of periodic acid for 20 min and subsequently Schiff reagent for 20 min. Then, the slides were rinsed with tap water; finally, a counterstain with Mayer’s haematoxylin (5 min) was used to visualize the nuclei. Expression determination of kidney-injury molecule-1 (KIM-1), villin, p53 and Bax. RNA isolation  Three precision cut kidney slices from each treatment group were pooled and snap-frozen in RNase free Eppendorf tubes. RNA was isolated with the Maxwell® 16 simplyRNA Tissue Kit (Promega, Leiden, the Netherlands). Slices were homogenised in homogenisation buffer using a minibead beater. The homogenate was diluted 1:1 with lysis buffer. The mixture was processed according to the manufacturer’s protocol using 	 193	the Maxwell machine. RNA concentration was quantified on a NanoDrop One UV-Vis Spectrophotometer (Thermoscientific, Wilmington, US) right before conversion to cDNA.    cDNA generation  RNA samples were diluted 0,5 μg in 8,5 μl of RNAsa free water. cDNA was generated from RNA using random primers with TaqMan Reverse Transcription Reagents Kits (Applied Biosystems, Foster City, CA). To each sample the following solutions were added: 2,5 μL 5x RT-buffer, 0,25 μL 10mM dNTP’s, 0,25 μL Rnasin (10 units), 0,5 μL M-MLV Reverse Transcriptase (100 units), 0,5 μL random primers.  qPCR  Real-time quantitative PCR was used to determine relative mRNA levels of KIM-1, villin, p53 and Bax. PCR was performed using SensiMixTM SYBR Low-ROX kit (Bioline, London, UK) with the QuantStudio 7 Flex Real-Time PCR System (Thermoscientific, Wilmington, US) with 1 cycle of 10 min at 95°C, 40 cycles of 15 sec at 95°C and 25 sec at 60°C, with a final dissociation stage of 15 sec at 95°C, 1 min at 60°C and 15 sec at 95°C. cDNA for each sample was diluted to 10 ng/μl and measured in triplicate. All primers were purchased from Sigma-Aldrich. Fold induction of each gene was calculated using the housekeeping gene GAPDH.  The primer sequences used in qPCR were: KIM-1: 5′-GTGAGTGGACAAGGCACAC-3′ (forward), and  5′-AATCCCTTGATCCATTGTTT-3′ (reverse);  villin: 5′- GCTCTTTGAGTGCTCCAACC-3′ (forward), and  5′-GGGGTGGGTCTTGAGGTATT′ (reverse);  p53: 5′- CCCCTGAAGACTGGATAAC-3′ (forward), and  5′-AACTCTGCAACATCCTGGGG-3′ (reverse);  Bax: 5′- ACAGGGGCCTTTTTGTTACAG-3′ (forward), and  5′-GGGGAGTCCGTGTCCACGTCA-3′ (reverse);  GAPHD: 5’-CGCTGGTGCTGAGTATGTCG-3’ (forward) and  5’-CTGTGGTCATGAGCCCTTCC-3’ (reverse).  All primers were purchased from Sigma-Aldrich. Fold induction of the evaluated genes was calculated using as reference the housekeeping gene GAPDH.  ICP-MS analysis After incubation with different concentrations of cisplatin or the Au(III) compound, PCKS were washed with ice-cold Krebs-Henseleit buffer and snap-frozen and stored at -80°C until the analysis.  Sample preparation  The tissue samples were digested with 100 µL nitric acid overnight, all samples were completely dissolved. 100 µL hydrochloric acid and 800 µL milliQ were added to produce a volume of 1 mL. Prior to analysis the samples were diluted 20 times with 0.65% HNO3/0.1% HCl.  Pt and Au determination  The Pt and Au contents were quantitated applying a Perkin Elmer (Waltham, MA, USA) Sciex Elan DRC-e ICP-MS instrument, equipped with a Cetac ASX-110FR autosampler, a 0.2 mL min-1 MicroMist U-series pneumatic concentric nebulizer (Glass Expansion, West Melbourne Vic, Australia) and a PC3 cyclonic spray chamber (Elemental Scientific Inc., Omaha, NE, USA). ICP-MS RF power, lens voltage and nebulizer gas and flow were optimized on a daily basis and other settings were: 1 sweep/reading, 25 readings/replicate, 5 replicates, 50 ms dwell time. The 197Au+, 195Pt+, and 194Pt+ isotopes were monitored. Pt and Au concentrations were determined by external calibration (0-20 ppb Pt and Au). LODs were 0.1 and 0.2 µg L-1 for Pt and Au, (3*SD on blank, n=10) 	194	and the spike recovery were 102% and 99% for Pt and Au (n = 3), respectively. Pt and Au single element PlasmaCAL standards (SCP Science, Quebéc, Canada) were used and the standards were prepared in a mixture of 0.1% HCl and 0.65% subboiled HNO3 in MilliQ water. This mixture was furthermore used to dilute samples after digestion and as blank solution.  Temperature dependency  For the evaluation of temperature dependency, after the pre-incubation at 37°C for 1 h, rat kidney slices were incubated for 10, 30 and 60 min with the metal complexes at 37°C or 4°C, and were subsequently washed with ice-cold Krebs-Henseleit buffer and snap-frozen as described above.  Statistics A minimum of three independent experiments were performed using slices in triplicates from each rat kidney. The TC50 values were calculated as the concentration reducing the viability of the slices by 50%, relative to the untreated samples using a nonlinear fitting of log(concentration compound) vs response and is presented as a mean (± SD) of at least three independent experiments. Statistical testing was performed with repeated measures ANOVA and Bonferroni as post hoc test to compare the treated samples with the untreated controls. A p-value of ≤ 0.05 was considered to be significant. In all graphs and tables, the mean values and standard deviation (SD) are shown.     References 1 J. T. Hartmann and H.-P. Lipp, Expert Opin. Pharmacother., 2003, 4, 889–901. 2 R. P. Miller, R. K. Tadagavadi, G. Ramesh and W. B. Reeves, Toxins, 2010, 2, 2490–2518. 3 N. Pabla and Z. Dong, Kidney Int., 2008, 73, 994–1007. 4 D. M. Townsend, M. Deng, L. Zhang, M. G. Lapus and M. H. Hanigan, J. Am. Soc. Nephrol. JASN, 2003, 14, 1–10. 5 L. Kelland, Nat. Rev. Cancer, 2007, 7, 573–584. 6 M. D. Hall, M. Okabe, D.-W. 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Howell, Metallomics, 2016, 8, 951–962. 		198 	  Chapter B4 Investigation of the Molecular Accumulation Mechanisms of an Au(III) Cyclometallated Compound Compared to Cisplatin in vitro: Are OCT2 and CTR1 involved?              S. Spreckelmeyer,a,b M. van der Zee,a B. Bertrand,a,c E. Bodio,c S. Stürup,d and A. Casinia,e a Dept. Pharmacokinetics, Toxicology and Targeting, Research Institute of Pharmacy, University of Groningen, Antonius Deusinglaan 1, 9713 AV Groningen, The Netherlands; E-Mail: a.casini@rug.nl b Medicinal Inorganic Chemistry Group, Department of Chemistry, University of British Columbia, 2036 Main Mall, Vancouver, British Columbia V6T 1Z1, Canada. c Institut de Chimie Moléculaire de l’Université de Bourgogne, ICMUB UMR CNRS 6302, 9 avenue A. Savary, 21078 Dijon, France.  d    Dept. of Pharmacy, University of Copenhagen, Universitetsparken 2, 2100 Copenhagen, Denmark e   School of Chemistry. Cardiff University. Main Building, Park Place, CF103AT Cardiff, United Kingdom.         	 199 1. Abstract The molecular mechanisms of toxicity and cellular transport of anticancer metallodrugs, including platinum-based agents, have not yet been fully elucidated. Here, we studied the toxic effects and metal content (by ICP-MS) of a novel Au(III) compound of the (C^N) theme compared to cisplatin in a panel of cancer cell lines as well as one healthy cell line. A correlation between the concentration dependent toxicity and metal content could be observed. The most sensitive cell lines, A2780 and A2780 cisR, were selected for further transporter studies in vitro. The involvement of OCTs and MATE transporters as well as the copper transporters CTR1 and ATP7A/B in the accumulation of both metal compounds in A2780 cells was evaluated by co-incubating the cells with cimetidine or CuCl2, respectively. The Au(III) compound 1 seems to be more potent compared to cisplatin, when evaluated after 24h and 72h incubation. In addition, an increase in toxicity was seen after 72h for both compounds co-incubated with CuCl2. After 72h, also cimetidine had an effect on the accumulation of both compounds. Uptake studies between 0-120 min incubation showed no effect of either cimetidine or CuCl2 on metal compound accumulation. To conclude, the Au(III) compound 1 and cisplatin might have different mechanisms of transport. The uptake transporter OCT2 and CTR1 are most likely not involved in both accumulations mechanisms, but the efflux transporter MATE and ATP7A/B should be investigated further.                               	200 2. Introduction In the field of metallodrugs, platinum(II) compounds (cisplatin, carboplatin, oxaliplatin and nedaplatin) are still the gold-standard for numerous types of cancer,1 mostly applied in combination with other chemotherapeutics. Unfortunately, their use is limited due to intrinsic or acquired resistance and severe side-effects (eg. nephrotoxicity in the case of cisplatin). Consequently, to develop a potent and specific drug that limits the number of disadvantages described above, new metallodrugs are designed as anticancer agents that incorporate different metals. Many of them are in preclinical studies, but only a very small number reached clinical trials, including NAMI-A and KP1019, which are ruthenium(III) based compounds (Figure 1).2	               Figure  1.  Metallodrug used in the clinic (cisplatin) and metallodrugs containing ruthenium that reached clinical trials (NAMI-A and KP1019).  Among other metal-based complexes, gold(I) and gold(III) compounds raised interest in the last years showing promising anticancer properties in vitro, which prompted the investigation of their molecular mechanisms of biological action.3,4 In Figure 2, representative examples of cytotoxic gold(I) complexes (Auranofin) and gold(III) complexes are presented. Auranofin is currently used as an antirheumatic drug and bears Au(I) as the reactive metal species, stabilized by a thioglucose and a phosphine ligand coordinated in a linear fashion. The complex is highly cytotoxic in vitro against different types of cancer cell lines.5 Nitrogen donor atoms are found to greatly stabilize the Au(III) ions compared to softer donor atoms like sulphur, the latter preferring Au(I). Thus, a vast number of Au(III) complexes with N-donor ligands have been reported so far, including polypyridyl ligands such as [Au(bipy)(OH)2][PF6] (Figure 2). In medicinal inorganic chemistry, the use of Au(I) and Au(III) based organometallic complexes is advantageous due to their redox and thermodynamic stability. Additionally, by modification of the ancillary ligands or steric and electronic properties, the lipophilic character of the anionic cyclometallated ligands can easily be tuned.3            Figure  2.  Au(I) (Auranofin) and Au(III) compounds discussed in this work.  [Au(pyb-H)Cl2] represents an organometallic cyclometallated C^N Au(III) compound, with an even greater stability in aqueous solution compared to [Au(bipy)(OH)2][PF6], due to the direct metal to carbon bond. This Au(III) AuS POOOOOOOOOAuranofinN NAuHO OH[Au(bipy)(OH)2][PF6]PF6-+N AuCl Cl[Au(pyb-H')Cl2]Cisplatin NAMI-A KP1019NHNN NHRuCl ClClClNHNNHNRuDMSOClClCl Cl HNNPtH3N ClH3N Cl	 201 compound was used as the precursor for the synthesis of the Au(III) compound 1 investigated in this chapter. In general, Au(III) compounds exhibit relevant reactivity with several biomolecules, including DNA and proteins, consisting of either ligand exchange processes or redox processes. Depending on the specific nature of the various Au(III) complexes, their electrochemical profile, and the type of reacting species in the target molecule, these reactions may lead either to the formation of tight Au(III)-biomolecule coordinative bonds or to the oxidation and damage of the involved biomolecule itself.5 Protein targets which are relevant for Au(III) complexes are thioredoxin reductase (TrxR), cathepsin cysteine protease and deubiquitinases. Au(III) complexes can both oxidize TrxR at various amino acid sites or form a coordination bond directly with the Seleno-Cys residue and sometimes also with Cys residues in the active site. Au(I) complexes can only form a coordination bond directly with the seleno-cys residue. For this reactions it is essential that both Au(III) or Au(I) complexes undergo ligand exchange reactions to bind to the Seleno-Cys.3 Overall, the interactions of the gold complexes with different biological targets induce direct DNA damage, modification of the cell cycle, mitochondrial damage, proteasome inhibition, modulation of specific kinases, and other cellular processes, which eventually trigger apoptosis. These processes seem to play a major role in the mechanism of the cytotoxic action of gold compounds. 	   However, very little is known about the mechanisms of cellular accumulation (uptake and efflux) of the gold complexes in cancer cells or in healthy tissue. Only one in vitro study has been published thus far, evaluating the role of hOCT1-2 (the human organic cation transporter 1 and 2), hCTR1 (the human copper transporter 1) and endocytotic processes in the uptake of Au(I) NHC (N-heterocyclic carbene) complexes by the 518A2 melanoma cell line, using the MTT assay for toxicity determination as surrogate marker for uptake.6 This clearly shows the lack of extensive mechanistic cellular accumulation studies.7 In general, the differential toxicity of different anticancer drugs in certain organs, such as the nephrotoxicity of cisplatin, may be related to a different Pt accumulation in the cells of these tissues. It is worth mentioning that a higher drug accumulation in a particular cell type might be caused by either a higher uptake or a lower efflux8,9, which emphasizes the need for mechanistic information on the transport mechanisms involved. Even for the anticancer platinum drugs, knowledge about their mechanism of accumulation in cancer cells as well as in healthy cells is incomplete. The OCT2 and CTR1 have been postulated to be involved in the uptake of cisplatin, whereas multi extrusion protein (MATE) and ATP7A/B seem to be involved in its efflux, as recently reviewed by Spreckelmeyer et al.7  In this work, we present the synthesis and characterization of a novel Au(III) cyclometallated compound featuring a coumarin ligand endowed with fluorescence properties for imaging in cells by fluorescence microscopy and to evaluate cell accumulation. The evaluation of the toxic effects of the Au(III) complex compared to cisplatin was performed using a panel of cancer cell lines as well as non-cancer cells in vitro. Moreover, the involvement of OCT2, MATE, ATP7A/B and CTR1 in the uptake and excretion of the gold compound was studied via transporter inhibition experiments in vitro and the accumulation of the compounds was further analyzed via ICP-MS in two A2780 cell lines, one sensitive and one resistant to cisplatin.  3. Results and Discussion 3.1. Synthesis and Characterization A novel cyclometallated Au(III) compound 1 was synthesized adapting already established procedures for similar complexes (Figure 3).10 The C^N precursor [Au(pyb-H)Cl2] 11 was reacted with a phosphine, adding 3-[4-(diphenylphosphino)phenyl]-7-methoxy-2H-chromen-2-one (PPh2Arcoum) to yield the fluorescent Au(III) compound 1.12         	202                 Figure  3.  Synthesis route of Au(III) compound 1.  The complexation of the phosphine ligand with [Au(pyb-H)Cl2] as well as the isomeric purity of Au(III) compound 1 was analyzed by 31P(1H) NMR spectroscopy. The NMR spectrum shows a singlet at 31.5 ppm of the coordinated phosphorous shifted downfield by 35 ppm with respect to the corresponding precursor.10 Moreover, the 1H NMR spectrum of Au(III) compound 1 shows a downfield shift of the signal of the pyridine proton in position 6 by 0.05 ppm associated with a morphological change of the pyridine moiety. A splitting of the signals corresponding to the carbons of the two phenyl rings was observed due to their diastereotopic feature induced by the fixed geometry of the cyclometallated ligand as depicted in Figure 4. Moreover, another splitting of the signal is observed due to the coupling of the phosphorous atom with carbons and hydrogens.         Figure  4. General scheme of the two possible boat-like stereoisomers of [Au(pyb-H)ClP].   3.2. Fluorescence   Au(III) compound 1 in dichloromethane exhibits a typical absorption at 381 nm and a fluorescence emission band of the coumarin chromophore at 429 nm with ΦF = 38 % (Figure 5). Interestingly, this value is higher than the one of the free phosphine ligand (ΦF = 20 %).12 We already noticed this phenomenon when we previously complexed Au(I) on the same coumarin-phosphine ligand.13 Moreover, the emission wavelength is not sensitive to complexation of the chromophore to [Au(pyb-H)Cl2] (λem = 426 nm for the free phosphine ligand in methanol). H2CC NC CAu ClPCH2CNCCAuClPN Au ClClAuNPPH2ClOOOH2PPOOOAcetone, r.t., 1.5 hKPF6,KPF6Au(III) compound 1Pos. 6	 203  Figure  5.  Absorption, excitation and emission spectra of Au(III) compound 1 (dichloromethane, 2.5 x 10-5 M).  3.3. Antiproliferative effects The antiproliferative effects of Au(III) compound 1 were studied in different human cancer cell lines in comparison to cisplatin, via a classical 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyl tetrazolium bromide (MTT) assay after 72 h incubation. Specifically, the selected human cancer cell lines were A2780 (ovarian carcinoma), HCT116 p53 +/+ and -/- (colon carcinoma), MCF 7 (breast carcinoma) and A549  (lung carcinoma), as these cancer types are the most common ones. The tumour protein p53 is a tumour suppressor gene and known to be downregulated in cisplatin resistant cancer tissues.14 The obtained IC50 values are summarized in Table 1. Au(III) compound 1 exhibits a toxicity profile in all tested cell lines which was also found for other Au(III) cyclometallated compounds from the same class with a (C^N) theme.10 However, compared to cisplatin, Au(III) compound 1 shows a higher toxic effect against HCT116 p53 -/- and a significant higher toxic effect against MCF7, while it has a lower toxicity in HCT p53+/+ and a significant lower toxicity in A549 cells. No difference was observed for the toxicity of Au(III) compound 1 in A2780 and against the non-tumorigenic HEK-293T cells with respect to cisplatin. Overall, the observed activity profiles might indicate different mechanisms of action compared to cisplatin as well as different accumulation mechanisms. Due to the highest toxicity against the ovarian cancer A2780 cells, these were selected for further experiments to investigate the transport mechanisms for the Au(III) compound 1 and cisplatin.  Table  1.  Toxicity of Au(III) compound 1 (expressed as IC50 valuesa) compared to cisplatin against different human cancer cell lines and against non-tumorigenic HEK-293Tcells, after 72 h of incubation. For statistical analysis, the t-test was used. * (p ≤ 0.05) indicates the difference is statistically significant when compared to cisplatin treated samples  Compound A2780 HCT116  p53 +/+ HCT116  p53 -/- MCF7 A549 HEK-293T 1 2.4 ± 0.3 9.8 ± 0.9 18.4 ± 1.1 12.6 ± 3* 25.9 ± 5* 10.1 ± 3 Cisplatin 2.3 ± 0.5 5.3 ± 0.2 22.9 ± 2.3 20.0 ± 3 12.1 ± 1 8.6 ± 1.3    3.4. Studies on the mechanisms of transport  3.4.1. Competition experiments 300 400 500 6000.00.51.0wavelength [nm]Normalized  corrected intensity AbsorbanceExcitationEmission	204 The toxicity of the new Au(III) compound 1 and cisplatin were evaluated in human ovarian cancer cells sensitive (A2780) and resistant (A2780cisR) to cisplatin. Both A2780 and A2780cisR cells were recently evaluated from Sorensen et al.15 for the CTR1, OCT2 and ATP7A/B transporters expression levels, as summarized in Figure 6 (reproduced with permission from reference15). The organic cation transporter 2 (OCT2) is an uptake transporter for many substrates, mainly small cationic compounds, and the copper transporter 1 (CTR1) is an uptake transporter for mainly Cu2+ cations. Instead, ATP7A and ATP7B are pumps known to be involved in the efflux of Cu2+. Notably, A2780cisR cells show a significantly higher expression of the copper efflux transporters ATP7A (1.3 fold) and ATP7B (5-fold) as well as the cation uptake transporter OCT2 (1.3 fold). The copper uptake transporter CTR1 is on the other hand lower expressed in A2780cisR cells compared to the A2780 wild type (0.4 fold). These data suggest that the resistance observed in the A2780cisR cells is at least partly caused by a different balance between uptake and efflux of cisplatin resulting from a decreased influx via CTR1 and increased efflux of cisplatin by ATP7B, thereby decreasing the intracellular exposure of the cells to cisplatin.  Figure 6.  Relative drug transporter expression levels in A2780 and A2780cisR cells, reproduced with permission from reference.15  The IC50 values of Au(III) compound 1 and cisplatin were determined after 24h and 72h incubation time in order to observe the development of the toxicity over time (Table 2).  Overall, Au(III) compound 1 is more toxic in the A2780 cells, than in the resistant A2780cisR cells. In addition, after 24h incubation, Au(III) compound 1 is 5-fold more toxic than cisplatin in A2780 cells, and 7-fold more toxic in A2780cisR cells. However, after 72h incubation, Au(III) compound 1 and cisplatin show the same toxicity in A2780, but the Au(III) compound 1 is still ca. 3-fold more toxic in A2780cisR than cisplatin. In addition, there is no significant difference between the IC50s at 24h and 72h treatment of A2780cisR cells with Au(III) compound 1, leading to the assumption that the Au(III) compound does not show accumulation of toxic effects during subchronic exposure. In A2780 cells, the Au(III) compound 1 shows a 2-fold decrease of the IC50 values from 24h to 72h incubation., indicating accumulation of toxicity during chronic exposure.            	 205 Table  2.  IC50 values of Au(III) compound 1 and cisplatin against A2780 and A2780cisR cells, incubated in absence and presence of CuCl2 or cimetidine (Cim), recorded after 24h and 72h incubation. The reported values are the mean ± SD of three independent experiments. For statistical analysis, the Two-way ANOVA was used. * (p ≤ 0.05), ** (p ≤ 0.01) indicate the difference is significant when compared to samples treated with the metallodrugs only (control).  24 h 72 h Compound A2780 A2780cisR A2780 A2780cisR 1 5.8 ± 1.5 15 ± 4 2.4 ± 0.3 11 ± 0.5 1 + Cim 6.6 ± 0.3 18.2 ± 6.1 0.7 ± 0.2** 8.1 ± 0.5* 1 + CuCl2 4.1 ± 0.3 6.1 ± 1.9* 0.2 ± 0.1** 3.1 ± 1.0** Cisplatin 26 ± 2 103 ± 3 2.3 ± 0.5 30 ± 1 Cisplatin + Cim 22.0 ± 2.1 91.7 ± 4.2 1.0 ± 0.1** 21.3 ± 1.1** Cisplatin + CuCl2 22.2 ± 1.3 51.0 ± 4.9** 0.5 ± 0.2** 18.0 ± 0.6**  In order to evaluate the involvement of OCT2 and CTR1 uptake transporters in the accumulation of Au(III) compound 1 and cisplatin, the compounds’ toxicity was further tested in the presence of transporter inhibitors or competitor substrates, respectively. If these transporters are involved in the drug uptake, their inhibition should lead to a reduced intracellular accumulation of the metal complexes and to a decrease in cytotoxic effects and thus an increase in IC50. Cimetidine (300 µM) was selected as inhibitor for OCTs and MATE, and CuCl2 (30 µM)7 as competitive substrate of CTR1 and ATP7A/B. Experiments were performed in A2780 and A2780cisR cells after 24h and 72h incubation with the compounds. CuCl2 and cimetidine do not show a toxic effect on A2780 or A2780cisR cell lines at the used concentrations. The obtained results are summarized in Table 2 and in Figures 7 and 8. After 24h incubation, neither cimetidine nor CuCl2 showed an effect on the toxicity of Au(III) compound 1 or cisplatin in A2780 cells (Figure 7A). However, in A2780cisR cells (Figure 7B), both cisplatin and Au(III) compound 1 showed a significantly increased toxicity when co-incubated with CuCl2. For cisplatin the resistance was only partly reduced, but for Au(III) compound 1 the resistance was fully compensated resulting in a similar toxicity in both cell lines. These results support the hypothesis that CuCl2 is inhibiting de-toxification mechanisms, possibly the copper efflux transporters (ATP7A/B), which are expressed at much higher levels in the A2780cisR cells than in the wild type A2780 cells (Figure 6). Apparently, these efflux transporters play a minor role in the wild type A2780 cells. The lack of effect of cimetidine on the toxicity of cisplatin or Au(III) compound 1 in both cell lines might indicate that the uptake by OCTs and efflux by MATE is not limiting for the toxicity.  	206       Figure  7. IC50 values of Au(III) compound 1 (black) and cisplatin (grey) on (A) A2780 and (B) A2780cisR after 24h incubation. Data are expressed as mean ± SD (n=3). For statistical analysis, the Two-way ANOVA was used. * (p ≤ 0.05), ** (p ≤ 0.01) indicate the difference is significant when compared to samples treated with the metallodrugs only (control).   After 72 h incubation of the A2780 cells, both cimetidine and CuCl2 increased the toxicity of Au(III) compound 1 significantly, 3.5-fold and 12-fold, respectively. For cisplatin, a 2.5-fold and 4.5-fold increased toxicity could also be observed with cimetidine and CuCl2, respectively (Figure 8A). The same effect was observed in A2780cisR cells, but to a lower extent (Fig 8B). This supports the hypothesis, that for both compounds, at 72 h, incubation efflux mechanisms are inhibited by CuCl2 and cimetidine, possibly via inhibition of ATP7A/B and MATE, respectively.                           Control+ 300 µM cimetidine+ 30 µM CuCl 2Control+ 300 µM cimetidine+ 30 µM CuCl 201020304050IC50 [µM]ACisplatinCompound 1Control+ 300 µM cimetidine+ 30 µM CuCl 2Control+ 300 µM cimetidine+ 30 µM CuCl 2020406080100120IC50 [µM]***BCisplatinCompound 1Control+ 300 µM cimetidine+ 30 µM CuCl 2Control+ 300 µM cimetidine+ 30 µM CuCl 20102030IC50 [µM]*******B CisplatinCompound 1Control+ 300 µM cimetidine+ 30 µM CuCl 2Control+ 300 µM cimetidine+ 30 µM CuCl 2024IC50 [µM]** ******ACisplatinCompound 1**	 207  Figure  8. IC50 values of Au(III) compound 1 (black) and cisplatin (grey) on (A) A2780 and (B) A2780cisR after 72h incubation. Data are expressed as mean ± SD (n=3). For statistical analysis (Two-way ANOVA) * (p ≤ 0.05), ** (p ≤ 0.01) indicate the difference is significant when compared to the metallodrugs treated samples (control).   3.4.2. Metal content determination After evaluating the toxic effects of both compounds in cancer cells in the presence and absence of transport inhibitors/competitors, the metal content was determined by inductively coupled plasma mass spectrometry (ICP-MS) to gain further insights into the drug accumulation mechanisms. For these experiments two concentrations of the metal compounds were chosen based on their IC50 and the IC50 times two (IC50 x 2). Interestingly, no differences in Au content could be observed between the two cell lines treated with the same concentrations of Au(III) compound 1 (5 µM or 10 µM at 24 h or 3 µM and 6 µM at 72 h) (Table 3). However, increased accumulation of Au was observed in both cell lines treated with 10 µM of Au(III) compound 1 for 24 h and co-incubated with CuCl2 (Figure 9A, B). The effect of the CuCl2 co-incubation was most marked in the A2780 cells with a 5-fold increase in Au uptake compared to the controls. A 2-fold increase was observed at 5 µM, but this increase was not statistically significant. 	        Table  3. Metal content of Au(III) compound 1 and cisplatin in A2780 and A2780cisR cells, measured after 24h and 72h. The reported values are the mean ± SD of three independent determinations. nd = not determined  Pt or Au content /µg protein  24 h 72 h Compound A2780 A2780cisR A2780 A2780cisR 1 (5 µM) 0.04 ± 0.01 0.04 ± 0.01 nd nd 1 (10 µM) 0.12 ± 0.03 0.10 ± 0.02 nd nd 1 (3 µM) nd nd 0.08 ± 0.01 0.05 ± 0.01 1 (6 µM) nd nd 0.19 ± 0.02 0.15 ± 0.01 Cisplatin (15 µM) 0.07 ± 0.01 0.06 ± 0.01 nd nd Cisplatin (20 µM) 0.17 ± 0.04 0.08 ± 0.02 nd nd Cisplatin (5 µM) nd nd 0.04 ± 0.02 0.02 ± 0.01 Cisplatin (10 µM) nd nd 0.07 ± 0.02 0.05 ± 0.01  Co-incubation of Au(III) compound 1 with 300 µM cimetidine resulted in an increased accumulation of Au only in the A2780 wild type cells at 10 µM of Au(III) compound 1, but not at 5 µM and no effect was seen in the A2780cisR cells (Figure 9A, B).          Control+ 300 µM cimetidine+ 30 µM CuCl 2Control+ 300 µM cimetidine+ 30 µM CuCl 20.00.10.20.3ng Au/ µg proteinB***10 µM5 µMControl+ 300 µM cimetidine+ 30 µM CuCl 2Control+ 300 µM cimetidine+ 30 µM CuCl 20.00.20.40.6ng Au/ µg proteinA***10 µM5 µM	208         Figure  9.  Au content after 24h incubation of (A) A2780 (B) A2780cisR cells treated with 5 µM (black) and 10 µM (grey) of Au(III) compound 1. Data are expressed as mean ± SD (n=3). For statistical analysis the Two-way ANOVA was applied. * (p ≤ 0.05), ** (p ≤ 0.01) indicate the difference is significant when compared to the metallodrugs treated samples (control).  The same type of experiments was repeated after 72 h treatment of cells with Au(III) compound 1 at 3 and 6 µM. After 72 h incubation, in the presence of 30 µM CuCl2 a ca. 2-fold increase in Au content was observed both in A2780 cells and in A2780cisR cells at both concentrations of Au(III) compound 1, except for the co-incubation of 6 µM Au(III) compound 1 in the wild type cells (Figure 10 A, B). This may be explained by the fact that after 72 h, 6  µM is ca. 30 fold the IC50 value for Au(III) compound 1, which may induce extensive cell death, and cell membrane integrity may be altered and the Au compound may have leaked out of the cells during washing of the samples. Cimetidine did not have any significant effect on the Au content of the slices after 72 h of incubation.                 Figure  10. Au content after 72h incubation of (A) A2780 and (B) A2780cisR cells treated with 3 µM (black) and 6 µM (grey) of Au(III) compound 1. Data are expressed as mean ± SD (n=3). For statistical analysis the Two-way ANOVA was used. * (p ≤ 0.05), ** (p ≤ 0.01) indicate the difference is significant when compared to the metallodrugs treated samples (control).  Subsequently, we measured the Au content at different time points during the first 120 min to monitor metallodrug uptake in the cancer cells. Samples were taken after 10, 20, 30, 60 and 120 min in the case of A2780 cells (Figure 11). With cimetidine (Figure 11A) no effect could be observed on Au uptake. With CuCl2 (Figure 11B) only after 120 min, a significantly higher Au content could be observed. In the case of A2780cisR cells (Figure 12), no significant effect of cimetidine or CuCl2 on the Au content could be observed up to 120 min of incubation. Between 10-20 min incubation, a drop in Au content can be observed in A2780 cells. We don’t have an explanation for that thus far, but it should be further investigated in additional experiments.    Control+ 300 µM cimetidine+ 30 µM CuCl 2Control+ 300 µM cimetidine+ 30 µM CuCl 20.00.10.20.30.4ng Au/ µg proteinB****6 µM3 µMControl+ 300 µM cimetidine+ 30 µM CuCl 2Control+ 300 µM cimetidine+ 30 µM CuCl 20.00.10.20.30.4ng Au/ µg proteinA*6 µM3 µM	 209                 Figure  11.  Au content in 3 µM treated A2780 with/without (A) cimetidine (B) CuCl2. Data are expressed as mean ± SD (n=3). For statistical analysis the Two-way ANOVA was applied. * (p ≤ 0.05), ** (p ≤ 0.01) indicate the difference is significant when compared to its control (treatment with 3 µM Au(III) compound 1 taken at the same time).                  Figure  12.  Au content in 3 µM treated A2780cisR cells with/without (A) cimetidine (B) CuCl2. Data are expressed as mean ± SD (n=3). For statistical analysis the Two-way ANOVA was applied. * (p ≤ 0.05), ** (p ≤ 0.01) indicate the difference is significant when compared to its control (treatment with 3 µM Au(III) compound 1 taken at the same time).  When evaluating the Pt content in A2780 and A2780cisR cells treated with cisplatin after 24h (Figure 13A-B) or 72h incubation (Figure 14A-B) by ICP-MS, no significant difference could be observed after co-incubation with cimetidine or CuCl2. However, a marked increase in Pt uptake was observed in A2780 cells treated for 24h with the higher concentration of cisplatin (20 µM) compared to cells treated with the lower concentration (15 µM). This concentration dependent effect could not be observed in the case of the A2780cisR cells at either 24 or 72 h incubation, supporting the idea that higher detoxification and efflux mechanisms may be in place. In Figure 15 and Figure 16, the Pt content within the first 120 min of cisplatin incubation is shown co-incubated with cimetidine or CuCl2. No significant differences between cisplatin alone and co-incubated with the transporter inhibitors are shown, besides for cisplatin at 5 µM (Figure 15A), where cimetidine increased the Pt content at the 120 min time point.             Control+ 300 µM cimetidine+ 30 µM CuCl 2Control+ 300 µM cimetidine+ 30 µM CuCl 20.00.10.20.3ng Pt/ µg proteinB20 µM15 µMControl+ 300 µM cimetidine+ 30 µM CuCl 2Control+ 300 µM cimetidine+ 30 µM CuCl 20.00.10.20.3ng Pt/ µg proteinA 20 µM15 µM10 20 30 40 50 60 70 80 90 100 110 1200.000.050.100.150.20time [min]ng Au/ µg proteinA3 µM compound 1 + 300 µM cimetidine10 20 30 40 50 60 70 80 90 100 110 1200.000.050.100.150.20time [min]ng Au/ µg proteinA3 µM compound 1 + 300 µM cimetidine10 20 30 40 50 60 70 80 90 100 110 1200.000.050.100.150.20time [min]ng Au/ µg proteinB3 µM compound 1 + 30 µM CuCl210 20 30 40 50 60 70 80 90 100 110 1200.000.050.100.150.20time [min]ng Au/ µg proteinB**3 µM compound 1 + 30 µM CuCl2**	210        Figure  13.  Pt content after 24h incubation of (A) A2780 and (B) A2780cisR with 15 µM (black) and 20 µM (grey) cisplatin. Data are expressed as mean ± SD (n=3). For statistical analysis the Two-way ANOVA was used. No sign  differences were found between the controls and the inhibitor-treated cells.                                    Figure  14.  Pt content after 72h incubation in (A) A2780 (B) A2780cisR cells, treated with 5 µM (black) and 10 µM (grey) cisplatin. Data are expressed as mean ± SD (n=3). For statistical analysis the Two-way ANOVA was used. No sign differences were observed between the cells incubated with the inhibitors and their respective controls.            Control+ 300 µM cimetidine+ 30 µM CuCl 2Control+ 300 µM cimetidine+ 30 µM CuCl 20.000.050.10ng Pt/ µg proteinB10 µM5 µMControl+ 300 µM cimetidine+ 30 µM CuCl 2Control+ 300 µM cimetidine+ 30 µM CuCl 20.000.050.100.15ng Pt/ µg proteinA10 µM5 µM10 20 30 40 50 60 70 80 90 100 110 1200.000.010.020.030.04time [min]ng Pt/ µg protein5 µM cisplatin + 300 µM cimetidine*A10 20 30 40 50 60 70 80 90 100 110 1200.000.010.020.030.04time [min]ng Pt/ µg protein5 µM cisplatin + 30 µM CuCl2*B	 211        Figure  15.  Pt content in 5 µM treated A2780 cells with/without (A) cimetidine (B) CuCl2. Data are expressed as mean ± SD (n=3). For statistical analysis (Two-way ANOVA) * (p ≤ 0.05), ** (p ≤ 0.01) indicate the difference is significant when compared to its control (5 µM cisplatin).                      Figure  16.  Pt content in 5 µM treated A2780cisR cells with/without (A) cimetidine (B) CuCl2. Data are expressed as mean ± SD (n=3). For statistical analysis the Two-way ANOVA was applied. * (p ≤ 0.05), ** (p ≤ 0.01) indicate the difference is significant when compared to its control (5 µM cisplatin).   Finally, it is also worth noting that metal accumulation is more efficient in the case of Au(III) compound 1 compared to cisplatin in both cancer cell lines over time (Table 3). This result is in line with the observed toxic effects which are more pronounced for Au(III) compound 1 compared to cisplatin at either 24h or 72h incubation (Table 2), suggesting that a more efficient uptake of compound 1 could explain at least a part of the difference in toxicity.   3.4.3. Passive/active mechanisms To investigate if carrier-mediated or passive molecular mechanisms are responsible for metal accumulation, cells were incubated with the Au(III) compound 1 or cisplatin at either 37°C or 4°C for 10 min, 20 min, 30 min, 60 min and 120 min. This experiment was performed only once (n=1) as a pilot experiment. At 4°C, carrier-mediated transport, including active mechanisms like ATP dependent transporters, is not functional, resulting in complete reduction of carrier-mediated uptake of the substrate, leaving only passive transport. Although at 4°C the viscosity of the cell membrane increases, leading to a decreased fluidity, which may also result in decreased passive diffusion, the reduction in uptake of by passive diffusion is much less than for transporter-mediated uptake.16 In Figure 17, the Au content in A2780 and A2780cisR cells treated with 3 µM Au(III) compound 1 is shown. The Au content is markedly higher at 37°C compared to 4°C in A2780 cells after 60 min incubation, suggesting that active mechanisms are important for Au(III) compound 1 uptake into A2780 cells. In A2780cisR cells, an effect temperature can only be seen after 120 min incubation. These experiments were only performed once, thus the results have to be interpreted with caution.     10 20 30 60 1200.000.020.040.060.080.10time [min]ng Au/µg proteinA10 20 30 60 1200.000.050.100.15time [min]ng Au/µg proteinB10 20 30 40 50 60 70 80 90 100 110 1200.000.020.040.060.08time [min]ng Pt/ µg protein5 µM cisplatin + 300 µM cimetidineA10 20 30 40 50 60 70 80 90 100 110 1200.000.020.040.060.08time [min]ng Pt/ µg protein5 µM cisplatin + 30 µM CuCl2**B	212                 Figure  17.  Au content in (A) A2780 and (B) A2780cisR cells treated with 3 µM Au(III) compound 1 ; red bar: 37°C, blue bar: 4°C (n=1).  Concerning cisplatin, the Pt content does not change at 37°C compared to 4°C, leading to the assumption that active mechanisms of metal accumulation do not play a crucial role within the first 120 min of incubation.       Figure  18.  Pt content in 5 µM cisplatin treated (A) A2780 WT and (B) A280 Res cells; red bar: 37°C, blue bar: 4°C (n=1)   3.5. Copper accumulation The viability results of cisplatin treated cells showed a decreased IC50 value after co-incubation with CuCl2. However, the ICP-MS data did not show an increased Pt content that could be responsible for such an effect. As inhibition of the copper transporters by cisplatin or by the Au(III) complex cannot be excluded, we investigated whether the Cu2+ content was increased due to the co-incubation with each metallodrug. Concerning copper uptake, CTR1 affinity to Cu is between 0.6 µM in fibroblasts and 13 µM in murine hepatocytes.17 ATP7B affinity to Cu is 2.5 * 10-17 M.18 Both transporters are not only selective for Cu, but also for example for Ag(I), Cd(II) and Fe(III).19 	The copper content was evaluated in both the wild-type A2780 and cisplatin resistant A2780cisR cells since the different CTR1/ATP7A/B transporter expression levels (see Figure 6) may have an effect in its accumulation. After 24h, the Cu content in the control cells is the same (0.056 ± 0.02 ng Cu in A2780 and 0.045 ± 0.01 ng Cu in A2780cisR). Overall, it could be observed that the copper content strongly increased in both cell lines upon treatment with the Au(III) compound 1 for 24 and 72h (Figure 19). The co-incubation of Au(III) compound 1 with CuCl2 showed an increase in toxicity compared to cells incubated with Au(III) compound 1 alone. With ICP-MS we showed an increased Au and Cu content after 24h and 72h incubation, that may be responsible for the observed increased toxicity. The effect of the increased Cu-content is not fully understood yet, but an additive or even synergic toxic effect with Au may be possible. To note, a lower Cu content was observed in the A2780 cells treated with 30µM CuCl2 after 24h incubation as well as 10 20 30 60 1200.000.010.020.03time [min]ng Pt/µg proteinA10 20 30 60 1200.000.010.020.03time [min]ng Pt/µg proteinB	 213 after 72h incubation. In order to prevent cell death, the cells might have downregulated the CTR1 uptake transporter due to an excess of CuCl2 in the medium.                                      CuCl2 - + - + - + Au - - 5 µM 5 µM 10 µM 10 µM                WT Res WT Res WT Res WT Res WT Res WT Res0.00.20.4ng Cu/µg proteinA 24h********WT Res WT Res WT Res WT Res WT Res WT Res0.00.20.4ng Cu/µg proteinB 72h** ****	214    CuCl2 - + - + - + Au - - 3 µM 3 µM 6 µM 6 µM  Figure  19.  Cu content in A2780 (WT) and A2780cisR (Res) cells after (A) 24h and (B) 72h incubation with 30 µM CuCl2 and different concentrations of Au(III) compound 1. Data are expressed as mean ± SD (n=3). For statistical analysis (Two-way ANOVA) * (p ≤ 0.05), ** (p ≤ 0.01) indicate the difference is significant when compared to its control (compared to incubation with Cu alone). Also for cisplatin treated A2780 cells (Figure 20), we could observe that, after 24h and 72h incubation, the Cu content increases at both concentrations of cisplatin. However this effect was absent in the A2780cisR cells incubated for 72h.    CuCl2 - + - + - + Pt - - 15 µM 15 µM 20 µM 20 µM                   WT Res WT Res WT Res WT Res WT Res WT Res0.000.050.100.150.20ng Cu/µg proteinA 24h***	 215                     Figure  20.  Cu content after (A) 24h and (B) 72h incubation with 30 µM CuCl2 and cisplatin. Data are expressed as mean ± SD (n=3). For statistical analysis (Two-way ANOVA) * (p ≤ 0.05), ** (p ≤ 0.01) indicate the difference is significant when compared to its control (incubation with 30 µM CuCl2 alone).   Overall, both metal compounds seem to have an effect on Cu accumulation in A2780 cells.  3.6. Fluorescence Microscopy In order to investigate the sub-cellular localization of the Au complex, we performed fluorescence microscopy experiments on A2780 cells treated with Au(III) compound 1 and the two inhibitors cimetidine and CuCl2. Propidium iodide (PI) staining was used to stain for nucleic acids and shows the nuclei. The pictures show the A2780 cells after 2h incubation with Au(III) compound 1 at IC50 x 2 concentration (Figure 21). The compound itself does not co-localize with PI in the nuclei, but clearly enters the cells and is localized in the cytoplasm. This observation points towards a different mechanism of action compared to cisplatin, and suggests that the DNA damage may not play a pivotal role in the toxicity of the Au compound. Co-incubation with either cimetidine or CuCl2 did not change the intracellular accumulation pattern.    Figure 21. Fluorescence microscopy of A2780 cells treated for 2 h with Au(III) compound 1 with/without 300 µM cimetidine or with/without 30 µM CuCl2. Cond. Compound 1                   PI staining Overlay            5 µM    CuCl2 - + - + - + Pt - - 5 µM 5 µM 10 µM 10 µM WT Res WT Res WT Res WT Res WT Res WT Res0.000.050.100.150.200.25ng Cu/µg proteinB 72h****	216 5 µM + 300 µM cimetidine    5 µM + 30 µM CuCl 2      4. Conclusions  In the past years, several studies have been carried out to elucidate the mechanisms of uptake and efflux of cisplatin in cancer cells and kidney slices (eg. Chapter B3). Nonetheless, such transport mechanisms are not yet fully understood.7 Moreover, the mechanisms leading to toxicity and accumulation of new generation anticancer gold complexes have not been fully elucidated. Therefore, we investigated and compared the toxicity and the accumulation of cisplatin and a cytotoxic experimental organometallic Au(III) complex in a panel of cancer cell lines. Additionally, we evaluated the involvement of OCTs, MATEs, CTR1 and ATP7A/B using either their inhibitor cimetidine or competitor CuCl2. The cyclometallated Au(III) compound 1 is more toxic than cisplatin in a number of cell lines, including HCT16 p53 -/-, MCF7 and A2780cisR cells. However, it has a similar or lower toxicity for several other cancer cell lines. The compound is also moderately toxic towards the non-tumorigenic HEK cells. The A2780cisR cell line is not only resistant to cisplatin but also to the Au compound, and this may be based on the differential expression of transporters like OCTs and ATP7A/B with respect to the A2780 cells.  The intracellular accumulation of the Au(III) compound 1 is more efficient than that of cisplatin, as judged from the accumulation of Au and Pt respectively, in both A2780 and A2780cisR cells as demonstrated by ICP-MS. Overall, after 24h incubation with Au(III) compound 1, co-incubation with CuCl2 but not with cimetidine, showed a significant higher toxicity in A2780cisR cells, whereas in A2780 cells no effect was seen of either CuCl2 or cimetidine. However, after 72h, co-incubation of Au(III) compound 1 with either cimetidine or CuCl2 showed a higher toxicity. After 72h incubation, co-incubation of Au(III) compound 1 with CuCl2 resulted in an increase in toxicity and increase in Au and Cu content in both cell lines as demonstrated by ICP-MS. Based on these results two explanations can be suggested for these effects: Au(III) compound 1 is a substrate for ATP7A/B and inhibition of the efflux transporter ATP7A/B by Cu ions results in a higher Au accumulation and higher toxicity, and/or the increased toxicity is a result of additive or synergistic toxicity of Au and Cu accumulation. A direct involvement of the CTR1 or OCT2 in the uptake of the drug could not be shown by inhibition by CuCl2 or cimetidine. The increased toxicity of Au(III) compound 1 in the presence of cimetidine after 72 h of incubation, but not after 24h, is difficult to explain, as no concomitant increased accumulation of Au was observed by ICP-MS. Studies with other, more selective, inhibitors for each single drug transporter are needed 	 217 to confirm these results. Au(III) compound 1 localizes mainly in the cytoplasm in A2780 cells (in the presence or not of CuCl2 and cimetidine) as demonstrated by fluorescence microscopy.  Similarly to Au(III) compound 1, 24h incubation of cisplatin with CuCl2 shows a significant decrease in viability in A2780cisR cells, although not as strong as for Au(III) compound 1. After 72h, co-incubation of cisplatin with either cimetidine or CuCl2 leads to an increase in toxicity, but no increase in Pt content could be observed in both cell lines as demonstrated by ICP-MS. This result, together with the evidence for increased Cu content in A2780 cells, leads to the hypothesis that copper accumulation is the reason for the increased toxicity in this cell lines. A direct involvement of OCTs or CTR1 in the uptake of cisplatin could not be concluded from our results. If this would be the case then inhibition by cimetidine should have resulted in lower toxicity and lower accumulation in the cells. Holzer et al. showed that, as a consequence of cisplatin incubation, the CTR1 transporter might be sequestered away from the membrane, leading to a decreased Pt uptake into cells. However, a decreased Pt uptake was not confirmed by our results.20 Similarly, a direct involvement of MATE or ATP7A/B in the accumulation of cisplatin could not be confirmed. Concerning passive vs. active uptake for Au(III) compound 1 and cisplatin within the first 120 min of incubation, very preliminary results suggest that active mechanisms might play a role for Au(III) compound 1 uptake into A2780 cells, whereas in A2780cisR this might not be the case. For cisplatin, no difference between 37°C and 4°C incubated cells could be observed, leading to the assumption, that active transport mechanisms might not play a crucial role in these cell lines.   5. Experimental section Synthesis General Remarks All reactions were carried out under an atmosphere of purified argon using Schlenk techniques. Solvents were dried and distilled under argon before use. The precursor [Au(pyb-H)Cl2]11 has been synthesized according to literature procedure. All other reagents were commercially available and used as received. All the analyses were performed at the “Plateforme d’Analyses Chimiques et de Synthèse Moléculaire de l’Université de Bourgogne”. The identity and purity (≥ 95%) of the complexes were unambiguously established using high-resolution mass spectrometry and NMR. Exact mass of the synthesized complexes were obtained on a Thermo LTQ Orbitrap XL. 1H- (300.13, 500.13 or 600.23 MHz), 13C- (125.77 or 150.90 MHz) and 31P- (121.49, 202.45 or 242.94 MHz) NMR spectra were recorded on Bruker 300 Avance III, 500 Avance III or 600 Avance II spectrometers. Chemical shifts are quoted in ppm (δ) relative to TMS (1H and 13C) using the residual protonated solvent (1H) or the deuterated solvent (13C) as internal standards. 85% H3PO4 (31P) was used as an external standard. Infrared spectra were recorded on a Bruker Vector 22 FT-IR spectrophotometer (Golden Gate ATR) and far infrared spectra were recorded on a Bruker Vertex 70v FT-IR spectrophotometer (Diamant ATR). X-ray diffraction data for the Au(III) compound 1 were collected on a Bruker Nonius Kappa CCD APEX II at 115 K.  Au(III) compound 1, [Au(pyb-H)(PPh2Archrom)Cl].PF6 A round-bottom flask was charged with the precursor [Au(pyb-H)Cl2]   (50 mg, 0.115 mmol), KPF6 (106 mg, 0.573 mmol, 5 eq.) and 3-[4-(diphenylphosphino)phenyl]-7-methoxy-2H-chromen-2-one (PPh2Ar) (50 mg, 0.115 mmol, 1 eq.) in suspension into 5 mL of distilled acetone under argon atmosphere. Starting Au complex was solubilized after some minutes. The reaction was maintained at room temperature for 1.5 h; afterward 10 mL of dichloromethane were added and the yellow solution was filtrated through Celite® and concentrated under vacuum. The pure product was obtained after recrystallization from a dichloromethane/pentane mixture as a yellow powder (91 mg, 80 % yield).  1H NMR (Acetone-d6, 500.13 MHz, 298 K): 3.97 (s, 3 H, OCH3), 4.49 (d, 1 H, 2JH-H = 15.6 Hz, CH2-PyrBz), 5.05 (d, 1 H, 2JH-H = 15.6 Hz, CH2-PyrBz), 6.52 (dt, 1 H, 3JH-H = 8.5 Hz, 4JH-H = 1.5 Hz, H5’), 6.88 (dd, 1 H, 3JH-H = 7.5 Hz, 4JH-H = 3.0 Hz, H6’), 6.98 (d, 1 H, 4JH-H = 2.5 Hz, HD), 7.00 (dd, 3JH-H = 8.5 Hz, 4JH-H = 2.5 Hz, HC), 7.03 (dt, 3JH-H = 8.5 Hz, 4JH-H = 0.5 Hz, H4’), 7.33 (dd, 1 H, 3JH-H = 8.5 Hz, 4JH-H = 1.5 Hz, H3’), 7.60-7.66 (m, 4 H, Hortho-Ph), 7.70 (d, 3JH-H = 8.5 Hz, HB), 7.73-7.79 (m, 2 H, Hortho-pC6H4), 	218 7.81 (t, 3JH-H = 8.5 Hz, H5), 7.89-8.01 (m, 8 H, Hmeta/para-Ph + Hmeta-pC6H4), 8.06 (d, 1 H, 3JH-H = 8.5 Hz, H3), 8.27 (s, 1 H, HA), 8.31 (dt, 1 H, 3JH-H = 8.5 Hz, 4JH-H = 1.5 Hz, H4), 9.25 (broad s, 1 H, H6).  13C(1H) NMR (Acetone-d6, 125.77 MHz, 300 K): 47.9 (s, CH2-PyrBz), 56.5 (s, O -CH3), 101.2 (s, CHD), 113.8 (s, CHC), 114.0 (s, Cquat-coum), 122.9 (s, Cquat-p-C6H4), 124.0 (d, 1JP-C = 84.3 Hz, Cipso-Ph), 124.5 (d, 1JP-C = 83.0 Hz, Cipso-Ph), 124.8 (d, 1JP-C = 70.4 Hz, Cipso-p-C6H4),   125.4 (d, 4JP-C = 3.8 Hz, CH5), 127.3 (d, 4JP-C = 3.8 Hz, CH3), 128.8 (s, CH4’), 128.9 (d, 4JP-C = 2.5 Hz, CH5’),129.8 (s, CHpara-Ph), 129.9 (s, CH3’), 130.2 (d, 2JP-C = 10.1 Hz, CHortho-Ph), 130.3 (d, 2JP-C = 10.1 Hz, CHortho-Ph), 131.0 (s, CHB), 133.7 (d, 3JP-C = 7.5 Hz, CH6’), 134.5 (d, 3JP-C = 2.5 Hz, CHortho-p-C6H4), 134.6 (s + d, 3JP-C = 2.5 Hz, Cquat-Bz + CHortho-p-C6H4), 136.1 (s, CHmeta-Ph), 136.2 (s, CHmeta-Ph + CHmeta-p-C6H4), 136.4 (s, CHmeta-p-C6H4), 141.6 (d, 2JP-C = 2.5 Hz, C-Au), 143.1 (s, CHA), 144.2 (s, CH4), 150.8 (s, Cquat-coum), 152.4 (s, CH6), 156.8 (s, Cquat-pyr), 157.9 (s, Cquat-coum), 160.4 (s, Cquat-coum), 164.6 (s, Cquat-coum).   31P(1H) NMR (Acetone-d6, 202.45 MHz, 300 K): 31.5 (s, 1 P, PPh3-Coum), -144.2 (h, 1 P, PF6).  ESI-MS (DMSO-MeOH), positive mode exact mass for [C40H31NO3PAuCl]+ (836.13901): measured m/z 836.13656 [M-PF6]+.  IR (ATR & FIR, cm-1): 1725, 1613, 1569, 1437, 1362, 1025, 836, 751, 311, 229. Anal. Calc. for C40H31NO3P2F6AuCl: C, 48.92, H, 3.18, N, 1.43 %. Found: C, 48.40, H, 2.70, N, 1.52 %.   Cell experiments Cell viability assay The human breast cancer cell line MCF7, human lung cancer cell line A549, human colon cancer cell lines HCT116 p53+/+ and human ovarian cancer cell lines A2780 and A2780cisR (obtained from the European Centre of Cell Cultures ECACC, Salisbury, UK) were cultured in DMEM (Dulbecco’s Modified Eagle Medium) and the A2780 cells in RPMI containing GlutaMax supplemented with 10 % FBS and 1 % penicillin/streptomycin (all from Invitrogen), at 37°C in a humidified atmosphere of 95 % of air and 5 % CO2 in an incubator i (Heraeus, Germany). Non-tumoral human embryonic kidney cells HEK-293T were kindly provided by Dr. Maria Pia Rigobello (CNRS, Padova, Italy) and were cultivated in DMEM medium, added with GlutaMax (containing 10 % FBS and 1 % penicillin/streptomycin (all from Invitrogen) and incubated at 37°C and 5 % CO2. For evaluation of toxicity, cells were seeded in 96-well plates (Costar, Integra Biosciences, Cambridge, MA) at a concentration of 10.000 cells/well (A2780, MCF-7, HEK-293T) or 6000 cells/well (HCT116 p53 +/+, A549) and grown for 24 h in the appropriate medium mentioned above. Solutions of the compounds were prepared by diluting a freshly prepared stock solution (10-2 M in DMSO) of the corresponding compound in aqueous media (RPMI or DMEM for the A2780 or A549, MCF-7, HCT116 p53+/+ and HEK-293T). Afterwards, 200 µL of these dilutions of the compounds were added to the wells to obtain a final concentration ranging from 0 to 120 µM, and the cells were incubated for 72 h. Following 24 or 72 h drug exposure, 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT) was added to the cells at a final concentration of 0.5 mg ml-1 and incubated for 2 h, then the culture medium was removed and the violet formazan (artificial chromogenic precipitate of the reduction of tetrazolium salts by dehydrogenases and reductases) dissolved in how much ml? DMSO. The optical density of each well (96-well plates) was quantified in triplicates at 550 nm using a multi-well plate reader, and the percentage of surviving cells was calculated from the ratio of absorbance of treated to untreated cells. The IC50 value was calculated as the concentration reducing the proliferation of the cells by 50 % using which calculation program? and it is presented as a mean (± SE) of at least three independent experiments. ICP-MS The concentrations of platinum or gold were measured by inductively coupled plasma mass spectrometry (ICP-MS) using an Elan 6000 spectrometer (Perkin Elmer Sciex, Concord, ON, Canada) and Micromist Nebulizer and a cyclonic spraychamber (Glas Expansion Pocasset, MA, U.S.). Nebulizer gas, lens voltage and RF power were optimized daily using a 10 µg/L Pt standard in 0,1% (V/V) HCl with 0.65% (V) HNO3, normal settings were: Rf power 1350 W, nebuliser gas flow rate 0.95 L min−1 , and lens volgate 10 V. Acquisition parameters were 50 ms dwell time, 1 sweeps reading, 5 	 219 reading replicates and 25 readings monitoring all or a subset of 195Pt+, 196Pt+, 197Au+, and 63Cu+The instrument was tuned at the beginning of each analysis to ensure optimal operation.  Samples were dried by vacuum centrifugation (eppendorf, concentrator plus) at 60°C for 2 hours. Cell samples were digested by 200 µL 65% HNO3 and 50 µL 30% H2O2 overnight until the solution is clear. Samples were prepared for analysis by dilution to 5 mL with 0.65% (V/V) HNO3 and 0,1% (V/V) HCl. The external standards were prepared from a 10 mg/L Au andPt stock solution (CPI International, Peak Performance, 4400-120213WG01, Lot# 12B214) or Cu stock solution (Plasma CAL, SCL SCIENCE, Q.C. no 4, Cat# 140-102-045, Lot SC5322198) with 0.65% (V/V) HNO3 and 0,1% (V/V) HCl. All reagents were of the highest available purity.  Cell culture  Human ovarian sensitive (A2780) and cisplatin resistant (A2780cisR) cells were grown in 75 cm2 culture flasks (CellStar, Grenier Bio, Germany) in Roswell Park Memorial Institute (RPMI) 1640 medium supplemented with 10% fetal bovine serum (FBS), 1% penicillin/streptomycin and 2 mM L148 glutamine. Both cell lines were kept at 37°C, 5% CO2, and 100% humidity. Cells were passed on every 3-4 days using 0.25% trypsin in phosphate-buffered saline (PBS, pH 7.4). To maintain resistance, cisplatin-resistant A2780cisR cells were treated with 1 μM cisplatin between every third passage.     Au/ Pt content after 24h/72h incubation with metal compound +/- inhibitor A2780 or A2780cisR cells were plated into 25cm2 culture flasks (CellStar, Grenier Bio, Germany) with RPMI 1640 medium up to a volume of 5 mL per flask. The next day, the cells were treated with 15 µM/ 20 µM cisplatin for 24 hours and 5 µM/ 10 µM cisplatin for 72 hours +/- cimetidine or +/- CuCl2. Cimetidine was used in a nontoxic concentration of 300 µM and CuCl2 was used at 30 µM. The same set-up is used for Au(III) compound 1, but with 5 µM/ 10 µM for 24 hours incubation and 3 µM/ 6 µM for 72 hours incubation. After 24 hours or 72 hours, the medium was removed and the cells were washed two times with 3 mL ice-cold PBS. 500 µL lysis buffer was added and the samples were stored on ice for 20 minutes. Afterwards, cells were detached by using a scrubber policemen and the cell suspension was transferred to a 1.5 mL eppendorf tube. 200 µL of the sample was transferred to a new eppendorf tube for ICP-MS analysis. 5 µL of cell suspension was used for protein determination.  Au/ Pt accumulation during 120 min  incubation with metal compound +/- inhibitor A2780 or A2780cisR cells were plated into 25 cm2 culture flasks (CellStar, Grenier Bio, Germany) with RPMI 1640 medium up to a volume of 5 mL per flask. The next day, the cells were treated with 5 µM cisplatin (+/- cimetidine or +/- CuCl2) for 10’, 20’, 30’, 60’ and 120’ min. For compound 1, we used a concentration of 3 µM. The samples were processed as described above. Protein assay Protein determination was performed using BioRad DCTM Protein Assay reagents A, B and S (Bio-Rad, ).  Using a 96-well-plate, 5 µL of standard (0, 5, 10, 15, 20, 25 and 30 mg/ml bovine serum albumin) and samples were analyzed in duplicate. First, 25.5 µL reagent-mix A+S (25 µL A + 0.5 µL S) was added and then 200 µL reagent B. The absorbance at 610 nm was measured after 15 minutes (stored in dark) using a PlateReader (Perkin Elmer, EnVision, 2104 Multilabel Reader).  Fluorescence Microscopy A2780 cells were seeded (5 x 105 for each sample) and grown on 8 well microscope plates, coated with Poly-L-lysine hydrobromide (Sigma-Aldrich, P6516) with RPMI medium. After 24h, cells were incubated with various concentrations of Au(III) compound 1 in RPMI, without FCS for 1h at 37 °C. At the end of incubation, cells were rapidly washed with cold 	220 PBS and then fixed with 2 % paraformaldehyde for 30 min at 4°C. For visualization of the nuclei with PI, cells were permeabilized with 0.2% Triton X-100 for 20 min at 4°C and treated with 1 µg/µl of PI for 10 min at room temperature. Cells were washed once with PBS and then analyzed by confocal microscopy. As preparation for visualization, the plate wells were removed from the slide and glycerol was used to cover the slide with a glass cover slip. The fluorescence was analysed using a Leica DM4000 B Automated Upright Microscope, equipped with the appropriate filters. PI was excited at 547 nm (emission wavelength 572 nm) and compound 1 at 358 nm (emission wavelength 461 nm, DAPI filter). The acquired images were obtained using the individual filters and a combined image, overlaying the fluorescence was acquired using the Leica microscope software.   6. References  (1) Desoize, B.; Madoulet, C. Critical Rev. Oncol. Hematol. 2002, 42, 317-325. (2) Allardyce, C. S.; Dyson, P. J. Dalton Trans. 2016, 45, 3201-3209. (3) Bertrand, B.; Casini, A. Dalton Trans. 2014, 43, 4209-4219. (4) Ott, I. Coord. Chem. Rev. 2009, 253, 1670-1681. (5) Nobili, S.; Mini, E.; Landini, I.; Gabbiani, C.; Casini, A.; Messori, L. Med. Res. Rev. 2010, 30, 550-580. (6) Kaps, L.; Biersack, B.; Müller-Bunz, H.; Mahal, K.; Münzner, J.; Tacke, M.; Mueller, T.; Schobert, R. J. Inorg. Biochem. 2012, 106, 52-58. (7) Spreckelmeyer, S.; Orvig, C.; Casini, A. Molecules 2014, 19, 15584-15610. (8) Fletcher, J. I.; Haber, M.; Henderson, M. J.; Norris, M. D. Nat. Rev. Cancer 2010, 10, 147-156. (9) Russel, F. G. M. In Enzyme- and Transporter-Based Drug-Drug Interactions: Progress and Future Challenges, Pang, K. S.; Rodrigues, A. D.; Peter, R. M., Eds.; Springer New York: New York, NY, 2010, pp 27-49. (10) Bertrand, B.; Spreckelmeyer, S.; Bodio, E.; Cocco, F.; Picquet, M.; Richard, P.; Le Gendre, P.; Orvig, C.; Cinellu, M. A.; Casini, A. Dalton Trans. 2015, 44, 11911-11918. (11) Cinellu, M. A.; Zucca, A.; Stoccoro, S.; Minghetti, G.; Manassero, M.; Sansoni, M. Dalton Trans. 1995, 2865-2872. (12) Hanthorn, J. J.; Haidasz, E.; Gebhardt, P.; Pratt, D. A. Chem. Commun. 2012, 48, 10141-10143. (13) Ali, M.; Dondaine, L.; Adolle, A.; Sampaio, C.; Chotard, F.; Richard, P.; Denat, F.; Bettaieb, A.; Le Gendre, P.; Laurens, V.; Goze, C.; Paul, C.; Bodio, E. J. Med. Chem. 2015, 58, 4521-4528. (14) Li, J.; Wood, W. H., III; Becker, K. G.; Weeraratna, A. T.; Morin, P. J. Oncogene 2006, 26, 2860-2872. (15) Sorensen, B. H.; Dam, C. S.; Sturup, S.; Lambert, I. H. J. Inorg. Biochem. 2016, 160, 287-295. (16) Nipper, M. E.; Majd, S.; Mayer, M.; Lee, J. C. M.; Theodorakis, E. A.; Haidekker, M. A. Biochimica et Biophysica Acta (BBA) - Biomembranes 2008, 1778, 1148-1153. (17) Lee, J.; Pena, M. M.; Nose, Y.; Thiele, D. J. J. Biol. Chem. 2002, 277, 4380-4387. (18) Hilário-Souza, E.; Valverde, R. H. F.; Britto-Borges, T.; Vieyra, A.; Lowe, J. Internat. J. Biochem. Cell Biol. 2011, 43, 358-362. (19) Liang, Z. D.; Long, Y.; Chen, H. H. W.; Savaraj, N.; Kuo, M. T. J. Biol. Inorg. Chem. 2014, 19, 17-27. (20) Holzer, A. K.; Howell, S. B. Cancer Res. 2006, 66, 10944-10952.         

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