Organometallic ruthenium(II)-arene complexes with triphenylphosphine amino acid bioconjugates: Synthesis, characterization and biological properties

Article abstract of DOI:10.1016/j.bioorg.2019.03.048

(p-Cymene)-ruthenium bioconjugates ML (1) and ML₂ (2), bearing phosphane ligands substituted with chiral or non-chiral amino acid esters, L, were synthetized and characterized by instrumental methods (NMR, CD, MS) and DFT calculations (using the wB97xD functional). Cytotoxic activity of complexes 1 and 2 was investigated by using human cervical carcinoma cell line (HeLa) and MTT assay. Four (2_pG, 2_pA, 2_mG and 2_mA) out of ten synthesized ruthenium complexes showed significant toxicity, with IC₅₀ values of 5–30 μM. Evaluation of the potential biomolecular targets of bioconjugates 2 by UV–Vis, fluorescence and CD spectroscopy revealed no measurable interaction with DNA, but micromolar affinity for proteins. The cytotoxicity of bioconjugates 2 is in correlation with their BSA binding constants, i. e. bioconjugates with lower IC₅₀ values show higher binding affinities towards BSA. Compound 2_mG with value of IC₅₀ 16 μM was selected for further biological characterization. The higher level of toxicity towards tumor compared to normal cell lines indicates its selective activity, important characteristic for potential medical use. It was detected 2_mG caused increase of cells in the S phase of cell cycle and consequential decrease of cells in G0/G1 phase. Additionally, 2_mG caused dose- and time-dependent increase of SubG0/G1 cell population, suggesting its ability to induce programmed cell death. Further investigation determined autophagy as the mode of cell death. The role of GSH in HeLa cells response to investigated organometallic ruthenium complexes was confirmed using specific regulators of GSH synthesis, buthionine sulfoximine and N-acetyl-cysteine. Pre-treatment of cells with ethacrynic acid and probenecid emphasized the role of GSH in detoxification of 2_mG compound. The amount of total ruthenium accumulation in the cell did not correlate with toxicity of 2_pG, 2_pA, 2_mG and 2_mA suggesting structure dependent differences in either cell uptake or kinetics of ruthenium complexes detoxification. We speculate that ruthenium complexes bind protein-based biomolecules further triggering cell death. Based on the gained knowledge, the synthesis and development of more tumor-specific ruthenium-based complexes as potential anticancer drugs can be expected.

Full text of DOI:10.1016/j.bioorg.2019.03.048

Accepted Manuscript
Organometallic ruthenium(II)-arene complexes with triphenylphosphine amino
acid bioconjugates: Synthesis, characterization and biological properties
Margareta Pernar, Zoran Kokan, Juran Kralj, Zoran Glasovac, Lidija-Marija
Tumir, Ivo Piantanida, Domagoj Eljuga, Iztok Turel, Anamaria Brozovic,
Srećko I. Kirin
PII:
S0045-2068(18)30960-X
https://doi.org/10.1016/j.bioorg.2019.03.048
YBIOO 2874
DOI:
Reference:
Bioorganic Chemistry
To appear in:
Received Date:
Revised Date:
Accepted Date:
30 August 2018
5 March 2019
15 March 2019
Please cite this article as: M. Pernar, Z. Kokan, J. Kralj, Z. Glasovac, L-M. Tumir, I. Piantanida, D. Eljuga, I. Turel,
A. Brozovic, S.I. Kirin, Organometallic ruthenium(II)-arene complexes with triphenylphosphine amino acid
bioconjugates: Synthesis, characterization and biological properties, Bioorganic Chemistry (2019), doi: https://
doi.org/10.1016/j.bioorg.2019.03.048
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Organometallic ruthenium(II)-arene complexes with
triphenylphosphine amino acid bioconjugates:
Synthesis, characterization and biological properties
Margareta Pernar,a,# Zoran Kokan,b,# Juran Kralj,a Zoran Glasovac,c Lidija-Marija Tumir,c Ivo
Piantanida,c Domagoj Eljuga,d Iztok Turel,e Anamaria Brozovic,a,* Srećko I. Kirin,b,*
a
Division of Molecular Biology, Ruđer Bošković Institute, Bijenička cesta 54, HR-10000
Zagreb, Croatia
b
Division of Materials Chemistry, Ruđer Bošković Institute, Bijenička cesta 54, HR-10000
Zagreb, Croatia
c Division of Organic Chemistry and Biochemistry, Ruđer Bošković Institute, Bijenička cesta
54, HR-10000 Zagreb, Croatia
d
Department for Oncoplastic and Reconstructive Surgery, University Hospital for Tumors,
University Clinical Hospital Centre Sisters of Mercy, HR-10000 Zagreb, Croatia
e
Faculty of Chemistry and Chemical Technology, University of Ljubljana, Večna pot 113,
SLO-1000 Ljubljana, Slovenia
# equal contribution
*corresponding authors
E-mail: Anamaria.Brozovic@irb.hr, Srecko.Kirin@irb.hr
Key words: Ruthenium; p-Cymene; Phosphine ligands; Cytotoxicity; Cell death

Abstract.
(p-Cymene)-ruthenium bioconjugates ML (1) and ML2 (2), bearing phosphane ligands
substituted with chiral or non-chiral amino acid esters, L, were synthetized and characterized
by instrumental methods (NMR, CD, MS) and DFT calculations (using the wB97xD
functional). Cytotoxic activity of complexes 1 and 2 was investigated by using human
cervical carcinoma cell line (HeLa) and MTT assay. Four (2pG, 2pA, 2mG and 2mA) out of ten
synthesized ruthenium complexes showed significant toxicity, with IC50 values of 5 to 30
μM. Evaluation of the potential biomolecular targets of bioconjugates 2 by UV-Vis,
fluorescence and CD spectroscopy revealed no measurable interaction with DNA, but
micromolar affinity for proteins. The cytotoxicity of bioconjugates 2 is in correlation with
their BSA binding constants, i. e. bioconjugates with lower IC50 values show higher binding
affinities towards BSA. Compound 2mG with value of IC50 16 μM was selected for further
biological characterization. The higher level of toxicity towards tumor compared to normal
cell lines indicates its selective activity, important characteristic for potential medical use. It
was detected 2mG caused increase of cells in the S phase of cell cycle and consequential
decrease of cells in G0/G1 phase. Additionally, 2mG caused dose- and time-dependent
increase of SubG0/G1 cell population, suggesting its ability to induce programmed cell death.
Further investigation determined autophagy as the mode of cell death. The role of GSH in
HeLa cells response to investigated organometallic ruthenium complexes was confirmed
using specific regulators of GSH synthesis, buthionine sulfoximine and N-acetyl-cysteine.
Pre-treatment of cells with ethacrynic acid and probenecid emphasized the role of GSH in
detoxification of 2mG compound. The amount of total ruthenium accumulation in the cell did
not correlate with toxicity of 2pG, 2pA, 2mG and 2mA, suggesting structure dependent
differences in either cell uptake or kinetics of ruthenium complexes detoxification. We
speculate that ruthenium complexes bind protein-based biomolecules further triggering cell
death. Based on the gained knowledge, the synthesis and development of more tumor-specific
ruthenium-based complexes as potential anticancer drugs can be expected.

Introduction.
Besides the importance of ruthenium (Ru) complexes in catalysis, a major area of research
interest is their medical implementation. Although frequently in use, the platinum-based
compounds, as an example of prominent inorganic anticancer complexes, have several
limitations such as development of drug resistance, side effects and ineffectiveness towards
certain tumors (Brozovic and Osmak 2007, Galluzzi et al 2012). In recent years, ruthenium
complexes displayed some superior properties over platinum-based drugs (Bergamo and Sava
2015, Vajs et al 2015). Despite the fact that none of the ruthenium complexes is still in
clinical use as anticancer drug, prominent representatives like NAMI-A, KP1019, RAPTA
and TLD1433 scored success in in vitro and in vivo studies, prompting ruthenium complexes
to rapidly become a major area in anticancer drug innovation (Bergamo and Sava 2015,
Blunden et al 2014, Domotor et al 2013, Meier-Menches et al 2018, Sersen et al 2015). The
wide variety of ruthenium anticancer complexes studied in recent years includes derivatives
with amino acids (Rathgeb et al 2014, Scrase et al 2015) and derivatives with
triphenylphosphines (Biancalana et al 2017, Chaplin et al 2007, Martinez et al 2017, Millan
2019).
Nowadays, a number of anticancer metal complexes are in the developmental phase and are
mostly designed to mimic broadly used cisplatin, known to target DNA. Aside from the DNA
damage, cisplatin is able to induce reactive oxygen species (ROS) and endoplasmic reticulum
stress, as well as bind to peptides and proteins (Brozovic et al 2010). Similarly, recent studies
examining ruthenium anticancer compounds revealed that DNA is not always the primary
target and that these compounds bind proteins stronger than to DNA (Gasser et al 2011,
Novakova et al 2005, Ravera et al 2004, Scolaro et al 2007). Based on those findings,
different modes of ruthenium compounds cytotoxic actions occur, although the exact
mechanism is still not known. What is known is the fact that cancer cells are generally
growing and multiplying much faster than normal healthy cells, creating a reductive
environment due to the raised metabolic rate, higher levels of glutathione (GSH) and a lower
pH (Antonarakis and Emadi 2010). Moreover, it was shown that ruthenium complexes
interact with GSH (Wang et al 2005) and GSH-associated enzymes (Lin et al 2013), implying
the role of the GSH in cells’ defense against toxic damage (Sommer et al 2014).
Amino acid bioconjugates of triphenlyphosphanes are well known ligands for the
coordination of transition metals. We have studied the self-assembly and biological properties
of their palladium and platinum complexes (Kokan et al 2017, Kokan et al 2018) and
successfully used them as monodentate ligands in rhodium catalyzed asymmetric
hydrogenations (Kokan and Kirin 2012, Kokan and Kirin 2013, Kokan et al 2014, Opačak et
al 2019). A major feature of those metal complexes is the formation of chiral secondary
structures responsible for chiral induction in the catalytic cycle. Herein, we explore the
synthesis and characterization of (p-cymene)ruthenium complexes 1 and 2, bearing
phosphane ligands L with chiral or non-chiral amino acid esters. The spacer between the
phosphorus and the amino acid was varied in order to investigate hydrogen-bonding
propensity of the amino acid substituents. Structural and biological properties of the prepared
complexes will be reported.
Results and discussion.

Synthesis and characterization. The chemical synthesis of the presented organometallic
complexes was performed in solution in several steps, Scheme 1. First, the free carboxylic
acid of the phosphine precursors Ph2P-pC6H4-CO2H, Ph2P-mC6H4-CO2H or Ph2P-C2H4-
CO2H was reacted with N-unprotected amino acid methyl esters of achiral glycine or chiral
L-alanine in standard conditions for amide bond formation to yield amino acid
triphenylphosphine ligand bioconjugates L, Scheme 1. In the second step, ligand
bioconjugates L and ruthenium precursor [{(pCym)RuCl}2Cl2] were combined to give
mono-complexes 1, with 1:1 metal to ligand stoichiometry, general formula
[(pCym)RuLCl2]. Finally, in the last synthetic step, mono-complexes react with one
additional equivalent of the corresponding ligand and the bis-complexes 2 were obtained,
[(pCym)RuL2Cl](PF6), with 1:2 metal to ligand stoichiometry. In total, six ligands and ten
complexes were prepared. Unfortunately, complexes 2aG and 2aA, although clearly present in
the crude reaction mixture, could not be isolated pure enough for further studies.
O
O
O
O
O
O
O
O
Ru
Cl
Ru
Cl
OH
O
HN
O
HN
O
NH
O
HN
O
(a)
(b)
(c)
Cl
P
P
P
R1
R1
R1
R2
R2
R2
P
P
R1
R2
R1
PF6
LpG; R1 = p-Ph, R2 = H
LpA; R1 = p-Ph, R2 = Me
LmG; R1 = m-Ph, R2 = H
LmA; R1 = m-Ph, R2 = Me
LaG; R1 = CH2CH2, R2 = H
1pG; R1 = p-Ph, R2 = H
1pA; R1 = p-Ph, R2 = Me
1mG; R1 = m-Ph, R2 = H
1mA; R1 = m-Ph, R2 = Me
1aG; R1 = CH2CH2, R2 = H
1aA; R1 = CH2CH2, R2 = Me
2pG; R1 = p-Ph, R2 = H
2pA; R1 = p-Ph, R2 = Me
2mG; R1 = m-Ph, R2 = H
2mA; R1 = m-Ph, R2 = Me
LaA; R1 = CH2CH2, R2 = Me
Scheme 1, experimental conditions: (a) TBTU/HOBT/DIPEA/DCM, amino acid ester
hydrochloride, RT, 15 h; (b) Di-µ-chlorobis[(p-cymene)chlororuthenium(II)]/DCM, RT, 2 h;
(c) 1. NH4PF6/AcCN, reflux, 45 min, 2. L/DCM, RT, 15 h.
13
31
The ligands and complexes were characterized by NMR (1H, C and P), CD spectroscopy
and MALDI high resolution mass spectrometry. In particular, the 31P shifts of the phosphine
moiety in ligands L (δP ≈ -5 ppm), mono- and bis-complexes ML (δP ≈ 25 ppm) and ML2 (δp
≈ 22 ppm) clearly indicate binding to the ruthenium metal in the different stoichiometry. The
NH proton chemical shifts for L, ML and ML2 indicate only weak intramolecular hydrogen
bonding in ML2 complexes (δNH < 7.15 ppm in CDCl3). It is interesting to note that the C2
proton in 1mG and 1mA are unexpectedly highly deshielded (δH ≈ 8.9 ppm). Selected
characteristic NMR data are collected in Table S1.
Computational study. Relative Gibbs energies of the ruthenium complexes were calculated
using wB97xD density functional approach, Table S2. This functional was confirmed as very
good in rationalizing enantioselectivity in a rhodium catalyzed hydrogenation reaction and in
predicting geometries of organic molecules and transition metal complexes including
ruthenium complexes (Gridnev and Imamoto 2015, Minenkov et al 2012). Further, this
functional is the recommended method when weak interactions like π-π interactions are
expected (Qi et al 2016). Four complex cations with 2:1 ligand to metal stoichiometry, 2mG
,
2pG, 2mA and 2pA, were chosen for the calculations, containing meta- or para-substituted
triphenylphosphine ligand conjugated with glycine or alanine methyl ester, respectively and
several conformations were considered for each of the cations. Relative energies of all
optimized conformers of the ruthenium complex cations are collected in Table S2
(supplement). In addition, Cartesian coordinates of all computed structures are available as a
single separate text file, for visualization and analysis.

All calculated conformations contain aromatic stacking interactions ( –  stacking) between
two different triphenylphosphine moieties, but with a different number of intramolecular
hydrogen bonds between the ligands. In the case of 2mG, a representative conformation was
generated with the aid of NMR experiments. NOESY experiment suggests position of amide
protons in close proximity to the C2 and C4 atoms of the stacked phenylene subunit and also
to the cymene ligand. This information indicates a "syn-Phe" orientation of two 1,3-
phenylene subunits forming a curved shape of the complex. Chloride ligand is directed
outward the structure of complex.
Figure 1. Optimized structures of 2mGs-I and 2mAs-Ip.
In context of intramolecular H-bonds between the ligands, both "syn-Phe" (2mGs) and "anti-
Phe" (2mGa) conformers can exist in a Herrick (I) or van Staveren (II) arrangement. Several
conformers satisfying a "syn-Phe" arrangement were optimized and the lowest energy one
shows a Herrick-type of hydrogen bonding motif (2mGs-I). During our attempts to optimize
2mGs-II conformer, reorganization of the hydrogen bonds occurred and the optimization
ended up in "semi-Herrick" structure (2mGs-Ib) with only one NH-OCOMe hydrogen bond.
We were able to optimize van Staveren conformer 2mGs-II only with cymene ligand moved
away from the stacked phenylene fragment. However, this structure strongly disagrees with
the NOESY data and it is not likely present in solution. According to DFT calculations, 2mGs
-
I is more stable than 2mGa-I, 2mGs-II and 2mGa-II by 11, 13 and 33 kJ mol-1, respectively. The
lowest stability of conformer 2mGa-II, with only one hydrogen bond, is in good accordance
with our previous results on ferrocene amino acids (Kirin et al 2006). These results
additionally support the structural indications obtained by NOESY experiment.
In the case of 2pG, only one - stacking motif with either Herrick or van Staveren hydrogen
bonding scheme (2pG-I and 2pG-II, respectively) is possible. The calculations indicate the
somewhat lower energy for 2pG-I than for 2pG-II conformers (by ca 6 kJ mol-1, Table S2).
Inspection of the geometrical parameters reveals significantly longer hydrogen bonds in 2pG-
II than in 2pG-I (HB, above 2.0 Å vs. 1.919 Å) indicating origin of the energy difference.
The analogous conformers were also identified for the chiral L-alanine derivatives 2mAs, 2mAa
and 2pA. Only L-alanine derivatives were used for the calculations, to allow comparison with
compounds experimentally prepared herein. The starting structures were generated from the
glycine-based conformations by replacement of one C-proton with methyl group. Again, the
most stable structure is characterized by "syn-Phe" arrangement of the stacked phenylene
subunit with Herrick motif of hydrogen bonding (2mAs-Ip) with P-type of helical chirality.
This result confirms preferred "syn-Phe" arrangement in both aminoacid bioconjugates.

According to calculations, conformer 2mAs-Ip is more stable than 2mAs-Im and 2mAa-Im, by
20 and 18 kJ mol-1, respectively. Calculated lower stability of M- helical isomers of 2mAs-Im
with respect to its P- diastereomer is ascribed to the arrangement of two methyl groups of
alanine fragments. More stable structure (P-helical isomer) has these two methyl groups
relatively close to each other (d = 4.088 Å). Although one could expect pronounced sterical
repulsion, this is apparently not the case. It is somewhat surprising that 2mAs-IIp lies only 8
kJ mol-1 above 2mAs-Ip and it is more stable than 2mAa-Im in spite of reduced number of
hydrogen bonds. The geometry of 2mAs-IIp is similar to 2mGs-II and, if we assume similar
conformation of glycine and alanine derivatives, its presence in the solution is unlikely.
Finally, we shall briefly comment the structure of 2pA complex. The lowest Gibbs energy was
obtained for the Herrick-type complex 2pA-I and it is more stable than 2pA-II by ca 11 kJ
mol-1. Relative stabilities of two diastereomers (defined by P- and M- helicity) in comparison
to the isomer 2pA-II follows the same trend as for 2mAs-I derivative, P-helicity with the
distance between the alanine methyl groups of 4.060 Å is preferred over the M- isomer by 19
kJ mol-1. The latter is even less stable than 2pA-II by 5 kJ mol-1.
Cytotoxicity. The cytotoxicity of ligands L, as well as their ML and ML2 ruthenium
complexes, was examined by MTT assay; ruthenium precursor di-µ-chlorobis[(p-
cymene)chloro-ruthenium(II)] was included for comparison. For this purpose, the well-
known human cervical carcinoma cell line was used (HeLa), attested for screening of new
compounds (Brozovic et al 2014, Cimbora-Zovko et al 2011). The IC50 value (the
cytotoxicity of the compound expressed as the dose that reduced the survival of cells in 72 h
MTT assay to 50% of the value obtained for control cells) was above 33.3 μM for all ligands
L and mono-ligand ruthenium complexes ML (Table S3). The ruthenium precursor was not
toxic for the cells (also Table S3). However, the IC50 for bis-ligand ruthenium complexes
2pG, 2pA, 2mG and 2mA was 5-30 μM, making them interesting for further examination as
potential antitumor compounds (Table 1). The compound 2pG was the least toxic and the 2mA
was the most toxic one. In order to avoid the solvent’s toxicity, concentrations above 33.3
μM were not used. In that way, the highest concentration of the solvent in the well was under
0.5 %, which is not toxic for the HeLa cells.
Table 1. The cytotoxic activity of the compounds towards the HeLa cells (IC50 μM ±
SD), after 72 h incubation. The cytotoxicity was measured by MTT assay. The experiments
were repeated at least three times. Cp. = Compound.
Cp.
2pG
2pA
2mG
2mA
IC50 ± SD (μM)
30±3.2
15±3.7
16±2.5
5±2.7
The IC50 values of 5 to 30 μM measured in HeLa cells for ML2 complexes presented herein
were similar to the value obtained under same experimental conditions for broadly used
chemotherapeutic, cisplatin (Stojanovic et al 2013). Moreover, our results for additional
tumor cell lines investigated are similar to the values previously obtained for breast
carcinoma (MDA-MB-231) cells, treated with ruthenium complexes substituted with amino
acid. The authors concluded that the amino acids do not significantly affect the activity of the

complexes, but existence of the methyl groups in diamine increase their biological activity
(Dos Santos et al 2016). The presence of triphenylphospine group seems to increment the
antitumor properties of Ru complexes through intercalation in the DNA of the human acute
promyelocytic leukemia (HL60) cells with IC50 value 5.2 μM (Saez et al 2014).
One of the criterions necessary for a newly synthesized complex to be considered as a
potential antitumor compound is low toxicity for normal cells and non-selective toxicity for
different tumor cell lines. With the purpose to test this criterion, for further experiments we
decided to use 2mG, due to its medium value of toxicity within the studied series of
compounds. In order to examine the toxic capacity of 2mG on different types of human tumor
cells, the laryngeal carcinoma (HEp2), lung carcinoma (H460) and MDA-MB-231 cells were
used. The collected data showed that 2mG is similarly toxic to all examined cancer cell lines
(Table 2). However, the IC50 of 2mG for normal human cell line, keratinocytes, was far above
the toxicity measured for all examined compounds; the same result was obtained for human
fibroblasts (Table 2).
Table 2. The cytotoxic activity of the 2mG towards different tumor and two normal cell
lines (IC50 μM ± SD), after 72 h incubation. The cytotoxicity was measured by MTT assay.
The experiments were repeated at least three times. Cp. Compound.
MDA-
MB-231
12.3±0.9
Cp.
2mG
HEp2
H460
HeLa
keratinocyte fibroblast
>33.3 >33.3
14±4.7
15±0.6
16±2.5
Potential biomacromolecule target evaluation. For the small bioactive molecules,
determination of the main target within the living cell is a quite challenging and time-
consuming task, particularly for the structures presented herein, that contain DNA-targeting
and protein-targeting features. In general, condensed aryl-Ru complexes are well known
DNA binders (Gasser et al 2011); however, here presented structures do not contain
condensed arenes necessary for DNA intercalation, nor several positive charges which could
interact with DNA-backbone. Nevertheless, attached amino acid residues could form several
H-bonds within DNA minor groove. On the other side, general spherical shape of the studied
molecules is characterized by high hydrophobicity and thus could be nicely accommodated
within hydrophobic pockets of well-known carrier proteins like serum albumins, within
which amino acid residues from the studied small molecules could additionally contribute to
the protein-substrate complex formation by H-bonding.
To facilitate the planning and the analysis of demanding cellular experiments in respect of the
main target choice, we studied interactions of several Ru-complexes with model DNA (calf
thymus, ct-DNA) and model protein: serum albumin (BSA), as the most abundant protein in
blood plasma, that is responsible for the transport of many small molecules (among which are
drugs and probes) (Otagiri et al 2016, Peters et al 1995). Experiments were performed in
simple, well-defined conditions in cuvettes, at biologically relevant conditions (buffered pH
7) containing only studied Ru-complex (micromolar concentration) and biomacromolecule
(ct-DNA or BSA).
The studied Ru-complexes did not stabilize ct-DNA against thermal denaturation nor
changed the DNA CD spectrum, thus showing no measurable interaction (Figure S4).
However, addition of Ru-complexes induced measurable changes in the BSA CD spectrum at

240 – 290 nm (Figure S5), suggesting structural change in the protein chiral conformation;
non-linear dependence of the CD change agreed well with a non-covalent binding event.
Furthermore, the intrinsic fluorescence of BSA significantly quenched upon Ru-complex
addition. Multivariate analysis of the fluorimetric data by Specfit program (Gampp et al
1985, Maeder and Zuberbuhler 1990) revealed formation of 1:1 stoichiometry Ru-
complex/BSA complex with micromolar affinity (Figures S6-S9). Intriguingly, non-linear
dependence of the BSA CD spectrum changes at 260 nm (Figures S6-S9, Table 3) also
supported similar binding affinity. Thus, results obtained suggest that the observed cytotoxic
activity of compounds is not related to any interactions of complexes with the cellular DNA,
but more likely with the protein target.
Table 3. Stability constants (log Ks)a and spectroscopic properties (I)b of the complexes that
ruthenium(II)-arene compounds formed with BSA protein, calculated according to the
fluorimetric titrations (Na-cacodylate buffer, c = 0.05 mol dm-3, pH = 7.0; exc= 300 nm; em=
320-450 nm, c(BSA) = 5  10-7 mol dm-3).
Cp.
2pG
2pA
2mG
2mA
log Ksa
 Ib
-95%
-48%
-57%
-34%
5.08±0.07
5.99±0.07
5.84±0.05
6.50±0.09
a
Processing of titration data by Specfit program (Gampp et al 1985, Maeder and Zuberbuhler 1990) gave best
fit for 1:1 stoichiometry; for all titrations concentration range corresponded to ca. 20-80% complexation.
b Changes in fluorescence of BSA induced by complex formation (I = (Ilim- I0) 100 / I0; where I0 was
calculated emission intensity of free compound and Ilim was emission intensity of a complex calculated by
Specfit program.
A similar protein binding affinity was observed previously for RAPTA-C, the Ru containing
antimetastatic agent, that accumulates at specific histone site on the nucleosome core. It
seems that changing the ethylenediamine ligand of [Ru–cymene-Cl]+ to phosphaadamantane
(+Cl-) switches the adduct formation profile from primary targeting the DNA to targeting the
proteins associated with chromatin (Adhireksan et al 2014).
Interestingly, strong correlation between the cytotoxic activity of compounds towards HeLa
cells (Table 1) and the stability constants of compound-BSA complexes (Table 3) was
observed. Specifically, compound 2mA showed both the highest cytotoxic activity and the
highest affinity for BSA constant, while 2pG showed the lowest cytotoxicity and the lowest
stability constant toward BSA among examined compounds. This correlation additionally
supported protein as one of the main biological targets for ruthenium complexes in this study.
Cell cycle. Further, we were interested to investigate the mechanism of toxicity of bis-ligand
ruthenium complexes and for this purpose we treated HeLa cells with one of them; 16 μM
2mG during 24-72 h. Upon flow cytometric analysis it was shown that 2mG caused a slight
time-dependent increase of cells in S phase of the cell cycle that is more visible in dose-
response of cells to 2mG treatment, along with consequential decrease of cells in G0/G1 phase
(Figure 2A). This was accompanied by the increase of the cells present in the SubG0/G1

fraction, which represent the population of dead cells. The increase of the SubG0/G1 cell
fraction was confirmed by the treatment of HeLa cells during 72 h with different
concentrations of 2mG (Figure 2B). Deconvolution algorithm (Watson pragmatic) was used
for cell cycle data analysis. The results were confirmed by time- and dose-dependent cell
cycle analysis upon treatment of HEp2 cells with 2mG, where increase of the cell amount in S
phase of the cell cycle is more visible compared to the HeLa cells due to their higher
sensitivity to 2mG (Figure S11).
Figure 2. 2mG compound induces increase of HeLa cells in S phase and consequential
decrease of cells in G0/G1 phase of the cell cycle, and an increase of SubG0/G1 cell
population. Cells were treated either with 16 μM 2mG during 24-72 h (A) or with different
concentrations of 2mG during 72 h (B). The data of one from three performed experiments are
shown.
Cell death. Necrosis and apoptosis are two major types of cell death. Inappropriate
regulation of both processes can result in several diseases, including cancer (Nikoletopoulou
et al 2013). With the purpose to distinguish between necrotic and apoptotic cell death, HeLa
cells were treated with different concentrations of 2mG during 24-72 h and then stained with
Annexin V-FITC and propidium iodide (PI). As shown in Figure 3, the percentage of PI
negative (-)/Annexin V-FITC positive (+) and PI+/Annexin V-FITC+ cells increased with the
time (Figure 3A) as well as with the dose (Figure 3B) indicating apoptosis as mode of 2mG
induced cell death. Starved and/or heat shocked cells were always used as compensation
controls for Annexin V-FITC and PI staining for all performed experiments.
One of the most common signaling cascades involved in apoptosis is the activation of a
highly specialized family of cysteinyl-aspartate proteases (caspases), which are usually
present in the inactive zymogen forms. Once activated, caspases initiate cell death by
cleaving and thus activating effector caspases, which drive the process of apoptosis (Li and
Yuan 2008). Caspase mediated apoptotic cell death is accomplished through the cleavage of
several key proteins required for cellular functioning and survival (Fischer et al 2003).
PARP-1 is one of the several known cellular substrates of caspases. Cleavage of PARP-1 by
caspases is considered to be the hallmark of apoptosis (Kaufmann et al 1993, Tewari et al
1995). In order to test our conclusion that 2mG induces apoptosis, formulated based on the
data from Figure 3A and 3B, we treated HeLa cells with 22 μM concentration of the complex

during 24-72 h. Our data showed that 2mG does not induce PARP cleavage, which was
detected in HeLa cells treated 72 h with 5 μM cisplatin (cDDP) used as a positive control
(Figure 3C). Moreover, the pre-treatment of HeLa cells with pan caspase inhibitor that
irreversibly binds to the catalytic site of caspase proteases and can inhibit the induction of
apoptosis, carbobenzoxy-valyl-alanyl-aspartyl-[O-methyl]-fluoromethylketone (Z-VAD), did
not protect cells from 2mG induced cell death (Figure 3D). Additionally, the caspase
independent cell death was confirmed upon specifically measuring caspase 3 and 7 activity in
the HeLa cells treated 48 and 72 h with 22 μM 2mG (Figure S12A), and by two hours pre-
treatment of additional cell system, HEp2 cells, with Z-VAD prior to treatment with 2mG
(Figure S12B). The activity of caspase 3/7 upon 2mG treatment was not detected (Figure
S12A) and Z-VAD did not increase survival of HEp2 cell line upon 2mG treatment (Figure
S12B). The fact that we did not detect cleavage of PARP and that Z-VAD did not prevent
2mG induced cell death brought us to the hypothesis that 2mG probably induces caspase
independent cell death. The pre-treatment of cells with inhibitor of necroptosis, necrostatin-1
(Vandenabeele et al 2013), did not protect cells from death (Figure S13), excluding this type
of cell death from the involvement in the 2mG toxicity.
Figure 3. 2mG compound induces cell death in HeLa cells. Cells were treated either with 10
or 16 μM 2mG for 24-72 h (A) or 10, 16 and 22 μM 2mG for 72 h (B). The cell death was
measured by flow cytometry. The one of three performed experiments for each set of
conditions is shown. Starved (24 h cell growth without serum) or heat shocked (10 min, 56
°C) cells were always used as a compensation Annexin V-FITC/PI controls. PI-/Annexin V-

FITC- cells (lower left quadrant), PI-/Annexin V-FITC+ cells (lower right quadrant),
PI+/Annexin V-FITC+ cells (upper right quadrant). Cleavage of PARP was detected by
Western blot. β-Tubulin was used as a loading control. One out of three performed
experiments is presented. HeLa cells treated with 5 μM cisplatin (cDDP) during 72 h were
used as positive control for PARP cleavage (C). HeLa cells were pre-treated for two hours
with 6.25 μM Z-VAD prior to treatment with different concentrations of 2mG. The cell
survival was measured 72 h after the treatment by MTT assay. The experiments were
repeated at least three times (D).
From the literature is known that cells can enter autophagy prior to dying (Dikic and Elazar
2018, Levine and Kroemer 2008). Autophagy is a homoeostatic cellular process regulating
protein and organelle turnover by lysosomal destruction (Levine and Kroemer 2008). The
treatment of HeLa cells with 22 μM 2mG during 24-72 h induced increase in Beclin-1, which
is accompanied with the increase of proteolytic derivate LC3-II visible upon 72 h treatment
(Figure 4A). This is in line with the fact that Beclin-1 is involved in autophagosome
formation and LC3 is located in the cytosol (LC3-I) or in autophagosomal membranes (LC3-
II). LC3-II may thus be used to estimate the abundance of autophagosomes prior to their
destruction through fusion with lysosomes (Klionsky et al 2012). With the intention to verify
the possible role of autophagy in 2mG induced cell death, the cells were pretreated with
bafilomycin A (BAF A), which is a known inhibitor of the late phase of autophagy (Klionsky
et al 2012), and then treated with different concentrations of 2mG. BAF A decreased the toxic
effect of 2mG (Figure 4B) pointing towards the conclusion that 2mG induces autophagy.
Additional inhibitor of autophagy, 3-methyladenine (3-MA), which is an inhibitor of
phosphatidylinositol 3-kinases (PI3K), decreased the toxicity of 2mG as well (Figure S11).
PI3K plays an important role in many biological processes, including controlling the
activation of the mechanistic target of rapamycin (mTOR), a key regulator of autophagy (Wu
et al 2010). Autophagy as the mode of cell death upon 2mG treatment was confirmed in HEp2
cell line by MTT and SRB assays (Figure S14A and S14B). The obtained data imply the
autophagy as a part of 2mG triggered caspase independent programmed cell death. There are
multiple caspase-dependent and -independent mechanisms by which classical features of
programmed cell death are mediated (Liu et al 2003). Further investigation is needed to
understand 2mG triggered caspase independent programmed cell death and the correlation of
autophagy with it.

Figure 4. 2mG compound induces autophagy in HeLa cells. Cells were treated with 22 μM
2mG. The expression of Beclin-1, LC3-I and LC3-II was detected by Western blot. β-Tubulin
was used as a loading control. Data obtained with dDensitometric analysis are expressed as a
ratio between examined Beclin-1 or LC3-II and β-Tubulin. Non-treated cells were set as 1.0.
One out of three performed experiments for each protein is presented (A). HeLa cells were
pre-treated for two hours either with 6.25 μM Z-VAD or with 0.5 nM BAF A prior to
treatment with different concentrations of 2mG. The cell survival was measured 72 h after the
treatment by MTT assay. The experiments were repeated at least three times. Significance
was determined between the 2mG only and either Z-VAD or BAF A pre-treated and 2mG
treated cells (*, P < 0.05; **, P < 0.01) (B).
The role of glutathione. The fact that cancer cells are generally growing and multiplying
much faster than normal healthy cells and have a rapid production of new biomolecules,
makes them more vulnerable to compounds such as the one investigated. Also, tumor cells
produce a reductive environment due to the increased metabolic rate, feature that opens the
possibility to activate ruthenium complexes solely at the site of cancerous cells (Antonarakis
and Emadi 2010). The tripeptide GSH has the capacity to bind different metal-based drugs
(Brozovic et al 2010). We were interested in the possibility that GSH is involved in
detoxification of investigated compounds due to their capacity to bind proteins (Table 3). For
that purpose, HeLa cells were either pre-treated with specific inhibitor of GSH synthesis,
0.01μg/mL buthionine sulfoximine (BSO) overnight or with precursor of GSH synthesis, 5
mM N-acetylcysteine (NAC) for two hours prior to treatment with 2pG, 2pA, 2mG or 2mA. The
conditions used were tested previously to be effective (Brozovic et al 2008, Brozovic et al
2014). The obtained data showed decreased survival of HeLa cells in the case when GSH was
depleted by BSO and increased survival of cells in the case when the level of GSH was
increased due to the NAC (Figure 5A-D). Similar results were obtained for all examined bis-

ruthenium complexes, showing the importance of GSH in the toxicity of examined
compounds.
Figure 5. Glutathione reduced the toxicity of 2pG, 2pA, 2mG and 2mA. HeLa cells were pre-
treated either with 0.01 μg/mL BSO (overnight) or with 5 mM NAC (2 h) prior to treatment
with different concentrations of 2pG (A), 2pA (B), 2mG (C) or 2mA (D). The cell survival was
measured 72 h after the treatment by MTT assay. The experiments were repeated at least
three times. Significance was determined between the 2mG only and either BSO or NAC pre-
treated and 2pG/2pA/2mG/2mA treated cells (*, P < 0.05; **, P < 0.01).
Since GSH is one of the major endogenous antioxidants, we next measured the formation of
reactive oxygen species (ROS) upon treatment of HeLa cells with 2mG. We were not able to
detect increased formation of ROS (Figure S12A and S12B) by flow cytometry, what is in
line with our prediction that this type of chemical structure does not activate the formation of
ROS. Therefore, we concluded that GSH probably has a role, not as an antioxidant, but rather
as a detoxification mechanism in the cells. This hypothesis was confirmed with pre-treatment
of cells with two antioxidant compounds, tempol and trolox, which did not protect cells from
2mG toxicity (Figures S13A and S13B). The combination treatment of HeLa cells with well
accepted inhibitor of glutathione S-transferase (GST) (a detoxification enzymes that catalyze
the conjugation of GSH to a wide variety of endogenous and exogenous electrophilic
compounds) (Townsend and Tew 2003), 5 or 7.5 μg/mL ethacrynic acid (ETA) (Osmak et al
1998), decreased cell survival compared with cells treated with 2mG only (Figure 6A). The

data implied enzymatically-regulated formation of detoxification complex between GSH and
2mG. Moreover, pre-treatment of HeLa cells with probenecid, the inhibitor of MRP/GS-X
pumps (Zaman et al 1995), decreased survival of cells compared to the ones treated only with
2mG (Figure 6B). These results indicate efflux of the formed conjugates out of the cells and
the detoxification role of GSH.
Figure 6. Glutathione probably forms conjugates with the 2mG. HeLa cells were pre-
treated either with 5 or 7.5 µg/mL ETA (A) or with 0.2 or 0.625 mM probenecid (B) prior to
treatment with different concentrations of 2mG. The cell survival was measured 72 h after by
MTT assay. The experiments were repeated at least three times. Significance was determined
between the 2mG only and either ETA or probenecid pre-treated and 2mG treated cells (*, P <
0.05; **, P < 0.01).
Cellular accumulation. Further we were interested to check the correlation between cellular
accumulations of investigated compounds with their toxicity (Pongratz et al 2004, Puckett et
al 2010). The HeLa cells were treated with 150 μM 2pG, 2pA, 2mG or 2mA during 2-6 h and
analyzed by high-resolution inductively coupled plasma mass spectrometry (HR ICPMS).
Our results showed that the amount of ruthenium in the cells is the highest upon treatment
with 2pA and the lowest upon treatment with 2mG (Figure 7A). The obtained data were further
confirmed by investigation of dose-dependent accumulation of ruthenium for all compounds
(Figure 7B). The data also showed that the amount of ruthenium does not correlate with
toxicity of each compound (Table 1). In a line with that, compound 2mG, used for most of the
testing, was well chosen since it exhibits significant toxicity (Table 1) with the lowest
amount of ruthenium in the cell. The most likely reasons, which need further investigation,
could be different structural features of the compounds, which regulate different cellular
uptake or even different capacity of GSH to detoxify them. The fact that hydrophobicity is an
important pharmacological feature, closely related to the drug uptake and efflux, and the fact
that phenyl group is hydrophobic (Liu et al 2011) could explain why phenyl group in para-
position (2pA) correlates with accumulation and in meta-position (2mA) with toxicity. By
using the same model system, HeLa cells, it was shown that anti-metastatic effect of NAMI-
A is unrelated to the penetration of the compound in the cell (Sava et al 2004). The extent of
ruthenium uptake of three complexes did not correlate with their different cytotoxicity in
ovarian cancer A2780 cell line (Bugarcic et al 2008, Komor and Barton 2013).

Figure 7. The accumulation of 2pG, 2pA, 2mG and 2mA in HeLa cells is different. The HeLa
cells were either treated with 150 μM concentration of 2pG, 2pA, 2mG or 2mA from two to six
hours (A) or with 17.5-150 μM during two hours (B). The cells were collected at the
indicated time points and the amount of ruthenium was measured by HR ICPMS. Two out of
four independent experiments are presented.
Conclusions.
In this study, six triphenylphosphine ligands (L) conjugated to amino acids and ten
ruthenium(II)-cymene complexes thereof, including both ML and ML2 stoichiometry, were
prepared, purified and characterized by NMR (1H, 13C, 31P), CD and MS techniques and DFT
calculations (wB97xD, gas phase). Cytotoxicity screening using human cervical carcinoma
(HeLa) cells and MTT assay revealed the IC50 value of 5-30 μM for all examined ML2
complexes (2pG, 2pA, 2mG and 2mA), while for all other compounds (L, ML and Ru precursor)
the IC50 values were above 33.3 μM. Additional analysis performed with 2mG showed similar
toxicity on several other human carcinomas (laryngeal carcinoma HEp2, lung carcinoma
H460, and breast carcinoma MDA-MB-231), but no toxicity for normal human cell lines
(both keratinocytes and fibroblasts). Studies of non-covalent interactions (thermal melting,
CD and fluorescence titrations) of ruthenium complexes with ct-DNA and BSA, as model
systems for potential biomolecular targets, revealed no measurable interaction with DNA, but
a micromolar affinity for proteins. Interestingly, bioconjugates with lower IC50 values
showed higher binding affinities towards BSA.
Further evaluation of biological activity showed that 2mG increased cells in S phase and
subsequently decreased them in G1 phase of the cell cycle, what was followed by increase of
the SubG0/G1 cell fraction. Staining with Annexin V-FITC and PI indicated programed cell
death as a mode of 2mG action. Caspase independent cell death was deduced from (a) the lack
of PARP-1 cleavage and (b) existence of cell death despite the Z-VAD presence. In addition,
a similar effect obtained by pre-treatment of cells with necrostatin-1 indicated absence of
necroptosis, as another possible mode of cell death. Increase of Beclin-1 as well as LC3-II
expression indicated that 2mG induced autophagy as part of cell death. Moreover, the
autophagy inhibitors BAF-A and 3-MA decreased the toxic effect of 2mG, confirming the
involvement of autophagy in its toxic effect.
Importance of GSH in the cell response to all ML2 complexes (2pG, 2pA, 2mG and 2mA) was
shown using BSO and NAC, revealing decreased (BSO) and increased (NAC) survival,
respectively. Detoxification role of GSH was suggested by the absence of ROS formation and

no effect of two antioxidants, tempol and trolox, on cell survival. On the other hand,
increased survival was detected when ethacrynic acid was used, indicating detoxification role
of GSH. This was further confirmed with inhibition of GSH-2mG conjugates efflux by
probenecid, which had the same effect on cell survival. Cellular uptake, analyzed by HR
ICPMS, showed different dose- and time-dependent cellular accumulation of the studied
complexes. It is particularly interesting to mention that para-substituted complexes (2pG and
2pA) are characterized by higher uptake into the cell, while meta-substituted complexes (2mG
and 2mA) are more toxic.
Here, we described for the first time newly synthesized ruthenium complexes which are
causing non oxidative protein damage and trigger autophagy, and also induce cells protection
mechanism through GSH led detoxification. Taken together, ML2 complexes deserve further
investigation as potential chemotherapeutic agents for different types of cancer.
Experimental.
General. Reactions were carried out in ordinary glassware and chemicals were used as
purchased from commercial suppliers without further purification. Reactions were monitored
by TLC on Silica Gel 60 F254 plates and detected with UV lamp (254 nm); organic
compounds were purified using automated flash chromatography equipped with a UV
detector (254 nm) and prepacked silica columns. NMR spectra were obtained on
1
spectrometer operating at 300.13 or 600.13 MHz for H, 242.92 MHz for 31P and 150.92
13
MHz for C nuclei. Mass spectra were measured on a HPLC-MS system coupled with 6410
triple-quadrupole mass spectrometer, operating in a positive ESI mode. High-resolution mass
spectra were obtained on a MALDI TOF-TOF instrument using a CHCA matrix. The CD
spectra were recorded on a JASCO J-810 spectropolarimeter equipped with Peltier
thermostat, using 1 cm Suprasil quartz cells with a scanning speed of 200 nm min−1. The
buffer background was subtracted from each spectrum, while each spectrum was a result of
three accumulations. The electronic absorption spectra of ruthenium complexes and thermal
melting experiments were measured on a Varian Cary 100 Bio spectrometer. Fluorescence
titrations were recorded on Varian Cary Eclipse fluorimeter. UV-Vis and fluorescence spectra
were recorded using appropriate 1 cm path quartz cuvettes.
Ligands, general procedure. The corresponding 4-(Diphenylphosphino)benzoic acid (1
mmol) was dissolved in dichloromethane (50 mL). HOBt × H2O (1 mmol), TBTU (1 mmol)
and DIPEA (0.5 mL, 4 mmol) were added and the mixture was left stirring at room
temperature. After 1 h methyl ester (1 mmol) was added and the mixture was left stirring for
the indicated period. After that, the reaction mixture was washed with NaHCO3 (3 × 50 mL,
sat. aq.) and subjected to flash chromatography (hexane/EtOAc).
Ph2P-pC6H4-CO-Gly-OMe, LpG; (Kokan and Kirin 2012). 4-(Diphenylphosphino)benzoic
acid (301.6 mg, 1 mmol), dichloromethane (50 mL), HOBt × H2O (135.5 mg, 1 mmol),
TBTU (322.1 mg, 1 mmol), DIPEA (0.5 mL, 4 mmol) and H-Gly-OMe × HCl (129.4 mg, 1
1
mmol), 72 h. Yield: 332.1 mg (88 %). H NMR (300.13 MHz, CDCl3) δ/ppm: 7.75 (dd, 2H,
J1 = 8.5 Hz, J2 = 1.5 Hz), 7.26–7.37 (m, 12 H), 6.62 (t, 1H, J = 4.5 Hz), 4.25 (d, 2H, J = 5
Hz), 3.8 (s, 3H).
Ph2P-pC6H4-CO-Ala-OMe, LpA; (Kokan et al 2014). 4-(Diphenylphosphino)benzoic acid
(302.3 mg, 1 mmol), dichloromethane (50 mL), HOBt × H2O (134.2 mg, 1 mmol), TBTU

(320.3 mg, 1 mmol), DIPEA (0.5 mL, 4 mmol), H-Ala-OMe × HCl (132.1 mg, 1 mmol), 24
1
h. Yield: 296.7 mg (76 %). H NMR (300.13 MHz, CDCl3) δ/ppm: 7.73–7.76 (m, 2H), 7.26–
7.37 (m, 12H), 6.70 (d, 1H, J = 7 Hz), 4.75–4.84 (m, 1H),3.79 (s, 3H), 1.51 (d, 3H, J = 7 Hz).
Ph2P-mC6H4-CO-Gly-OMe, LpG. 3-(Diphenylphosphino)benzoic acid (305.5 mg, 1 mmol),
dichloromethane (50 mL), HOBt × H2O (136.7 mg, 1 mmol), TBTU (322.3 mg, 1 mmol),
DIPEA (0.5 mL, 4 mmol), H-Gly-OMe × HCl (127.2 mg, 1 mmol), 24 h. Yield: 291.8 mg
(77 %). 1H NMR (300.13 MHz, CDCl3) δ/ppm:7.75–7.82 (m, 2H), 7.26–7.44 (m, 12H), 6.51–
13
6.55 (m, 1H), 4.20 (d, 2H, J = 5 Hz), 3.79 (s, 3H). C NMR (CDCl3, 75.46 MHz) δ/ppm:
3
2
41.9 (Cα), 52.6 (OMe), 127.7 (C4), 128.8 (d, JCP = 7 Hz, C3'), 129.0 (d, JCP = 5.5 Hz, C5),
129.2 (C4'), 132.3 (d, JCP = 24 Hz, C2), 133.9 (d, 2JCP = 20 Hz, C2'), 134.0 (C3, overlapped
2
with C2' peak), 136.5 (d, 1JCP = 11 Hz, C1'), 136.9 (d, 2JCP = 16 Hz, C6), 138.7 (d, 1JCP = 13.5
Hz, C1), 167.3 (C(O)NH), 170.5 (COOMe). 31P NMR (CDCl3, 242.93 MHz) δ/ppm: –4.81 (s,
1P).
Ph2P-mC6H4-CO-Ala-OMe, LpA; (Kokan et al 2014). 3-(Diphenylphosphino)benzoic acid
(304.1 mg, 1 mmol), dichloromethane (50 mL), HOBt × H2O (136.2 mg, 1 mmol), TBTU
(321.7 mg, 1 mmol), DIPEA (0.5 mL, 4 mmol), H-Ala-OMe × HCl (142.3 mg, 1 mmol), 24
1
h. Yield: 300.1 mg (77 %). H NMR (300.13 MHz, CDCl3) δ/ppm: 7.76–7.80 (m, 2H), 7.26–
7.44 (m, 12H), 6.61 (d, 1H, J = 8), 4.70–4.80 (m, 1H), 3.77 (s, 1H), 1.48 (d, 3H, J = 7 Hz).
Ph2P-C2H4-CO-Gly-OMe, LaG. 3-(Diphenylphosphino)propionic acid (262.3 mg, 1 mmol),
dichloromethane (50 mL), HOBt × H2O (136.4 mg, 1 mmol), TBTU (324.3 mg, 1 mmol),
DIPEA (0.5 mL, 4 mmol), H-Gly-OMe × HCl (127.2 mg, 1 mmol), 24 h; Rf (Hexane/EtOAc
= 7:3) = 0.10. Yield: 310.2 mg (94 %). 1H NMR (300.13 MHz, CDCl3) δ/ppm: 2.26–2.43 (m,
4H), 3.75 (s, 3H), 4.01 (d, J = 5 Hz, 2H), 5.93 (t, J = 4 Hz, 1H), 7.31–7.36 (m, 6H), 7.39–
13
1
7.46 (m, 4H). C NMR (CDCl3, 75.46 MHz) δ/ppm: 23.4 (d, JCP = 12 Hz, C1 phosphine),
2
3
32.6 (d, JCP = 18.5 Hz, C2 phosphine), 41.4 (Cα), 52.5 (OMe), 128.7 (d, JCP = 7 Hz, C3'
phosphine), 129.0 (C4 phosphine), 132.9 (d, JCP = 18.5 Hz, C2' phosphine), 137.9 (d, JCP
12.5 Hz, C1' phosphine), 170.6 (COOMe), 172.5 (d, 3JCP = 13.5 Hz, C(O)NH).
2
1
=
Ph2P-C2H4-CO-Ala-OMe, LaA. 3-(Diphenylphosphino)propionic acid (261.9 mg, 1 mmol),
dichloromethane (50 mL), HOBt × H2O (136.9 mg, 1 mmol), TBTU (327.3 mg, 1 mmol),
DIPEA (0.5 mL, 4 mmol), H-Ala-OMe × HCl (145.3 mg, 1 mmol), 24 h. Rf (Hexane/EtOAc
= 7:3) = 0.13. Yield: 273.7 mg (80 %). 1H NMR (300.13 MHz, CDCl3) δ/ppm: 1.37 (d, J = 7
Hz, 3H), 2.23–2.41 (m, 4H), 3.74 (s, 3H), 4.52–4.61 (m, 1H), 5.97 (d, J = 7 Hz, 1H), 7.31–
13
7.36 (m, 6H), 7.39–7.46 (m, 4H). C NMR (CDCl3, 75.46 MHz) δ/ppm: 18.7 (Cβ), 23.4 (d,
1JCP = 12 Hz, C1 phosphine), 32.7 (d, 2JCP = 18.5 Hz, C2 phosphine), 48.2 (Cα), 52.6 (OMe),
2
128.7 (d, 3JCP = 7 Hz, C3' phosphine), 128.93, 128.95 (C4 phosphine), 132.86, 132.89 (d, JCP
1
= 18.5 Hz, C2' phosphine), 137.89, 137.96 (d, JCP = 13 Hz, C1' phosphine), 171.8 (d, 3JCP
=
14 Hz, C(O)NH), 173.7 (COOMe). 31P NMR (CDCl3, 242.93 MHz) δ/ppm: –14.94 (s, 1P).
ML complexes, general procedure. Ligand L was dissolved in DCM (3 mL), di-µ-
chlorobis[(p-cymene)chlororuthenium(II)] was added and stirred for 2 h. After the reaction,
the crude product was purified by column chromatography on a short silica column (10 g),
eluent DCM/MeOH.
[(iPr-pC6H4-Me)RuCl2(Ph2P-pC6H4-CO-Gly-OMe)], 1pG. Ligand LpG (201.5 mg, 0.53
mmol) and di-µ-chlorobis[(p-cymene)chlororuthenium(II)] (142.3, 0.23 mmol); eluent
1
DCM/MeOH 2 %, Rf (DCM/MeOH 3 %) = 0.28. Yield: 279.9 mg (89 %). H NMR (300.13

MHz, CDCl3) δ/ppm: 7.69–7.95 (m, 8H), 7.37–7.47 (m, 6H), 6.64 (t, 1H, J = 5 Hz), 5.22 (d,
2H, J = 6 Hz), 4.98 (d, 2H, J = 6 Hz), 4.21 (d, 2H, J = 5 Hz), 3.78 (s, 3H), 3.49 (d, 1H, J =
5.5 Hz), 2.81–2.91 (m, 1H),1.86 (s, 3H), 1.12 (d, 6H, J = 7 Hz). 13C NMR (CDCl3, 150.92
MHz) δ/ppm: 18.0 (CH3, cymene), 22.0 (CHCH3, cymene), 30.4 (CHCH3, cymene),41.9
2
2
(Cα),52.6 (OMe), 87.5 (d, JCP = 5.5 Hz, C2, C6, cymene), 89.0 (d, JCP = 3 Hz, C3, C5,
2
3
cymene), 96.4 (C4, cymene), 111.6 (d, JCP = 3.5 Hz, C1, cymene), 126.5 (d, JCP = 10 Hz,
3
C3, phosphine), 128.4 (d, JCP = 10 Hz, C3', phosphine), 130.7 (C4', phosphine), 133.5 (d,
1JCP = 45 Hz, C1', phosphine), 134.3 (d, JCP = 9.5 Hz, C2', phosphine), 134.8 (d, JCP = 9.5
2
2
1
Hz, C2, phosphine), 135.1 (C4, phosphine), 137.7 (d, JCP = 44 Hz, C1, phosphine), 167.1
(C(O)NH), 170.4 (COOMe). 31P NMR (CDCl3, 242.93 MHz) δ/ppm: 25.32 (s, 1P). MALDI-
HRMS (m/z): calcd 613.1320 (C32H34NO3PRu+), found 613.1329.
[(iPr-pC6H4-Me)RuCl2(Ph2P-pC6H4-CO-Ala-OMe)], 1pA. Ligand LpA (191 mg, 0.49
mmol) and di-µ-chlorobis[(p-cymene)chlororuthenium(II)] (131.2, 0.21 mmol); eluent
1
DCM/MeOH 2 %, Rf (DCM/MeOH 3 %) = 0.28. Yield: 218.3 mg (75 %). H NMR (300.13
MHz, CDCl3) δ/ppm: 7.69–7.95 (m, 8H), 7.37–7.47 (m, 6H), 6.67 (d, 1H, J = 7.5 Hz), 5.21–
5.24 (m, 2H), 4.98 (d, 2H, J = 6 Hz), 4.71–4.80 (m, 1H),3.77 (s, 3H), 2.8–2.93 (m, 1H),1.87
(s, 3H), 1.49 (d, 3H, J = 7 Hz), 1.12 (d, 6H, J = 7 Hz). 13C NMR (CDCl3, 150.92 MHz)
δ/ppm: 18.0 (CH3, cymene), 18.6 (Cβ), 22.0 (CHCH3, cymene), 30.4 (CHCH3, cymene), 48.7
2
2
(Cα), 52.7 (OMe), 87.48, 87.52 (d, JCP = 5 Hz, C2, C6, cymene), 88.99, 89.04 (d, JCP = 3
Hz, C3, C5, cymene), 96.3 (C4, cymene), 111.6(d, 2JCP = 3.5 Hz, C1, cymene), 126.4 (d, 3JCP
= 10 Hz, C3, phosphine), 128.3 (d, JCP = 10 Hz, C3', phosphine), 130.7 (C4', phosphine),
133.51, 133.52 (d, JCP = 45 Hz, C1', phosphine), 134.4 (d, JCP = 9.5 Hz, C2', phosphine),
3
1
2
2
1
134.8 (d, JCP = 9.5 Hz, C2, phosphine), 135.28, 135.29 (C4, phosphine), 137.7 (d, JCP
=
44.5 Hz, C1, phosphine), 166.5 (C(O)NH), 173.4 (COOMe). 31P NMR (CDCl3, 242.93 MHz)
δ/ppm: 25.31 (s, 1P). MALDI-HRMS (m/z): calcd 662.1165 (C33H36ClNO3PRu+), found
662.1131.
[(iPr-pC6H4-Me)RuCl2(Ph2P-mC6H4-CO-Gly-OMe)], 1mG. Ligand LmG (182.2 mg, 0.48
mmol) and di-µ-chlorobis[(p-cymene)chlororuthenium(II)] (147.1, 0.24 mmol); eluent
1
DCM/MeOH 2 %, Rf (DCM/MeOH 3 %) = 0.26. Yield: 301.1 mg (92 %). H NMR (300.13
MHz, CDCl3) δ/ppm: 8.83 (dt, 1H, J1 = 12 hz, J2 = 1.5 Hz), 7.78 - 7.91 (m, 5H), 7.56–7.63
(m, 1H), 7.31–7.49 (m, 7H), 6.93 (t, 1H, J = 5.5 Hz), 5.22 (d, 2H, J = 6 Hz), 5.07 (d, 2H, J =
6 Hz), 5.19 (d, 2H, J = 5.5 Hz), 3.77 (s, 3H), 2.77–2.86 (m, 1H), 1.88 (s, 3H), 1.08 (d, 6H, J =
7 Hz). 13C NMR (CDCl3, 150.92 MHz) δ/ppm: 17.9 (CH3, cymene), 21.9 (CHCH3, cymene),
2
30.4 (CHCH3, cymene), 41.9 (Cα), 52.4 (OMe), 87.2 (d, JCP = 5.5 Hz, C2, C6, cymene),
2
2
89.5 (d, JCP = 2.5 Hz, C3, C5, cymene), 96.2 (C4, cymene), 111.6 (d, JCP = 2.5 Hz, C1,
3
3
cymene), 128.3 (d, JCP = 8.5 Hz, C5, phosphine), 128.5 (d, JCP = 9.5 Hz, C3', phosphine),
1
129.8 (C4, phosphine), 130.8 (C4', phosphine), 132.4 (d, JCP = 46 Hz, C1', phosphine), 132.9
(d, 3JCP = 11 Hz, C3, phosphine), 133.9 (d, 2JCP = 9.5 Hz, C2', phosphine), 134.1 (d, 1JCP = 46
2
Hz, C1, phosphine), 135.2 (d, JCP = 16 Hz, C2, phosphine), 136.73, 136.75 (C6, phosphine),
167.2 (C(O)NH), 170.3 (COOMe). 31P NMR (CDCl3, 242.93 MHz) δ/ppm: 25.50 (s, 1P).
+ESI MS (m/z): 648.1 ([M – Cl]+, 81 %). MALDI-HRMS (m/z): calcd 612.1241
(C32H33NO3PRu+), found 612.1226.
[(iPr-pC6H4-Me)RuCl2(Ph2P-mC6H4-CO-Ala-OMe)], 1mA. Ligand LmA (180 mg, 0.46
mmol) and di-µ-chlorobis[(p-cymene)chlororuthenium(II)] (141.6, 0.23 mmol); eluent
1
DCM/MeOH 2 %, Rf (DCM/MeOH 3 %) = 0.28. Yield: 302 mg (94 %). H NMR (300.13
MHz, CDCl3) δ/ppm: 8.75–8.80 (m, 1H), 7.76–7.78 (m, 5H), 7.61–7.68 (m, 1H), 7.31–7.47
(m, 7H), 6.88 (d, 1H, J = 8H), 5.26–5.30 (m, 2H), 5.17 (d, 2H, J = 6.5 Hz), 4.96 (d, 2H, J = 6

Hz), 4.65–4.75 (m, 1H), 3.76 (s, 3H), 2.75–2.89 (m, 1H), 1.88 (s, 3H), 1.52 (d, 3H, J = 7 Hz),
13
1.08 (t, 6H, J = 7.5 Hz). C NMR (CDCl3, 150.92 MHz) δ/ppm: 17.9 (CH3, cymene), 18.3
(Cβ), 21.8,22.0 (CHCH3, cymene), 30.4 (CHCH3, cymene), 48.8 (Cα), 52.6 (OMe), 86.987.6
(d, 2JCP = 5.5 Hz, C2, C6, cymene), 89.3, 89.4 (d, 2JCP = 2.5 Hz, C3, C5, cymene), 96.3 (C4,
2
3
cymene), 111.5 (d, JCP = 3 Hz, C1, cymene), 128.3 (d, JCP = 8.5 Hz, C5, phosphine),128.5
3
(d, JCP = 9.5 Hz, C3', phosphine), 129.6 (C4, phosphine),130.8 (C4', phosphine), 132.6 (d,
1JCP = 46 Hz, C1', phosphine), 133.1 (d, 3JCP = 11 Hz, C3, phosphine), 133.9–134.4 (C2', C1,
2
phosphine), 135.1 (d, JCP = 16 Hz, C2, phosphine), 136.72, 136.74 (C6, phosphine), 166.5
(C(O)NH), 173.4 (COOMe). 31P NMR (CDCl3, 242.93 MHz) δ/ppm: 25.49 (s, 1P). +ESI MS
(m/z): 662.1 ([M – Cl]+, 100 %). MALDI-HRMS (m/z): calcd 627.1476 (C33H36NO3PRu+),
found 627.1504.
[(iPr-pC6H4-Me)RuCl2(Ph2P-C2H4-CO-Gly-OMe)], 1aG. Ligand LaG (180 mg, 0.55mmol)
and di-µ-chlorobis[(p-cymene)chlororuthenium(II)] (167.9, 0.28mmol); eluent DCM/MeOH
1
1 % using 40 g silica, Rf (DCM/MeOH 3 %) = 0.34. Yield: 297.2 mg (85 %). H NMR
(300.13 MHz, CDCl3) δ/ppm: 0.98 (d, J = 7 Hz, 6H), 2.21–2.26 (m, 2H), 2.62 (septuplet, J =
7 Hz, 1H), 2.88–2.93 (m, 2H), 3.68 (s, 3H), 3.78 (d, J = 5.5 Hz, 2H), 5.09 (d, J = 6 Hz, 2H),
13
5.18 (d, J = 6 Hz, 2H), 6.21 (t, J = 5 Hz, 2H), 7.45–7.50 (m, 6H), 7.78–7.80 (m ,4H). C
1
NMR (CDCl3, 75.46 MHz) δ/ppm:17.7 (CH3, cymene), 21.8 (CHCH3, cymene), 22.9 (d, JCP
2
= 30 Hz, C1 phosphine), 30.2 (CHCH3, cymene), 30.9 (d, JCP = 2 Hz, C2 phosphine), 41.3
2
2
(Cα), 52.3 (OMe), 86.1 (d, JCP = 6 Hz, C2, C6, cymene), 90.1 (d, JCP = 4 Hz, C3, C5,
2
3
cymene), 95.1 (C4, cymene), 109.5 (d, JCP = 1 Hz, C1, cymene), 128.6 (d, JCP = 9 Hz, C3'
2
phosphine), 130.90, 130.93 (C4' phosphine), 133.3 (d, JCP = 9 Hz, C2' phosphine), 133.5 (d,
1JCP = 43 Hz, C1' phosphine), 170.2 (COOMe), 172.6 (d, JCP = 13 Hz, C(O)NH). P NMR
(CDCl3, 242.93 MHz) δ/ppm: 21.36 (s, 1P). MALDI-HRMS (m/z): calcd 600.1008
(C28H34ClNO3PRu+), found 600.1011.
3
31
[(iPr-pC6H4-Me)RuCl2(Ph2P-C2H4-CO-Ala-OMe)], 1aA. Ligand LaA (164 mg, 0.48mmol)
and di-µ-chlorobis[(p-cymene)chlororuthenium(II)] (146.3, 0.24mmol); eluent DCM/MeOH
1 % using 40 g. Yield: 196.8 mg (63 %). 1H NMR (300.13 MHz, CDCl3) δ/ppm: 0.96 (d, J =
7 Hz, 3H), 0.98 (d, J = 7 Hz, 3H), 1.28 (d, J = 7 Hz, 3H), 2.06–2.29 (m, 2H), 2.63 (septuplet,
J = 7 Hz, 1H), 2.85–2.94 (m, 2H), 3.67 (s, 3H), 4.26–4.36 (m, 1H),5.07–5.11 (m, 2H), 5.17–
5.20 (m, 2H), 6.14 (d, J = 7 Hz, 1H), 7.43–7.50 (m, 6H), 7.76–7.82 (m ,4H). 13C NMR
(CDCl3, 75.46 MHz) δ/ppm:17.7 (CH3, cymene), 18.2 (Cβ), 21.7, 21.9 (CHCH3, cymene),
1
22.9 (d, JCP = 31 Hz, C1 phosphine), 30.2 (CHCH3, cymene), 31.0 (C2 phosphine), 48.2
2
2
(Cα), 52.4 (OMe), 86.1 (d, JCP = 5.5 Hz, C2, C6, cymene), 90.1 (d, JCP = 4 Hz, C3, C5,
2
3
cymene), 95.2 (C4, cymene), 109.5 (d, JCP = 1 Hz, C1, cymene), 128.59, 128.62 (d, JCP
9.5 Hz, C3' phosphine), 130.88–130.92 (m, C4' phosphine), 133.3, 133.4 (d, JCP = 8 Hz, C2'
=
2
1
3
phosphine), 133.56, 133.62 (d, JCP = 43 Hz, C1' phosphine), 172.0 (d, JCP = 14 Hz,
31
C(O)NH), 173.3 (COOMe). P NMR (CDCl3, 242.93 MHz) δ/ppm: 21.35 (s, 1P). MALDI-
HRMS (m/z): calcd 614.1165 (C29H36ClNO3PRu+), found 614.1180.
ML2 complexes, general procedure. Mono complex 1 and NH4PF6 were dissolved in
CH3CN (5 mL) and refluxed for 35–45 min. CH3CN was evaporated, the residue dissolved in
DCM (5 mL) and filtrated through Celite. Ligand L was dissolved in DCM (2 mL), added to
the filtrate and the mixture was stirred for 24 h. The crude product was purified by column
chromatography on silica (40g), using DCM/MeOH 1 % as eluent.
[(iPr-pC6H4-Me)RuCl(Ph2P-pC6H4-CO-Gly-OMe)2]PF6, 2pG. Mono complex 1pG (51.2
mg, 0.075 mmol), NH4PF6 (18.2 mg, 0.11 mmol) and ligand LpG (70 mg, 0.19 mmol). Rf

1
(DCM/MeOH 3 %) 0.18. Yield: 32.6 mg (37 %). H NMR (300.13 MHz, CDCl3) δ/ppm:
7.23–7.60 (m, 28H), 7.12 (t, 2H, J = 5.5 Hz), 5.53 (d, 2H, J = 4 Hz), 5.14 (d, 2H, J = 6H),
4.21 (d, 4H, J = 5.5 Hz), 3.78 (s, 6H), 2.68–2.77 (m, 1H), 1.24 (d, 6H, J = 7 Hz), 1.09 (s, 3H).
13C NMR (CDCl3, 150.92 MHz) δ/ppm: 15.5 (CH32, cymene), 21.6 (CHCH3, cymene), 31.6
(CHCH3, cymene), 41.9 (Cα), 52.5 (OMe), 89.3 (t, JCP3 = 5 Hz, C2, C6, cymene), 97.6 (C3,
C5, cymene), 101.0 (C4, cymene), 126.97, 127.01 (d, JCP = 5 Hz, C3, phosphine), 128.64,
128.68, 128.85, 128.88 (d, 3JCP = 5 Hz, C3', phosphine), 131.2, 131.6 (C4', phosphine), 131.8
(C1, cymene), 133.2–133.9 (m, C1' phosphine), 134.12, 134.16, 134.26, 134.30, 134.49,
2
2
134.52 (d, JCP = 4 Hz, C2, C2', phosphine), 136.2 (C4, phosphine), 136.4 (d, JCP = 45 Hz,
C1, phosphine), 167.2 (C(O)NH), 170.3 (COOMe). 31P NMR (CDCl3, 242.93 MHz) δ/ppm: –
143.93 (septuplet, hexafluorophosphate), 21.42 (s, 1P). MALDI-HRMS (m/z): calcd
991.2579 (C54H5455N2O6P2Ru+), found 991.2537.
[(iPr-pC6H4-Me)RuCl(Ph2P-pC6H4-CO-Ala-OMe)2]PF6, 2pA. Mono complex 1pA (70 mg,
0.1 mmol), NH4PF6 (24.5 mg, 0.15 mmol) and ligand LpA (100 mg, 0.25 mmol). Rf
1
(DCM/MeOH 3 %) 0.22. Yield: 84.5 mg (71 %). H NMR (300.13 MHz, CDCl3) δ/ppm:
7.26–7.58 (m, 28H), 7.03 (d, 2H, J = 7 Hz), 5.55 (t, 2H, J = 4.5 Hz), 5.24 (d, 1H, J = 6 Hz),
5.09 (d, 1H, J = 6 Hz), 4.66–4.76 (m, 2H), 3.78 (s, 6H), 2.67–2.76 (m, 1H), 1.54 (d, 6H, J = 7
13
Hz), 1.24 (t, 6H, J = 7 Hz), 1.08 (s, 3H). C NMR (CDCl3, 150.92 MHz) δ/ppm: 15.4 (CH3,
cymene), 17.9 (Cβ), 21.54, 21.58 (CHCH3, cymene), 31.6 (CHCH3, cymene), 49.0 (Cα), 52.7
3
(OMe), 89.3, 89.6 (d, 2JCP = 9.5 Hz, C2, C6, cymene), 97.4, 97.7 (d, JCP = 3 Hz, C3, C5,
cymene), 101.0 (C4, cymene), 126.9–127.0 (m, C3, phosphine), 128.6–128.8 (m, C3',
phosphine), 131.19, 131.22, 131.47 (C4', phosphine), 131.8 (C1, cymene), 133.1–133.9 (m,
C1', phosphine), 134.0–134.5 (m, C2, C2', phosphine), 136.3, 136.4 (C4, phosphine), 136.5
2
(d, JCP = 43 Hz, C1, phosphine),166.58, 166.62 (C(O)NH), 173.4 (COOMe).31P NMR
2
(CDCl3, 242.93 MHz) δ/ppm: –143.96 (septuplet, hexafluorophosphate), 21.26 (d, 1P, JPP =
2
52 Hz), 21.81 (d, 1P, JPP = 52 Hz). MALDI-HRMS (m/z): calcd 1019.2892
(C56H58N2O6P2Ru+), found 1019.2852.
[(iPr-pC6H4-Me)RuCl(Ph2P-mC6H4-CO-Gly-OMe)2]PF6, 2mG. Mono complex 1mG (80.3
mg, 0.12 mmol), NH4PF6 (30.3 mg, 0.19 mmol) and ligand LmG (100 mg, 0.26 mmol). Rf
1
(DCM/MeOH 3 %) 0.22. Yield: 46.5 mg (33 %). H NMR (300.13 MHz, CDCl3) δ/ppm:
7.96 (t, 2H, J = 5.5 Hz), 7.78 (d, 2H, J = 7.5 Hz), 7.2–7.54 (m, 24H), 7.04 (t, 2H, J = 5 Hz),
5.58 (d, 2H, J = 5.5 Hz), 5.3 (d, 2H, J = 6 Hz), 4.17 (d, 4H, J = 5.5 Hz), 3.79 (s, 6H), 2.70–
2.80 (m, 1H),1.23 (d, 6H, J = 7 Hz), 1.07 (s, 3H). 13C NMR (CDCl3, 150.92 MHz) δ/ppm:
15.4 (CH3, cymene), 21.6 (CHCH3, cymene), 31.5 (CHCH3, cymene), 41.8 (Cα), 52.5 (OMe),
89.5 (m, C2, C6, cymene), 97.5 (C4, cymene), 100.9 (C3, C5, cymene), 128.6–128.9 (m, C5,
C3', phosphine), 129.6–129.7 (m, C4, phosphine), 131.2, 131.6 (C4', phosphine), 131.4 (C1,
cymene),132.5 (d, 3JCP = 7 Hz, C3, phosphine), 133.5–133.9 (C1, C1', phosphine), 134–134.6
(C2, C2', phosphine), 136.94, 137.0 (d, 2JCP = 4 Hz, C6, phosphine), 166.6, 166.7 (C(O)NH),
170.3 (COOMe). 31P NMR (CDCl3, 242.93 MHz) δ/ppm: –143.97 (septuplet,
hexafluorophosphate), 21.95 (s, 1P). +ESI MS (m/z): 1025.3 (M+, 23 %). MALDI-HRMS
(m/z): calcd 991.2579 (C54H54N2O6P2Ru+), found 991.2537.
[(iPr-pC6H4-Me)RuCl(Ph2P-mC6H4-CO-Ala-OMe)2]PF6, 2mA. Mono complex 1mA (78.9
mg, 0.11 mmol), NH4PF6 (27 mg, 0.17 mmol) and ligand LmA (110 mg, 0.28 mmol). Rf
1
(DCM/MeOH 3 %) = 0.26. Yield: 81.2 mg (60 %). H NMR (300.13 MHz, CDCl3) δ/ppm:
7.92–8.01 (m, 2H), 7.71 (d, 2H, J = 7.5 Hz), 7.18–7.56 (m, 24H), 6.81–6.91 (m, 2H), 5.58–
5.62 (m, 2H), 5.21–5.30 (m, 4H), 4.62–4.72 (m, 2H), 3.79 (d, 6H, J = 4Hz), 2.69–2.78 (m,
13
1H),1.51 (dd, 6H, J1 = 7 Hz, J2 = 1 Hz), 1.21–1.25 (m, 6H), 1.11 (s, 3H). C NMR (CDCl3,

75.48 MHz) δ/ppm: 15.5 (CH3, cymene), 18.17, 18.22 (Cβ), 21.5, 21.6 (CHCH3, cymene),
31.5 (CHCH3, cymene), 48.79, 48.81 (Cα), 52.7 (OMe), 89.68, 89.80, 89.85, 89.95 (m, C2,
2
C6, cymene), 97.4, 97.5 (d, JCP = 2.5 Hz, C3, C5, cymene), 101.2 (C4, cymene), 128.6,
3
128.7 (C5, phosphine), 128.9 (d, JCP = 16.5 Hz, C3', phosphine), 129.3, 129.5 (C4,
phosphine), 131.4, 131.6, 131.7 (C4', phosphine), 131.8 (C1, cymene), 132.3, 132.9 (C1’,
3
phosphine), 133.4–133.7 (m, C2, phosphine), 133.8 (C1, phosphine), 134.0 (d, JCP = 14.5
Hz, C3’, phosphine), 134.4 (C3, phosphine),134.5–134.8 (m, C2’, phosphine), 136.9–137.3
31
(C6, phosphine), 166.0 (C(O)NH), 173.6, 173.7 (COOMe). P NMR (CDCl3, 242.93 MHz)
2
δ/ppm: –143.84 (septuplet, hexafluorophosphate), 22.08 (d, 1P, JPP = 52 Hz), 22.40 (d, 1P,
2JPP = 52 Hz). +ESI MS (m/z): 1053.3 (M+, 3 %). MALDI-HRMS (m/z): calcd 1019.2892
(C56H58N2O6P2Ru+), found 1019.2852.
[(iPr-pC6H4-Me)RuCl(Ph2P-C2H4-CO-Gly-OMe)2]PF6, 2aG. Mono complex 1aG (81.1 mg,
0.13mmol), NH4PF6 (32.1 mg, 0.20mmol) and ligand LaG (104 mg, 0.32mmol). Rf
(DCM/MeOH 5 %) = 0.42. Yield: 60.9 mg (44 %). As mentioned in the discussion, this
1
compound could not be isolated with stratifying purity. H NMR (300.13 MHz, CDCl3)
δ/ppm: 1.22 (d, J = 7 Hz, 6H), 1.84–1.95 (m, 4H), 2.49 (ws, 2H), 2.75 (septuplet, J = 7 Hz,
1H), 3.03–3.15 (m, 2H), 3.72 (s, 3H), 3.91 (d, J = 5.5 Hz, 4H), 5.18 (d, J = 6 Hz, 2H), 5.63–
5.67 (m, 2H), 6.06 (t, J = 5 Hz, 2H), 7.36–7.63 (m, 20H).13C NMR (CDCl3, 150.92 MHz)
δ/ppm:14.7 (CH3, cymene), 21.5 (CHCH3, cymene), 22.3–22.5 (m, C1 phosphine), 31.0 (C2
2
phosphine), 31.6 (CHCH3, cymene), 41.4 (Cα), 52.4 (OMe), 87.4 (t, JCP = 5 Hz, C2, C6,
2
cymene), 97.1 (C4, cymene), 97.7 (C3, C5, cymene), 129.3 (d, JCP = 46 Hz, C1', phosphine),
3
129.30, 129.34, 129.71, 129.75 (d, JCP = 5 Hz, C3' phosphine), 131.6, 131.9 (C4,
2
phosphine), 132.16, 132.19, 132.95, 132.98 (d, JCP = 4 Hz, C2' phosphine), 132.8
3
31
(C1,cymene), 170.3 (COOMe), 171.14, 171.18 (d, JCP = 6 Hz, C(O)NH). P NMR (CDCl3,
242.93 MHz) δ/ppm: –143.91 (septuplet, hexafluorophosphate), 20.41 (s, 1P).
[(iPr-pC6H4-Me)RuCl(Ph2P-C2H4-CO-Ala-OMe)2]PF6, 2aA. Mono complex 1aA (65.5 mg,
0.10mmol), NH4PF6 (24.9 mg, 0.15mmol) and ligand LaA (85 mg, 0.25mmol). Rf
(DCM/MeOH 5 %) 0.45. Yield: 64.1 mg (42 %). As mentioned in the discussion, this
1
compound could not be isolated with stratifying purity. H NMR (300.13 MHz, CDCl3)
δ/ppm: 1.22 (d, J = 7 Hz, 6H), 1.84–1.95 (m, 4H), 2.49 (ws, 2H), 2.75 (septuplet, J = 7 Hz,
1H), 3.03–3.15 (m, 2H), 3.72 (s, 3H), 3.91 (d, J = 5.5 Hz, 4H), 5.18 (d, J = 6 Hz, 2H), 5.63–
5.67 (m, 2H), 6.06 (t, J = 5 Hz, 2H), 7.36–7.63 (m, 20H).
Computational details. All calculations were performed using the Gaussian09 program
package (see references section). Geometries were optimized using the wB97xD density
functional (Chai and Head-Gordon 2008) in conjunction with the 6-31G(d,p) for the first-row
elements and 6-31+G(d,p) basis set for phosphorus and chlorine, while the SDD effective
core potential (Andrae et al 1990) was employed for ruthenium. The nature of the stationary
points was verified by vibrational analysis at the optimized geometries and no imaginary
frequencies were obtained. Total Gibbs energies (Gtot) were calculated by summing
electronic energies with Gibbs energy correction as obtained from the calculations with
default calculation settings and without any scaling of the vibrational frequencies.
Visualization of the optimized structures was done by MOLDEN 5.0. (Schaftenaar and
Noordik 2000).
Biological testing. All examined organometallic ruthenium complexes were dissolved in
DMSO, (c = 10 mM), and stored at –20 °C. Just before use, these stock solutions were diluted
with growth medium to the appropriate concentrations; only concentrations of the complexes

below 33.3 μM were used, with the highest DMSO concentration well under 0.5%.
Ethacrynic acid (ETA; Sigma-Aldrich, USA), N-Benzyloxycarbonyl-Val-Ala-Asp(O-Me)
fluoromethyl ketone (Z-VAD-FMK; Fisher Scientific, USA) and Bafilomycin A1 (BAF A;
InvivoGen, USA) were dissolved in DMSO (Sigma-Aldrich) and kept at –20 °C. Caspase
3/7-Glo® Assay System (Promega, USA) was dissolved according to producer instruction and
kept at –20 °C. Buthionine sulfoximine (BSO; Sigma-Aldrich), N-acetylcysteine (NAC;
Sigma-Aldrich) and probenecid were dissolved in water and kept at –20 °C. 3-(4,5-dimethyl-
2-thiazolyl)-2,5-diphenyl-2H-tetrazolium bromide (MTT) was purchased by Sigma-Aldrich,
dissolved in phosphate-buffered saline and kept by 4 °C.
Cell culture. Human cervical carcinoma (HeLa) and laryngeal carcinoma (HEp2) cells were
obtained from cell culture bank (GIBCO BRL-Invitrogen, USA). Large cell lung carcinoma
(H460) and human breast adenocarcinoma (MDA-MB-213) cells were obtained from
American Type Culture Collection (ATCC, USA). Normal human skin keratinocyte were
obtained from the foreskin of healthy boys, aged 3–8 years. Foreskin samples were non-
inflamed and the children were free of any therapy at least 1 month before the surgery. The
cells were obtained at the Neurochemical Laboratory, Department of Chemistry and
Biochemistry, School of Medicine, University of Zagreb (Gabrilovac et al 2004). Normal
human skin fibroblasts were isolated from the upper arm of a 7-years-old female donor at the
Neurochemical Laboratory, Department of Chemistry and Biochemistry, School of Medicine,
University of Zagreb. They were used for the cytotoxicity assay at 35 and 40 population
doublings. All cell lines were grown as a monolayer culture in Dulbecco’s modified Eagle’s
medium (DMEM; Sigma-Aldrich, USA), supplemented with 10% fetal bovine serum (FBS;
Sigma-Aldrich) in a humidified atmosphere of 5% CO2 at 37 °C and were sub-cultured every
3–4 days.
Cytotoxic assay. Cytotoxic activity of newly synthesized organometallic ruthenium
complexes was determined by MTT assay (Mickisch et al 1990) modified accordingly. In
short, the cells were seeded into 96-well tissue culture plates. The next day, different
concentrations of compounds were added to each well in quadruplicate. Upon 72 h incubation
at 37 °C, the medium was aspirated, and the MTT dye (Sigma-Aldrich) was added. Three
hours later, the formed formazan crystals were dissolved in DMSO, the plates were
mechanically agitated for 5 min and the optical density at 545 nm was determined on a
microtiter plate reader (Awareness Technology Inc., USA). Sulforhodamine B (SRB; Sigma-
Aldrich) was used in order to perform SRB assay according to the protocol (Skehan et al
1990). In short, cultures fixed with trichloroacetic acid were stained for 30 minutes with 0.4%
(wt/vol) SRB dissolved in 1% acetic acid. Unbound dye was removed by four washes with
1% acetic acid, and protein-bound dye was extracted with 10 mM unbuffered Tris base [tris
(hydroxymethyl)aminomethane] for determination of optical density in a computer-
interfaced, 96-well microtiter plate reader (560 nm, Awareness Technology Inc.). The percent
of cell survival for each tested concentration of the compounds was calculated according to
the absorption value of non-treated control cells, which was set as 100 %.
Potential biomolecular target evaluation. Polynucleotide and protein were purchased as
noted: calf thymus (ct)-DNA (Sigma) and BSA (bovine serum albumin, Sigma), and
dissolved in sodium cacodylate buffer, pH = 7.0, I = 0.05 M. The calf thymus ct-DNA was
additionally sonicated and filtered through a 0.45 mm filter (Chaires et al 1982). The ct-DNA
concentration was determined spectroscopically at 260 nm using a molar extinction
coefficient (ε) value of 6550 M−1 cm−1 and it was expressed as the concentration of
phosphates (Bresloff and Crothers 1981).

All examined organometallic ruthenium complexes were dissolved in DMSO (c = 10-2 M or
c= 10-3 M), and stored in refrigerator. Stock solutions were diluted with buffer during the
experiment or immediately before. The highest DMSO content in solution was ≤1%.
Refrigerated DMSO stock solutions were stable longer than a month, while refrigerated
DMSO solutions diluted with buffer were checked to be stable for more than one week.
Concentrations of ruthenium complexes below 2  10-5 M were used to avoid intermolecular
association. At given experimental conditions the absorbance of measured compounds was
proportional to their concentrations.
Thermal melting curves for ct-DNA and its complexes with studied organometallic
ruthenium complexes were determined following the absorption change at 260 nm as a
function of temperature (Mergny and Lacroix 2003). Absorbance scale was normalized. Tm
values were the midpoints of the transition curves determined from the maximum of the first
derivative and checked graphically by the tangent method. The Tm values were calculated
by subtracting Tm value of the free polynucleotide from Tm value of the complex. Every
Tm value was the average of at least two measurements. The error in Tm is ± 0.5 °C.
CD titrations were performed by adding portions of the compound stock solution into the
solution of polynucleotide (c = 2 × 10−5 M) or protein (c = 2 × 10−6 M). Scanning speed was
200 nm min−1, the buffer background was subtracted from each spectrum, while each
spectrum was a result of three accumulations.
Fluorimetric titrations were performed by adding aliquots of organometallic ruthenium
complexes stock solution into the buffered solution of the BSA protein and by monitoring
fluorescence of protein. Excitation wavelength of λexc = 300 nm was used to avoid absorption
of excitation light caused by increasing absorbance of the organometallic ruthenium complex.
After mixing protein with studied compounds it was observed that equilibrium was reached
in less than 120 seconds. Due to low concentrations of studied compounds and protein used
in fluorimetric titrations no precipitation occurred. Emission was collected in the range
λem=320 – 450 nm. Titration data were processed by non-linear least-square fitting program
SPECFIT (Gampp et al 1985, Maeder and Zuberbuhler 1990) that gave the best fit of 1:1
stoichiometry of complexes. The binding constants of complexes of examined compounds
with BSA protein were calculated for the concentration range corresponding to ca. 20–80%
complexation.
Cell cycle analysis. HeLa cells were seeded into tissue culture plates and treated with either
10 μM of 2mG during 24-72 h or different concentrations of the compound during 72 h, in
order to analyze the cell cycle progression. Thereafter, both adherent and floating cells were
collected, washed with PBS, fixed in 70% ethanol and left overnight at –20°C. Fixed cells
were treated with RNase A (0.1 mg/mL, Sigma-Aldrich) for one hour at room temperature
and afterward stained with propidium iodide (50 μg/mL, Sigma-Aldrich) for 30 min in the
dark. The DNA content was analyzed by flow cytometry NaviosTM (Beckman Coulter,
Miami, Fl, USA). Data were analyzed with FlowLogic software (Inivai Technologies,
Victoria, Australia). Propidium iodide stained samples were analyzed using Watson
pragmatic algorithm for modeling cell cycle data.
Cell death analysis. Twenty-four hours after the seeding, HeLa cells were treated either with
10 and 16 μM concentrations of 2mG during 24-72 h or with different concentrations of 2mG
during 72 h. Starved or heat shocked cells were always used as a positive (compensation)

controls. After indicated time point, both adherent and floating cells were collected,
centrifuged and washed with PBS. The cell suspension was incubated with Annexin V-FITC
(BD Biosciences, USA; according to producer’s protocol) and propidium iodide (5μg/mL,
Sigma-Aldrich). Upon 30 min incubation at room temperature in dark, the viable, early
apoptotic, late apoptotic/necrotic and necrotic cell populations were detected and counted by
flow cytometry NaviosTM (Beckman Coulter, Miami, Fl, USA). Data were analyzed with
FlowLogic software (Inivai Technologies, Victoria, Australia).
Twenty-four hours after the seeding, HeLa cells were treated with 22 μM 2mG. The total cell
lysates were collected 24-72 h upon treatment and loaded onto a 10% SDS polyacrylamide
gel and run for two hours at 35 mA. Separated proteins were transferred onto a 0.2 mm
nitrocellulose membrane (Schleicher and Schull, Germany) in a Bio-Rad blot cell (Bio-Rad,
USA), using buffer consisting of 25 mM Tris/HCl, 86 mM glycine and 20% methanol. To
avoid nonspecific binding, the membrane was incubated in blocking buffer (5% non fat dry
milk, 0.1% Tween 20 in PBS) for one hour at room temperature and then incubated with anti-
PARP (Cell Signaling Technology, USA), Beclin-1 (Santa Cruz Biotechnology, USA) or
LCR-I/II antibody (MBL, USA) at room temperature for two hours. After washing the
membrane with 0.1% Tween 20 in TBS and incubation with corresponding horseradish
peroxidase-coupled secondary antibody (Amersham Pharmacia Biotech, Germany), the
proteins were visualized with ECL (Perkin Elmer, USA) according to the manufacturer’s
protocol. All membranes were incubated with β-Tubulin, which was used as a loading control
(Sigma-Aldrich, USA). The role of caspases in 2mG induced cell death was investigated by
MTT assay. HeLa cells were pre-treated for two hours with 6.25 μM specific pan-caspase
inhibitor, Z-VAD. Upon pre-treatment, different concentrations of 2mG were added and the
cytotoxicity effect was determined 72 h later as described above.
The role of autophagy in 2mG induced cell death was also investigated by MTT assay. HeLa
cells were pre-treated for two hours with 0.5 nM specific inhibitor of autophagy, BAF A.
Upon pre-treatment, different concentrations of 2mG were added and the cytotoxicity effect
was determined 72 h later as described above.
Determination of glutathione role in cell response. The role of intracellular GSH in cell
response to 2pG, 2pA, 2mG and 2mA was investigated by MTT assay. HeLa cells were either
pre-treated overnight with specific inhibitor of GSH synthesis, 0.01 μg/mL BSO or for two
hours with precursor in GSH synthesis, 5 mM NAC. Upon pre-treatment, different
concentrations of indicated organometallic complexes were added and the cytotoxicity effect
was determined 72 h later as described above. The capacity of GSH to form the detoxification
conjugates through enzymatic reaction with 2mG was investigated by treatment of HeLa cells
with combination of 5 or 7.5 μg/mL ETA (the inhibitor of glutathione S-transferase; the
enzyme involved in reaction of GSH with compounds) and different concentrations of 2mG
.
The cell survival was examined 72 h after. The optimal concentrations of used modulators of
GSH synthesis and glutathione S-transferase reaction were determined previously (Brozovic
et al 2008, Brozovic et al 2013, Osmak and Eljuga 1993). The activity of GSH pumps to
efflux the GSH-2mG conjugates was examined by pre-treatment of HeLa cells for one hour
with 0.2 or 0.625 mM probenecid, what was followed by treatment of HeLa cells with
different concentrations of 2mG. Seventy-two hours after, the effect of combination treatment
to 2mG treatment alone was determined.
Determination of total cell ruthenisation. Total cell ruthenisation was measured as
described previously for measurement of total cell platination (Brozovic et al 2013) with

modifications. Briefly, the cells were treated with either 150 µM of 2pG, 2pA, 2mG and 2mA
during 2-6 h or 17.5-150 µM 2pG, 2pA, 2mG and 2mA during 2 h, rinsed with ice-cold PBS, and
harvested into 10 ml of ice-cold PBS using a rubber policeman. After centrifugation, the cells
were re-suspended in PBS, an aliquot was used for determination of cell number, and the
remainder was digested in 70% nitric acid. Cell lysates were heated for 2 h at 75 °C, diluted
to 5% nitric acid, and assayed for ruthenium content. The amount of ruthenium was measured
by a validated high-resolution inductively coupled plasma mass spectrometry (HR ICPMS)
using the Element 2 (Thermo Finnigan, Germany). Calibration standards were prepared from
RuCl3 × 3H2O diluted in 1:4 hydrochloric acid and water (1000 μg/mL; Agilent, USA).
Statistical analysis. Data were analyzed by Student’s t-test, and expressed as the mean ±
standard error of the mean. Data were considered significant when P values were lower than
0.05, and in the figures these are designated as * = P < 0.05 or ** = P < 0.01. Experiments
were performed in triplicate and repeated at least twice.
Acknowledgement. These materials are based on work financed by the Croatian Science
Foundation (CSF, project numbers IP-2014-09-1461, IP-2016-06-1036 and DOK-2018-01-
8086), the Terry Fox Foundation, Croatian League Against Cancer and in part by a Croatian-
Slovenian bilateral project. The authors would like to thank Professor Maja Osmak and Dr.
Mario Cindrić (both Ruđer Bošković Institute) for helpful comments, Ernest Sanders (Ruđer
Bošković Institute) for technical assistance and COST Action CM1105 for financial support.
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