# Selection of Ruthenium Polypyridyl Complex-Modified Aptamers for Photodynamic Therapy against Streptococcus Pneumonia

Marie Flamme, Germain Niogret, Luke Mckenzie, Usman Akhtar, Fabienne Levi-Acobas, Albert Gandioso, Emma Clarke, Pierre Nicolas Bizat, Michael Connor, Mélanie A Hamon, et al.

<figure><img src="image_1.png" /></figure>

## ► To cite this version:

Marie Flamme, Germain Niogret, Luke Mckenzie, Usman Akhtar, Fabienne Levi-Acobas, et al. Selection of Ruthenium Polypyridyl Complex-Modified Aptamers for Photodynamic Therapy against Streptococcus Pneumonia. Journal of the American Chemical Society, In press, 10.1021/jacs.5c13224. [hal-05357414](https://hal.science/hal-05357414v1)

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Submitted on 10 Nov 2025

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# Selection of Ruthenium Polypyridyl Complex-modified Aptamers for Photodynamic Therapy against *Streptococcus pneumoniae*

Marie Flamme,<sup>a,b<sup>1</sup></sup> Germain Niogret,<sup>a,b<sup>1</sup></sup> Luke McKenzie,<sup>a,b</sup> Usman Akhtar,<sup>a,c</sup> Fabienne Levi-Acobas,<sup>a</sup> Albert Gandioso,<sup>b</sup> Emma Clarke,<sup>a,b</sup> Pierre Nicolas Bizat,<sup>a</sup> Michael Connor,<sup>d</sup> Mélanie A. Hamon,<sup>d</sup> Gilles Gasser,<sup>b*and</sup> Marcel Hollenstein<sup>a*</sup>

<sup>a</sup> Institut Pasteur, Université Paris Cité, CNRS UMR3523, Department of Structural Biology and Chemistry, Laboratory for Bioorganic Chemistry of Nucleic Acids, 28, rue du Docteur Roux, 75724 Paris Cedex 15, France

<sup>b</sup> Chimie ParisTech, PSL University, CNRS, Institute of Chemistry for Life and Health Sciences, Laboratory for Inorganic Chemical Biology, 75005 Paris, France

<sup>c</sup> Department of Pharmacy, Forman Christian College (A Chartered University), Zahoor Elahi Rd, Gulberg III, Lahore, 54600, Pakistan

<sup>d</sup> Institut Pasteur, Université Paris Cité, Chromatin and Infection Laboratory, 28, rue du Docteur Roux, 75015, Paris, France

<sup>1</sup> these authors contributed equally

*To whom correspondence should be addressed. E-mail: gilles.gasser@chimieparistech.psl.eu and marcel.hollenstein@pasteur.fr

## Abstract

Photodynamic therapy (PDT) harnesses the combination of light, oxygen, and photosensitizers to induce cell death via reactive oxygen species (ROS) formation. Given its intrinsic properties, PDT represents an alluring way of stymieing the increasing surge of antimicrobial resistance (AMR). Despite favorable assets, various hurdles need to be circumvented before PDT can efficiently be used to combat AMR. Here, we have evaluated the possibility of generating aptamers equipped with ruthenium polypyridyl complexes against entire Gram-positive *Streptococcus pneumoniae* bacteria. This combination is hypothesized to improve the poor specificity of photosensitizers, increase PDT efficiency, and potentially penetrate biofilms. Towards these aims, we first prepared nucleotides equipped with various ruthenium complexes and investigated their capacity at serving as substrates for polymerases for enzymatic DNA synthesis. Depending on the nature of the polypyridyl ligands, strong intercalation into dsDNA was observed even when connected to negatively charged nucleotide backbones. We then carried out SELEX and identified two unmodified aptamers that bound to the fixed bacterial target with $K_d$ values of 118 nM and 541 nM. The SELEX experiment with the ruthenium-modified nucleotide led to the identification of one aptamer. The enzymatic synthesis of the modified aptamer was complicated by the formation of a very stable secondary structure confirmed by UV melting experiments ($T_m$ of 84°C). The modified aptamer displayed a high affinity ($K_d$ value of 125 nM) for fixed *Streptococcus pneumoniae* bacteria. Collectively, these results highlight the possibility of using nucleotides equipped with large modifications such as ruthenium polypyridyl complexes in SELEX to raise potent aptamers against entire bacterial targets. These findings open directions to convert aptamers into potent devices to combat AMR via PDT-based approaches.

**Key words:** aptamers; modified nucleotides; polypyridyl complexes; *Streptococcus pneumoniae*; AMR; PDT
Introduction

Photodynamic therapy (PDT) is a clinically approved treatment modality for different types of
medical conditions including cancer. This therapeutic methodology is based on the
administration of a photosensitizer (PS) followed by its excitation by light irradiation to trigger
the formation of cytotoxic reactive oxygen species (ROS).<sup>1-3</sup> Compared to traditional treatment
modalities such as surgery, chemotherapy, and radiotherapy, PDT offers many advantages.
For instance, PDT offers a large degree of spatial and temporal control due to the selective
activation of the PS at a defined wavelength, leading to low systemic toxicity. So far, a dozen
of different PSs have been approved for clinical PDT applications.<sup>3</sup> Given the intrinsic
properties and advantages of this method, other medical conditions than cancer can be treated
by application of PDT. For instance, PDT appears to be particularly well-suited to combat
antimicrobial resistance (AMR) which is one of the most serious threats to human health.<sup>4</sup> As
PDT relies on the excitation of a photosensitizer for its mechanism of action, and not a
particular bacterial or viral target, it is particularly well suited to combat rising antimicrobial
resistance (AMR) while limiting the likelihood of acquired therapy resistance.<sup>5-6</sup> Several studies
have already demonstrated that antimicrobial photodynamic therapy (aPDT) is directly
amenable to different resistant microorganisms including bacterial strains resistant against
antibiotics.<sup>7-12</sup> Despite very promising assets, various hurdles need to be lifted before aPDT
can be applied and efficiently combat AMR on a large number of bacterial and viral targets.
For instance, Gram-negative bacteria are difficult targets for aPDT because the defensive
membrane prevents PS penetration.<sup>13-14</sup> Similarly, the capacity of bacteria to form biofilms
prevents PSs from forming ROS and requires relatively high concentrations of the
photosensitizing agents. Such high concentrations are often difficult to achieve which leads to
reduced aPDT efficiency.<sup>15</sup> Another severe limitation is the lack of selectivity of the treatment.<sup>16-17</sup>
Indeed, the PSs are often unspecific compounds, which prevents them from distinguishing
pathogenic species from human cells as well as unrelated bacteria. This lack of specificity
results in loss of efficiency of the treatment and is accompanied by undesired side-effects.<sup>9, 18-20</sup>
Some degree of specificity and selectivity can be restored by coupling PSs to vectors such
as aptamers.<sup>17</sup> Aptamers are single-stranded nucleic acid sequences capable of binding to a
broad range of targets with high affinity and specificity.<sup>21-23</sup> These functional nucleic acids are
generated by a Darwinian *in vitro* selection method called SELEX (systematic evolution of
ligands by exponential enrichment).<sup>24-25</sup> Due to their ease of selection and chemical
modification, aptamers have advanced as potent drug delivery systems and form the so-called
aptamer-drug-conjugates (ApDCs).<sup>26-30</sup> In the context of PDT, aptamers have been decorated
with organic- and metal-based PSs and shown to improve the specificity of the treatment by
guiding the photosensitizing agents to their intended targets.<sup>31-38</sup> However, application of
ApDCs to the aPDT strategy is timid and the full potential of chemical modification available to
oligonucleotides has not yet been explored in aptamer-driven PS delivery. Indeed, most
modification strategies so far involve the post-SELEX appendage of one or multiple drugs/PS
agents to the 3'/5'-termini of aptamers. While this approach does convey a degree of specificity
to PSs, it involves post-SELEX optimization steps, which often require structure-activity
relationship studies and permits the addition of only a limited number of PSs. In addition, post-
SELEX modification of existing aptamers can also result in loss of binding affinity and/or
specificity. On the other hand, modifications can also be integrated directly into the SELEX
process which abrogates the need for post-SELEX optimization and permits the integration of
multiple PSs in the aptameric scaffold. In this approach coined mod-SELEX, nucleosides
equipped with modifications at the level of the nucleobase replace one or multiple unmodified
nucleotide counterparts during enzymatic preparation of the naïve libraries.<sup>39-46</sup>

Herein, we have evolved an aptamer modified with ruthenium polypyridyl-based PSs against
the Gram-positive bacterium *Streptococcus pneumoniae*. The choice of the use of a metal-
based PS, and more specifically of ruthenium polypyridyl complexes is due to the great results
obtained with the ruthenium-based PS TLD1433, which is currently in phase II clinical trial.<sup>47-48</sup>
To do so, we first synthesized nucleoside triphosphates equipped with different ruthenium
polypyridyl complexes and linker arms. We investigated the biochemical properties of these
nucleotide analogs and observed that incorporation efficiency was strongly dependent on the
nature of the ligands on the ruthenium complex. After the SELEX experiment, we identified two
unmodified aptamers and one modified sequence that bound to S. pneumoniae with
dissociation constants (K_d values) in the 100 nM range. Collectively, this work demonstrates
the possibility of using mod-SELEX against bacterial targets and paves the way to further
improve the selectivity of aPDT to combat AMR.

**Materials and methods**

**Chemical synthesis of complexes, nucleosides, and nucleotides**

Synthesis, purification, and characterization of ligands, ruthenium complexes, and nucleoside and nucleotide analogs are all provided in the Supplementary Information. Chemical characterization of all compounds including NMR and mass spectra is also included in the Supplementary Information.

**General protocol for PEX reactions**

A 5'-FAM-labelled primer (10 pmol) was hybridized with the corresponding template (15 pmol) in DNase/RNase-free ultrapure water. This was achieved by elevating the temperature to 95°C and then allowing it to slowly return to ambient conditions over an hour. Subsequently, the designated DNA polymerase (0.5 to 1 µL), suitable reaction buffer, and the required dNTP(s) were added for a 10 µL total reaction volume. This mixture was then incubated for the given duration at the polymerase-specific optimal temperature. The temperatures for various polymerases were as follows: Phusion, Hemo KlenTaq, Taq, Bst, Q5 DNA polymerase, Therminator, Vent (exo⁻) - all at 60°C; Dpo4 and Deep Vent both at 55°C; and Phi29 and Kf (exo⁻) both at 37°C. All reactions were quenched by the addition of 10 µL of a solution containing formamide (70%), EDTA (50 mM), bromophenol (0.1%), and xylene cyanol (0.1%). The reaction products were analyzed by gel electrophoresis on a denaturing 20% polyacrylamide gel, that was run in 1x TBE buffer (pH 8) and urea (7 M) and visualized by phosphorimager analysis.

**Large scale synthesis and purification of modified sequence S1**

Following a similar protocol, 5'-FAM-labelled primer **P3** (10 nmol) was hybridized with template **T8** (15 nmol), in DNase/RNase-free ultrapure water. Taq DNA polymerase (500 U), natural dNTPs (200 µM), and **dU**<sup>Ru(bpy)TP</sup> **1** (60 µM) were then added for a total reaction volume of 100 µL. The reaction mixture was then incubated at 60°C for 3h after splitting the reaction mixture into 10 separate PCR tubes. After quenching with the same solution containing formamide (70%), EDTA (50 mM), and both dyes, the reaction products were subjected to gel electrophoresis (PAGE 20%) for 4.5 hours. After gel migration, the gel was visualized under a Bioblock Scientific UV module, TF35 L. The fluorescent band, corresponding to the desired DNA product, was carefully excised from the gel. For DNA extraction, the excised gel fragment was immersed in an elution buffer containing 0.5 M ammonium acetate and 10 mM magnesium acetate. Following elution, the solution underwent filtration, evaporation, and subsequent resuspension in distilled water. The DNA was then isolated by ethanol precipitation. Post-centrifugation, the supernatant was discarded, leaving the DNA pellet which was dried and reconstituted in distilled water. The final yield and product purity were ascertained using UV spectroscopy, with yields commonly ranging between 10-20% for the modified single-stranded DNA product.
## General protocol for PCR

The constituents for the PCR mix were as follows: primers at 5 µM each, a template at 0.01 µM, dNTP(s) at 200 µM, Mg²⁺ at 2 mM, polymerase (1 µL), and the designated polymerase buffer (either 2 µL of 10X or 4 µL of 5X), in a total volume of 20 µL. The PCR amplification protocol commenced with an initial 5-minute denaturation step at 94°C. This was succeeded by amplification (25 cycles for modified nucleotides and 10 to 15 for canonical nucleotides), each consisting of a 30 second denaturation at 94°C, annealing for 30 seconds at 52°C, and an elongation for 2 minutes at 72°C. The cycles finished with a final extension period of 5 minutes at 72°C. Following amplification, the PCR products were visualized by 4% agarose gel electrophoresis, supplemented with 1X E-GEL sample loading buffer (loading: 1 to 5 pmol).

## Fluorescent assays of ethidium bromide substitution studies

The fluorescent assays for ethidium bromide substitution studies were conducted using a high sensitivity fluorimeter (Quantamaster QM4CW, Photon Technology International). These measurements were carried out in the presence of varying concentrations of ruthenium complexes ranging from 1 to 30 µM. The concentration of EB (ethidium bromide) was 20 µM, and that of ct-DNA was 100 µM in a buffer containing 50 mM of NaCl and 5 mM of Trizma. The ruthenium complexes were gradually introduced, from 0 to 20 µM, after 10 successive additions. An excitation wavelength of 280 nm was utilized for these studies.

## Preparation of bacteria

Experimental starters of *S pneumoniae* R6⁴⁹ were generated by growing a swath of bacteria from an overnight culture on 5% Columbia blood agar plates (Biomerieux reference no. 43041) at 37°C + 5% CO₂ in Todd-Hewitt + 50mM HEPES (THH; 4-6 h). Midlog bacteria were pelleted (1500 xg; 10 minutes), and diluted to 0.6 OD₆₀₀ in 15% final THH+ glycerol stocks for freezing at -80°C.⁵⁰ To generate 4% paraformaldehyde (PFA) killed R6 for SELEX, experimental starters were grown revived and grown in THH for 3 h. Cultures were pelleted, washed in DPBS, and quantified spectrophotometrically using c.f.u / mL the conversion factor 0.6 OD₆₀₀ = 3x10⁸ c.f.u/mL. Total bacteria were incubated at room temperature in 4% PFA for 30 minutes, prior to three DPBS washes and frozen as pellets at -80°C. Fixation of bacteria was determined by plating 10 µl on 5% Columbia blood agar plates for overnight growth at 37°C + 5% CO₂. PFA inactivates *S pneumoniae* while maintaining bacterial morphology.⁵⁰

## In vitro SELEX with modified nucleotide dU<sup>Ru(bpy)TP</sup> 1

After PCR amplification (by application of the general protocol) of library L1 (10 pmoles) with natural dNTPs and primers P3 and P7 (300 pmoles) followed by conversion to ssDNA by λ-exonuclease digestion (at 37°C for 2 h), 100 pmoles were used as template in PEX reactions with dU<sup>Ru(bpy)TP</sup> 1, Vent (exo⁻), and primer P8 (67 pmoles) and following the general protocol. Next, 125 µL of a streptavidin bead solution was magnetized and the supernatant was discarded. The beads were washed 3 times with 300 µL of TEN buffer (40 mM Tris-HCl, pH 7.5, 1 mM EDTA, 150 mM NaCl). The PEX products were added to the beads and left to incubate at room temperature for 30 minutes. The resulting solution was then magnetized and the supernatant was removed. The beads were washed as follows: 3 times with 300 µL TEN buffer, 5 times with 300 µL of 0.1 M NaOH + 1 mM EDTA, once with 600 µL TEN buffer, and once with 300 µL RNase/DNase free ultrapure water. The beads were then incubated with 45
µL of NaOH 2 M and 60 µL water at 65°C for 25 minutes. The solution was neutralized by adding 90 µL of HCl 1 M followed by Nucleospin column purification. The resulting solution was dried under vacuum. The dried products were dissolved in 40 µL of 1X PBS and denatured at 95°C for 7 minutes and then cooled on ice for 5 minutes.

10⁸ fixed bacteria were suspended in 1 mL PBS 1X buffer and were centrifuged at 10000 rpm for 5 min and the supernatant was removed. 40 µL of PBS 1X buffer was added. The previously treated PEX products were added to the suspended bacteria and the resulting mixture was shaken at room temperature for 1 h (for all the 12 rounds of SELEX), depending on the round of selection. The mixture was then centrifuged at 10000 rpm for 5 min and the supernatant was removed. The bacteria were resuspended in 100 µL of PBS 1X buffer. This step was repeated three times. The bacteria were then resuspended in 75 µL water and heated at 95°C for 20 min. After centrifugation at 13000 rpm for 6 min, the supernatant was collected. The supernatants were then PCR amplified using the general protocol. PCRs products were concentrated to a volume of 50 µL. 7 µL of λ-exonuclease buffer 10X, 11 µL of water and 4 µL of λ-exonuclease polymerase (5 U) were added and the mixture was incubated at 37°C for 30 min. After Nucleospin purification, the mixture was quantified by absorbance and used as a template for subsequent rounds of SELEX. The twelfth generation was then amplified using Taq DNA polymerase with unlabeled primer **P2** and primer **P9** (2 µM) and natural dNTPs (300 µM). These amplicons were then TA-cloned using the TOPO® TA Cloning® Kit (Invitrogen) by application of the recommendation of the vendor and similar protocols.⁵¹ Briefly, after adding S.O.C. medium, TOPO® Cloning reaction mixtures (containing PCR products, topoisomerase, and competent cells) were heated at 37°C for 1 h. The transformations were then plated on LB Agar containing 50 µg/mL ampicillin and 40 mg/mL X-gal and incubated at 37°C overnight. Individual, white or light blue colonies were then picked and PCR amplified with unlabeled primer **P2** and primer **P9** (1 µM) and natural dNTPs (500 µM). PCR products of correct size were then subjected to Sanger sequencing (Eurofins) after Nucleospin column purification.

## In vitro SELEX with natural nucleotides

SELEX with unmodified nucleotides proceeded similarly as that described with modified nucleotides. However, library **L1** was directly prepared by PCR amplification with primers **P3** and **P7** and directly converted to ssDNA by λ-exonuclease digestion and was not heat denatured prior to SELEX. Incubation time with target bacteria was 1 h for rounds 1-8, and 30 min for rounds 9 and 10. After PCR preparation of the enriched 10th round of SELEX, sequencing of the products was performed at the Biomics platform of Institut Pasteur following a previously developed method.⁵² Briefly, a suitable library (average length of 245 nucleotides) was obtained using PCR Free kits from Illumina. The resulting, purified PCR products were quantified by Quant-iT PicoGreen dsDNA fluorometric quantification (Thermo Fisher Scientific, Waltham, MA, USA) and pooled for sequencing in paired-end (2x162 bases) on the Illumina MiSeq platform using the 500-cycle v2 chemistry (Illumina, San Diego, CA, USA). Sequencing of samples yielded about 700,000 paired reads each, which after phix removal led to 297,518 reads. Starting from 297,518 reads, 154,096 unique sequences were observed. Of all these reads, there are 14 unique sequences with a count greater to 1000 (Supporting Information) and the three most representative sequences are indicated in Table 2.

## Thermal melting measurements by ultraviolet spectroscopy

$T_m$ measurements were recorded on a UV-VIS CARY 3500 COMPACT PELTIER from Agilent Technologies. All experiments were performed in a PBS 1X buffer solution (137 mM NaCl, 2.7 mM KCl, 4.3 mM Na₂HPO₄, and 1.47 mM KH₂PO₄, pH = 7.4 in H₂O). The solutions contained unmodified sequence **S1** (1 µM), modified sequence **S1** (0.5 µM), or a mixture of unmodified **S1** (1 µM) and dU<sup>Ru(bpy)TP 1</sup> (60 µM) in a final volume of 100 µL. To avoid evaporation, 150
µL of paraffin oil was added on top of the solution. A control cuvette setup was used 1X PBS, into which a temperature probe was integrated. Absorbance was recorded at a wavelength of 260 nm and the heating rate was set to 1°C/min. A heating-cooling-heating cycle in the temperature range 10-95°C was applied and three cycles were carried out. The resulting melting curves were converted to hyperchromicities and were then normalized. The curve representing the first derivative of hyperchromicity as a function of temperature was subsequently plotted. The Tm values were identified at the peak of these curves. The definitive Tm values were obtained by averaging the results of the three independent experiments.

## Fluorescent binding assays

5'-FAM-labelled aptamer candidates and oligonucleotide controls were incubated with 10⁸ fixed bacteria at room temperature for 1h under constant stirring at 350 rpm. The sequence concentrations tested ranged from 0 to 1.25 µM in a 50 µL PBS solution. After incubation, the mixtures were subjected to centrifugation at 10,000 rpm, isolating the supernatant for downstream evaluation. After two washes, the first of which was retained for evaluation, bacterial samples were reconstituted in 50 µL PBS for fluorescence measurements. The fluorescence measurements were carried out using an EnVision Multilabel Plate Reader. The fluorescence detection utilized a single excitation filter with central wavelength (CWL) at 485 nm, bandwidth (BW) of 14 nm, and minimum transmission (Tmin) of 60%. The emission readings were carried out using an M520W filter. Fluorescence intensities were then plotted as a function of concentration. The ligand binding analysis function of the software SigmaPlot was used to calculate the apparent Kd of aptamers by fitting curves to the following equation:

$$Y = \frac{(B_{max} \times X)}{(K_{d} + X)}$$

where Y is the fluorescence intensity at each given concentration, Bmax is the maximum signal of the specific binding, and X is the concentration of aptamer in nM.

## Determination of binding affinity using flow cytometry

For the determination of the dissociation constants (Kd), various concentrations of 5' FAM-labelled aptamers and control sequences (0, 10, 50, 100, 200, 500, 700, 1000, and 1250 nM) were incubated with 1 × 10⁸ bacterial cells for 30 minutes. The cells were washed twice with DPBS and resuspended in DPBS for FACS analysis using the Attune NxT Flow Cytometer. The data were analyzed using FlowJo software. GraphPad Prism was employed to analyze the binding curves using nonlinear regression (curve fitting) based on a one-site total binding model. The Kd value, calculated with 95% confidence intervals, was used to determine the standard deviation (±). The Kd value of the modified aptamer S1 was also determined in the same manner using 5'-FAM-labeled and Ru-modified aptamer S1 at various concentrations (0, 10, 50, 100, 200, 500, and 700 nM). To determine whether the binding sites of aptamer S9 and Ru-modified aptamer S1 are the same or different, a competition assay was performed using an unlabelled S9 sequence with 5'-FAM-labeled aptamers (S9, modified S1, and unmodified S1). The 5'-FAM-labeled aptamers were incubated with 10⁸ bacteria for 30 minutes in the presence of unlabelled S9 at a final concentration of 250 nM. The binding assay was conducted following the same procedure described above for Kd determination. Mean fluorescence intensity was analyzed using FlowJo.

## Results

### Design and synthesis of modified nucleotides
In our design for the identification of aptamers compatible with aPDT, we opted for PSs based on ruthenium polypyridyl complexes. Compared to PSs based on organic compounds, metal complexes display alluring properties such as large Stokes shifts, chemical stability, better water solubility, and high ¹O₂ production capacities. In addition, these transition metals offer interesting scaffolds that can easily be modified to fine-tune their photochemical and photophysical properties.⁴⁷, ⁵³⁻⁵⁷ In particular, we decided to decorate nucleotides with [Ru(bpy)₂dppz]²⁺ (dU<sup>Ru(bpy)</sup>TP 1 in Fig. 1; bpy = bipyridine; dppz = dipyrido[3,2-a:2',3'-c]phenazine⁵⁸⁻⁵⁹ and [Ru(bphen)₂(bipy)]²⁺ (dU<sup>Ru(bph)</sup>TP 2 and dU<sup>Ru(bph)PEG</sup>TP 3 in Fig. 1; bphen = 4,7-diphenyl-1,10-phenanthroline)⁶⁰ due to their impressive photosensitizing properties. In order to produce nucleotides equipped with such modifications, we envisioned to conjugate the metal complexes via copper-catalysed alkyne-azide cycloaddition (CuAAC) directly on the alkyne moiety of the known nucleotide ethynyl-dUTP (EdUTP). This design follows a modification strategy previously shown to be compatible with SELEX.⁶¹⁻⁶² In addition to a simple, short linker arm connecting the complexes to the nucleobase, we also included a PEG spacer to further improve water-solubility of the rather large, hydrophobic ligands on the ruthenium complexes and potentially increase the incorporation efficiency of the modified nucleotides (vide infra).⁶³

<figure><img src="image_3.png" /><figcaption>**Fig. 1.** Chemical structures of the modified nucleotides dU<sup>Ru(bpy)</sup>TP 1, dU<sup>Ru(bph)</sup>TP 2, and dU<sup>Ru(bph)PEG</sup>TP 3.</figcaption></figure>

Consequently, we prepared ruthenium polypyridyl complexes equipped with azide moieties compatible with CuAAC. To do so, we applied a similar strategy as that used to prepare a previously reported azide functionalized version of [Ru(bpy)₂dppz]²⁺.³¹ Briefly, 2-azidoethylamine 5⁶⁴ and commercial azido-PEG₄-amine 6 were connected to 4'-methyl-2,2'-bipyridine-4-carboxylic acid (bpyAc) 4⁶⁵ by application of standard amide bond formation conditions (Scheme 1, Scheme S1, and Supporting Information). In parallel, complex 9 had to be synthesized to accommodate the modified bpyAc ligands in the coordination sphere. The
preparation of complex 9 first involved the replacement of the chloride ligands of commercially available ruthenium chloride trihydrate (RuCl₃·3H₂O) by DMSO to avoid cis-trans isomerization. The DMSO ligands of the resulting complex ((CH₃)₂SO)₄RuCl₂ were then replaced by bathophenanthroline (bph). This exchange leads to the formation of the desired complex [Ru(bph)₂Cl₂]²⁺ (compound 9 in Scheme 1).⁶⁶ Ligands 7 and 8 were then complexed to 9 in EtOH/H₂O solution, resulting in the two ruthenium azide-precursors 10 and 11.

<figure><img src="image_4.png" /></figure>

**Scheme 1.** Synthesis of ruthenium polypyridyl complexes containing azide functional groups. Reagents and conditions: i) EDC, NHS, Et₃N, anhydrous DCM, rt, 15 h, 62% for 7 and 45% for 8; ii) Bathophenanthroline, LiCl, DMF, reflux, 12 h, 69%; iii) EtOH/H₂O (8:2), 70°C, 15 h, 53% for 10 and 42% for 11.

With the suitable azide-modified metal complexes in hand, we carried out CuAAC with EdUTP by application of previously reported protocols (Scheme S2).³¹, ³⁹, ⁶¹ Nonetheless, worthy of note, these reactions are carried out under strictly inert atmosphere to maintain good conversion yields. Furthermore, the reactions must be conducted in a solvent mixture with a ratio of 6/2/2 DMF/MeCN/H₂O to ensure optimal solubility of all reactants. After a through HPLC purification protocol, dU²R²u(bpy)TP 1, dU²R²u(bph)TP 2, and dU²R²u(bph)PEGTP 3 could be obtained in moderate to good yields (20% for 1, 60% for 2, and 56% for 3). Importantly, these modified nucleotides displayed important stability and no degradation was observed over longer storage periods.

## Biochemical characterization of modified nucleotides

With the modified nucleotides in hand, we next sought to evaluate their compatibility with enzymatic DNA synthesis. To do so, we carried out primer extension (PEX) reactions using two primer/template systems of different lengths. A first, system involved the 15 nt long, 5'-
FAM-labelled primer **P1** along with the 31-mer template **T1** (see Supporting Information for sequence composition), which permits the incorporation of up to four modified dUMPs. The second system was designed to encompass up to 15 different modified nucleotides and consisted of the 19 nt long, 5'-FAM-labelled primer **P2** along with the 79-mer template **T2** (see Supporting Information for sequence composition). In addition to these systems, we evaluated the capacity of twelve different DNA polymerases to incorporate the modified nucleotides. Gel analysis of the reaction products revealed that **dU**<sup>Ru(bpy)TP 1</sup> was accepted as a substrate by several polymerases including Phusion, Hemo KlenTaq, Bst, Therminator, Vent (exo<sup>-</sup>) and Deep Vent (exo<sup>-</sup>) with both primer/template systems (Supplementary Fig. S1). In these reactions, the primer is completely consumed, and full-length products were obtained, accounting for 15 ruthenium polypyridyl complexes on a single oligonucleotide. This is a remarkable feat since so far only one<sup>67</sup> or four<sup>68</sup> ruthenium-modified nucleotides have been incorporated into DNA.<sup>69</sup> These results also suggest the possibility of using **dU**<sup>Ru(bpy)TP 1</sup> in SELEX since longer oligonucleotides are accessible by enzymatic synthesis. Unexpectedly, gel analysis of the reaction products obtained with **dU**<sup>Ru(bph)TP 2</sup> as a substrate displayed a completely different pattern than that observed not only with **dU**<sup>Ru(bpy)TP 1</sup> but also with other modified nucleotides (Supplementary Fig. S2). Indeed, while we noticed full consumption of the primer in most reactions, no clear band corresponding to full-length product could be detected. Instead, we observed the presence of truncated products for some of the reactions while others led to products that were retained in the wells of the gels and appeared to be incapable of gel migration (indicated by red arrows in Supplementary Fig. S2B). To rationalize these findings, we initially believed that the higher lipophilicity imparted by the larger bph ligands<sup>70-71</sup> combined with increased steric bulk led to limited solubility of the nucleotide, reduced recognition by the polymerase, and low electrophoretic mobility of the reaction products. Hence, to address these hypotheses, we carried out PEX reactions with **dU**<sup>Ru(bph)PEGTP 3</sup> equipped with a PEG linker, which was believed to alleviate some of these limitations. However, gel analysis of PEX reactions with both primer/template systems revealed a similar pattern to that observed with **dU**<sup>Ru(bph)TP 2</sup> (Supplementary Fig. S3), suggesting that other factors were involved. A similar outcome was observed when a slightly shorter primer/template system **P3/T3** was used (Fig. S3A).

In order to shed light into the differential substrate behavior of nucleotides equipped with
[Ru(bpy)₂dppz]²⁺ from those with [Ru(bphen)₂(bipy)]²⁺, we carried out PEX reactions with
different nucleotide concentrations (Supplementary Fig. S4). Gel analysis reveals a threshold
concentration (~200 µM) of dU<sup>Ru(bpy)TP 1</sup>, which leads to DNA polymerase inhibition. This
analysis also confirms that substrate tolerance of dU<sup>Ru(bph)TP 2</sup> is independent of its
concentration and that the reactions lead to products that lost their electrophoretic mobility. We
next carried out single nucleotide PEX reactions to investigate whether dU<sup>Ru(bph)TP 2</sup> and
dU<sup>Ru(bph)PEGTP 3</sup> were recognized as substrates by polymerases at all. For that, we used a
simple system consisting of the 5'-FAM-labelled, 19 nt long primer P4 and the 20-mer template
T4 (see the Supporting Information for the sequence composition). Gel analysis of the products
of these PEX reactions (Supplementary Figs. S5 and S6) clearly showed that i) both Ru-
modified nucleotides were incorporated into the primer, ii) long reaction times (3 h) were
required, and iii) the presence of [Ru(bphen)₂(bipy)]²⁺ induced an impressive gel mobility
retardation compared to reaction products obtained with unmodified dTTP. Finally, we carried
out PEX reactions with dU<sup>Ru(bph)PEGTP 3</sup> on six primer/template systems differing in terms of
sequence composition and number of possible incorporation sites. We also analyzed the
products by agarose gel electrophoresis (Fig. 2, B1/B2), which appears to be superior to PAGE
(Fig. 2A) for the detection of such substrates. These experiments clearly indicate that
dU<sup>Ru(bph)PEGTP 3</sup> is a substrate for Vent (exo<sup>-</sup>), leading to full length products in all systems
(Supplementary Fig. S7). However, it becomes apparent that the presence of as few as four
modified nucleotides completely abrogates gel electrophoretic mobility on PAGE, but not on
agarose gels. Also, products obtained with longer templates (i.e. T2 and T3) appear to consist
of different species as highlighted by the observed smears on agarose gels.
Taken together, the biochemical analysis of the three modified nucleotides clearly reveals that all these analogs are substrates for DNA polymerases but to various degrees. Chemical differences between the ligands on the complexes lead to important differences in gel mobility and potentially in enzymatic DNA synthesis. Given these differences, we opted to employ **dU<sup>Ru(bpy)</sup>TP 1** in mod-SELEX, which appears to be most suitable for enzymatic DNA synthesis and analysis of reaction products.

<figure><img src="image_5.png" /></figure>

**Fig. 2.** A) PAGE gel (20%) analysis of PEX reactions (3 h) using primer P3 (10 pmol) and template T3 (15 pmol). Reaction mixtures contained natural A, C, G nucleotides (200 µM final), each of the modified triphosphates (100 µM final), and 1 µL of Vent (exo⁻) (2 U) polymerase. The following nucleotides were evaluated: lane 1) **dU<sup>Ru(bpy)</sup>TP 1**; lane 2) **dU<sup>Ru(bph)</sup>TP 2**; lane 3) **dU<sup>Ru(bph)</sup>PEG<sup>1</sup>TP 3**; lane 4) positive control with natural dNTPs and Vent (exo⁻). The 19 nucleotide (nt) mark corresponds to primer P3, 71 nt to that of the unmodified full-length product, and 71 nt* to that of the modified full-length product. **B1-B2)** Agarose gel analysis (2% and 4%, respectively) of PEX reaction products shown in lanes 1 to 4 of the PAGE gel of this figure along with two additional controls consisting of 100 µM of unreacted **dU<sup>Ru(bph)</sup>TP 2** (lane 5) and **dU<sup>Ru(bph)</sup>PEG<sup>1</sup>TP 3** (lane 6).
Ruthenium(II) polypyridyl complexes have received considerable attention due to their ability to interact with dsDNA, often via intercalation or electrostatic interaction, which is seen as a key factor in their modes of biological action.⁷²⁻⁸⁰ However, to the best of our knowledge, the interaction of ruthenium(II) polypyridyl complexes-nucleotide conjugates with DNA has never been investigated. Evaluating a possible interaction is also important to understand the observed differences in incorporation efficiencies between the three nucleotides and avoid difficulties in strand dissociation during SELEX. Hence, we sought to establish whether dU<sup>Ru(bpy)TP 1</sup> and dU<sup>Ru(bph)TP 2</sup> could interact with double-stranded DNA via ethidium bromide (EB) displacement studies with calf thymus DNA (ct-DNA).⁸¹⁻⁸³ We also included the free complexes in this study to evaluate the effect of the presence of the negatively charged nucleotide backbone on the intercalation efficiency.

Following a series of analyses of emission and excitation spectra (data not shown), we opted for an excitation at 280 nm for subsequent experiments since the ruthenium complexes do not produce any significant emission between 500 and 800 nm when excited at this wavelength, thus minimizing any potential interference with the **EB** signal. Substitution experiments were carried out in the presence of different concentrations of the complexes [Ru(bpy)<sub>2</sub>dppz-N<sub>3</sub>]<sup>2+</sup> (Fig. 3A), dU<sup>Ru(bpy)TP 1</sup> (Fig. 3B), [Ru(bphen)<sub>2</sub>(bipy)-N<sub>3</sub>]<sup>2+</sup> **10** (Fig. 3C), and dU<sup>Ru(bph)TP 2</sup> (Fig. 3D). For each curve, the concentration of **EB** was set at 20 µM and that of ct-DNA at 100 µM ([ct-DNA]/**EB** = 5:1). Ruthenium complexes were added gradually, from 0 to 30 µM, over 10 successive additions.

<figure><img src="image_6.png" /><figcaption>**Fig. 3.** Emission fluorescence spectra obtained through ethidium bromide substitution by the ruthenium complexes in a solution containing 100 µM ct-DNA, 20 µM EtBr, 50 mM NaCl, and 5 mM Trizma. The inserts in each curve show the quenching plots, based on the linear Stern-Volmer equation, for the determination of K<sub>sv</sub>.</figcaption></figure>

The quenching constants (K<sub>sv</sub>) were calculated according to the linear Stern–Volmer (SV) equation.⁸²

$$ \frac{I_0}{I} = 1 + K_{SV} \cdot r $$

Where I₀ and I represent the fluorescence intensities in the absence and presence of the complexes, respectively and r represents the ratio of total complex concentration to DNA
concentration ([Ru]/[ct-DNA]).<sup>82</sup> The K<sub>sv</sub> constant serves as a comparative parameter for the displacement of **EB** from dsDNA and higher values indicate a stronger interaction with dsDNA. We have also used the known binding constant of **EB** for DNA (1.4 x 10<sup>6</sup> M<sup>-1</sup>)<sup>84-85</sup> to calculate the apparent binding constant (K<sub>app</sub>) of the complexes with the following formula:<sup>84, 86</sup>

$$K_{app} = K_{EB} \cdot \left( \frac{[EB]_{50\%}}{[Ru]_{50\%}} \right)$$

where K<sub>app</sub> is the apparent binding constant between the complex and dsDNA, while K<sub>EB</sub> represents the DNA binding constant for ethidium bromide. The concentrations required to achieve a 50% reduction in fluorescence are noted as [EB]<sub>50%</sub> and [Ru]<sub>50%</sub> for ethidium bromide and ruthenium complexes, respectively.

<table><caption>**Table 1.** Summary of binding (K<sub>app</sub>) and quenching (K<sub>sv</sub>) constants for the interaction of complexes with ct-DNA</caption><thead><tr><th>Complex</th><th>K<sub>sv</sub></th><th>[Ru]<sub>50%</sub><br>[EB]<sub>50%</sub></th><th>K<sub>app</sub> (M<sup>-1</sup>)</th></tr></thead><tbody><tr><td>[Ru(bpy)<sub>2</sub>dppz-N<sub>3</sub>]<sup>2+</sup></td><td>183.94</td><td>0.12</td><td>1.15 x 10<sup>7</sup></td></tr><tr><td>dU<sup>Ru(bpy)TP 1</sup></td><td>16.43</td><td>0.69</td><td>2.04 x 10<sup>6</sup></td></tr><tr><td>[Ru(bphen)<sub>2</sub>(bipy)-N<sub>3</sub>]<sup>2+</sup> 10</td><td>18.28</td><td>0.72</td><td>1.94 x 10<sup>6</sup></td></tr><tr><td>dU<sup>Ru(bpy)TP 2</sup></td><td>5.63</td><td>1.01</td><td>1.39 x 10<sup>6</sup></td></tr></tbody></table>

These studies clearly confirm the excellent affinity of [Ru(bpy)<sub>2</sub>dppz-N<sub>3</sub>]<sup>2+</sup> for dsDNA<sup>58, 87</sup> since both constants are ~10-fold larger than those for [Ru(bphen)<sub>2</sub>(bipy)-N<sub>3</sub>]<sup>2+</sup> 10 (Table 1). Interestingly, the appendage of a nucleoside triphosphate moiety onto a ligand of these complexes significantly reduced the affinity for dsDNA (~10-fold reduction of both constants for [Ru(bpy)<sub>2</sub>dppz-N<sub>3</sub>]<sup>2+</sup> and ~2 to ~4-fold for [Ru(bphen)<sub>2</sub>(bipy)-N<sub>3</sub>]<sup>2+</sup> 10, Table 1). Nonetheless, the apparent binding constants for nucleotides 1 and 2 are still ~10-fold higher than those reported for other Ru(II) bipyridyl complexes,<sup>83-84, 88-89</sup> suggesting a stronger interaction with dsDNA (Table 1). This affinity for dsDNA might be responsible for the reduction in enzymatic DNA synthesis observed in PEX reactions when concentrations are increased (Supplementary Fig. S4).

## Compatibility with SELEX

Biochemical characterization and intercalation studies have allowed us to identify dU<sup>Ru(bpy)TP 1</sup> as a suitable candidate for the preparation of modified libraries for SELEX as well as to refine the optimal conditions for enzymatic DNA synthesis. However, prior to carrying out SELEX, we needed to first ascertain 1) that the modified nucleotide could be used for the preparation of naïve libraries and 2) that modified libraries could be converted to unmodified DNA using PCR and unmodified dNTPs.

To do so, we first evaluated the compatibility of dU<sup>Ru(bpy)TP 1</sup> with PCR-mediated enzymatic synthesis since this would shorten the SELEX protocol. Therefore, we conducted PCR with template T3 and flanking primers P3 and P5 in the presence of several DNA polymerases. Agarose gel electrophoresis analysis of the reaction products (Supplementary Fig. S8) did not reveal any amplification of the template when the modified nucleotide was used instead of dTTP. Hence, dU<sup>Ru(bpy)TP 1</sup> does not appear to be compatible with enzymatic DNA synthesis under PCR conditions. Since we had established that the modified nucleotide was compatible
with several DNA polymerases under PEX reaction conditions, we next evaluated the
possibility of converting modified into unmodified DNA. To do so, we first carried out a PEX
reaction with dU<sup>Ru(bpy)TP</sup> 1 and Vent (exo<sup>-</sup>) as polymerase using template T2 along with the 5'-
phosphorylated version of primer P2 (P2P). The resulting modified double-stranded DNA was
then converted to single-stranded DNA by λ-exonuclease digestion. Using the modified ssDNA
as template, we carried out PCR with primers P2 and P5. Gel analysis (agarose 2%) clearly
revealed the presence of a strong band when the ruthenium-modified oligonucleotide was used
as template (lane 1 in Supplementary Fig. S9A) and which displayed an identical gel mobility
compared to that of the product obtained with unmodified dNTPs and template T2. Finally, we
carried out a similar experiment using a randomized library instead of a defined oligonucleotide
sequence. The library L1 consists of a 35 nucleotide long randomized region (corresponding
to 4<sup>35</sup> ≈ 10<sup>21</sup> possible sequence combinations) flanked by binding sites for primers P3 and P7
which is equipped with a 5'-phosphate handle.<sup>61</sup> Agarose gel analysis (Supplementary Fig.
S9B) also confirmed efficient PCR amplification when modified library L1 was used as a
template.

*In vitro* selection of aptamers

Infections by gram-positive *Streptococcus pneumoniae* bacteria can lead to life-threatening
diseases including pneumonia and meningitis. These diseases combined cause > 1 million
deaths worldwide.⁹⁰ Since pneumococcal infections are principally treated with antibiotics, a
strong increase in resistance to common drugs has been observed for numerous strains of S.
*pneumoniae*.⁹¹⁻⁹³ Consequently, S. pneumoniae appears as an appropriate target for the
identification of PS-modified aptamers. Particularly, we opted for the well-known and described
laboratory R6 strain which lacks the capsule as proof-of-principle.⁴⁹ After growing of S.
pneumoniae strain R6 under standard conditions,⁹⁴ the bacteria were fixed with 4%
paraformaldehyde (PFA).⁹⁵ This fixation method allows all extracellular proteins and structures
to remain associated to the bacterial cell. The resulting fixed bacteria were then subjected to
a mod-SELEX protocol that is inspired by a report from the Perrin lab, who identified the first
base-modified aptamer against a bacterium (Fig. 4).⁹⁶ Briefly, library L1 was PCR amplified
with primers P3 and P7 and converted to ssDNA by λ-exonuclease digestion. The resulting
library served as a template for PEX reactions with dU<sup>Ru(bpy)TP</sup> 1, Vent (exo<sup>-</sup>), and primer P8
equipped with a 5'-biotin and containing three ribonucleotides in the linker arm (see the
Supporting Information for sequence composition). The presence of the 5'-biotin allowed to
capture only the modified library obtained by PEX reaction via immobilization on streptavidin-
coated magnetic particles. After several washes and mild NaOH treatment to remove the
unmodified template strand, the modified library is recovered from the magnetic particles by a
more intensive NaOH treatment, which cleaves the ribophosphodiester linkages.⁹⁶ After
column purification, modified ssDNA (100 pmol) was added to 10⁸ fixed bacteria and the
mixture was shaken for 1 hour. After incubation and centrifugation, the unbound sequences
were removed by several washes. The pellet was resuspended and denatured at 95°C for 20
min followed by centrifugation, in order to elute the bound sequences. The single stranded
modified DNA was then amplified and converted to natural, double stranded DNA by PCR,
using primers P3 and P7. Presence of the phosphorylated group allowed the recuperation of
the strand of interest by λ-exonuclease digestion. The single stranded natural DNA was then
quantified and reintroduced into another round of selection. In parallel to mod-SELEX, we also
conducted an *in vitro* selection experiment with unmodified DNA. In addition to providing a
direct comparison to modified sequences, unmodified aptamers could be used as diagnostic
tools and/or PS-delivery systems. SELEX with unmodified DNA followed standard protocols
and involved the same L1 library but PCR production with primers P3 and P7 rather than primer
P8 used in the mod-SELEX experiment (see the Supporting Information for experimental
details).
After several adaptations and revisions, including refinement of PCR conditions and modulation of the number of cycles, a total of 10 rounds were carried out with standard DNA chemistry and 12 for the mod-SELEX experiment without counter-SELEX steps (see Supplementary Figs. S10 and S11, respectively). NGS sequencing of generation 10 of the SELEX with unmodified DNA was performed. Sequence analysis revealed the convergence of the selection with the detection of 26 families of sequences that are overexpressed in generation 10. A minimum of 1000 reads and a sequence similarity of 90% were required for the selection of cluster families (see Supplementary Fig. S19). The most representative sequences of the three main families identified by sequencing were chosen for further biological testing (sequences **S8**, **S10**, and **S12** in Table 2).

The population of round 12 stemming from the mod-SELEX experiment was analysed via TA-cloning followed by Sanger sequencing of 34 individual clones.<sup>51,96</sup> Analysis of the sequencing data revealed that sequence **S1** (Table 2) was present in 20% of all clones, suggesting significant enrichment of the initial library for this sequence.

<figure><img src="image_7.png" /><figcaption>**Fig. 4.** Schematic representation of the Mod-SELEX protocol used with libraries prepared with dU<sup>Ru(bpy)TP</sup> 1 against fixed S. pneumoniae R6.</figcaption></figure>

**Table 2.** Sequence composition of aptamer candidates identified by sequencing after SELEX.<sup>a</sup>
<table><thead><tr><th>Name</th><th>Sequence</th><th>Relative abundance (%)<sup>b</sup></th></tr></thead><tbody><tr><td>S8</td><td>5'-CGT ACG GTC GAC GCT AGC ACA CCC CCC AGG ATC ATA AAT TCT CTC GCT GTA GCC ACG TGG AGC TCG GAT GC-3'</td><td>3,8</td></tr><tr><td>S10</td><td>5'-CGT ACG GTC GAC GCT AGC ACC AGC CAT AGT CAC ATC AAA ACT AAC TCA CAT TCC ACG TGG AGC TCG GAT GC-3'</td><td>3,0</td></tr><tr><td>S12</td><td>5'-CGT ACG GTC GAC GCT AGC CCC ATA CTC ATC ACC ATT CAC ATC ACT CAC CTA GCC ACG TGG AGC TCG GAT GC-3'</td><td>2,9</td></tr><tr><td>S1</td><td>5'-GGA TCC GAG CTC CAC GTG GCT AGG TGA GTG ATG TGA ATG GTG ATG AGT ATG GGG CTA GCG TCG ACC GTA CG-3'</td><td></td></tr></tbody></table>

## Synthesis and characterization of the modified sequence

After identification of families of oligonucleotides and interesting sequence motifs, the most promising candidates are typically produced by solid-phase DNA synthesis prior to evaluating their binding affinities. While this approach is straightforward for unmodified DNA, the mod-SELEX experiment led to the identification of the 71 nt long aptamer candidate **S1**, which contains 13 heavily modified $dU^{Ru(bpy)}$ nucleotides and represents a challenging target for solid-phase synthesis. A valid alternative to produce enough of modified sequence **S1** is large-scale enzymatic synthesis via PEX reactions. In order to assess the compatibility of $dU^{Ru(bpy)}$TP 1 with such a protocol, we first identified the optimal nucleotide concentration (60 µM) required for the high yielding synthesis of the expected full-length product using template **T8** (corresponding to the reverse complement of **S1**) and primer **P3** (see Supplementary Fig. S12). Next, we carried out a screening experiment to identify the polymerase, which led to the highest conversion yields on this particular sequence (see Supplementary Fig. S13). Interestingly, Taq DNA polymerase led to the best results in terms of conversion yields and absence of truncated species both at low (10 pmol) and slightly higher (100 pmol) scale (see Supplementary Fig. S13A and S13B, respectively). Finally, we evaluated three different methods for the removal of the unmodified template **T8** and recovery of the modified sequence **S1**, i.e., λ-exonuclease digestion of the 5'-phosphorylated version of template **T8**, immobilization of duplex after PEX reaction via a 5'-biotinylated version of template **T8** followed by recovery of the product by mild NaOH washes, and PAGE gel purification (see Supplementary Fig. S15). Of the three methods, PAGE gel purification appeared to be the most suitable since PEX reactions on the 10 nmol scale could be purified (see Supplementary Fig. S14) and up to 3 nmoles of expected product could be obtained. Surprisingly, during these optimization steps, we observed the consistent formation of a faster running band on PAGE and agarose gel electrophoresis, even during quality control experiments using previously gel purified sequences (see Supplementary Fig. S15). Intrigued by these observations, we subjected purified Ru-modified sequence **S1** to a thermal UV-melting experiment along with the corresponding unmodified ssDNA version of sequence **S1** (Fig. 5).

<footnote><sup>a</sup>Primer binding regions are highlighted with green and blue letters and red bold-face **Ts** indicate the position of the modified nucleotides. <sup>b</sup>Determined as (Reads/Total reads) x 100.</footnote>
<figure><img src="image_8.png" /><figcaption>**Fig. 5.** Representative UV-melting curves (heating ramps only) monitored at 260 nm for single-stranded **dU**<sup>Ru(bpy)</sup>-modified **S1** (graph **A1**) and the corresponding first derivative (graph **A2**); unmodified ssDNA version of sequence **S1** (graph **B1**) and the corresponding first derivative (graph **B2**); unmodified ssDNA version of sequence **S1** but the presence of 15 µM of **dU**<sup>Ru(bpy)TP 1</sup> (graph **C1**) and the corresponding first derivative (graph **C2**). All experiments were carried out in PBS 1x (100 µL) and at 0.5 µM for **dU**<sup>Ru(bpy)</sup>-modified **S1** and 1 µM for unmodified ssDNA version of sequence **S1**.</figcaption></figure>

Interestingly, the melting curve of the Ru-modified sequence **S1** displayed a typical sigmoidal transition, suggesting the formation of a stable structure (Fig. 5A). What is more, a very high $T_m$ value (83.7 ± 0.6°C) could be determined for this highly modified single-stranded oligonucleotide. The unmodified version of sequence **S1** (obtained by a similar PEX-based protocol) also displays a sigmoidal transition, suggesting the formation of a secondary, stable structure (Fig. 5B). The influence of the Ru polypyridyl complexes on the formation of a stable secondary structure is undeniable since a dramatic decrease in thermal destabilization was observed for the unmodified sequence compared to the modified counterpart ($T_m$ = 43.5 ± 1.0°C; $\Delta T_m$ = 40.2 °C). Since the addition of **dU**<sup>Ru(bpy)TP 1</sup> to a solution of unmodified sequence **S1** precludes the formation of any stable secondary structure (Fig. 5C), the Ru polypyridyl complexes need to be covalently connected to the nucleotidic scaffold to exert their stabilization effect. Overall, these thermal denaturation experiments confirm the observation of a faster running band on PAGE, which can be ascribed to the formation of a stable secondary structure.

## Binding affinity determination of aptamers

After the successful SELEX experiments, we next sought to evaluate the binding capacity of unmodified and modified sequences. To do so, we first determined whether unmodified aptamer candidates could recognize and bind fixed *S. pneumoniae* R6 using a fluorescent binding assay. To do so, we evaluated the binding capacity of the three most abundant sequences **S8**, **S10**, and **S12** (Table 2) along with the corresponding truncated sequences **S9**, **S11**, and **S13** devoid of the primer binding regions (Supporting Information Table S1). For this evaluation, we employed a similar strategy as that outlined for the *in vitro* selection followed by quantification of fluorescence. We also included in this study a previously identified aptamer (Lyd-3) against *S. pneumoniae* strain 262 (Klein)⁹⁷ and two unmodified templates (**T2** and **T9**) as control sequences. All sequences were equipped with a 5'-FAM tag for detection purposes. Briefly, aptamer candidates are incubated with 10⁷ fixed bacteria in PBS buffer for one hour at
room temperature. After centrifugation and removal of unbound sequences, bacteria-aptamer complexes were subjected to two subsequent rounds of binding and centrifugation. The fluorescence intensity of all collected fractions was then determined using a spectrofluorometer (Supporting Information Figure S16).⁹⁷ This analysis clearly revealed that sequences **S9** and **S11**, the truncated versions of sequences **S8** and **S10**, respectively, efficiently bound to *S. pneumoniae* R6 in a concentration dependent manner. In addition, both sequences **S12** and **S13** displayed similar yet reduced binding capacity compared to **S9** and **S11**. Interestingly, parent sequences **S8** and **S10** did recognize *S. pneumoniae* R6 as target since an increase in fluorescent intensity could be observed with higher DNA concentration but were even less efficient at binding than **S12** and **S13**. Based on this initial screen, we conducted a more thorough characterization of sequences **S9** and **S11**. To do so, we first repeated the determination of the binding saturation curves of both aptamers but with 10⁸ instead of 10⁷ bacteria, which is identical to the conditions used during SELEX (Supporting Information Figure S17). This additional binding assay confirmed that both sequences were capable of binding to the *S. pneumoniae* R6 target with **S9** displaying more favorable properties than **S11**. We then determined the *K<sub>d</sub>* values of both truncated sequences **S9** and **S11** against 10⁸ fixed bacteria in PBS buffer (Fig. 6). This analysis revealed that i) binding curves adopted similar shapes to that of previously identified aptamer Lyd-3 against *S. pneumoniae* strain 262 (Klein)⁹⁷, ii) **S9** displayed better binding capacity than **S11** (*K<sub>d</sub>* value of 118 nM compared to 541 nM), iii) affinities compared favorably to that of the best binder Lyd-3 selected against *S. pneumoniae* strain 262 (Klein),⁹⁷ and iv) the *K<sub>d</sub>* values obtained were higher than that of recently identified pan-binding aptamers isolated against mixtures of *S. pneumoniae* strains.⁹⁸

<figure><img src="image_9.png" /><figcaption>**Fig. 6.** Binding saturation curves of aptamers against 10⁸ fixed *S. pneumoniae* R6 in PBS buffer. A) binding efficiency of sequence **S9**; *K<sub>d</sub>* = 118 ± 1 nM (R² = 0.97); B) binding efficiency of sequence **S11**; *K<sub>d</sub>* = 541 ± 1 nM (R² = 0.95). Experiments were carried out in triplicate with different bacterial samples.</figcaption></figure>

We next sought to determine the binding specificities of both unmodified aptamer sequences **S9** and **S11**. To do so, we carried out similar fluorescent binding assays but with different bacteria. We use *S. pneumoniae* strain 6B and TIGR4, which unlike SELEX target strain R6, they are equipped with a polysaccharide capsule, and we used an unrelated bacterium *E. coli*,⁹⁹ ¹⁰⁰⁻¹⁰¹ This analysis clearly revealed that none of the aptamers **S9** and **S11** displayed any propensity at binding to *E. coli* (Supporting Information Figures S18A and S18D), even at high DNA concentrations (Supporting Information Figure S18E). These findings are not entirely unexpected since *E. coli* is a gram-negative bacterium, which displays a radically different envelope both in terms of structure and composition compared to that of the gram-positive pathogens from *S. pneumoniae*. Similarly, only a very limited binding to *S. pneumoniae* strain 6B was observed for both aptamers **S9** and **S11** (Supporting Information Figures S18C and S18D), even at high DNA concentrations (Supporting Information Figure S18E). On the other hand, both sequences **S9** and **S11** displayed a degree of recognition for *S. pneumoniae* strain TIGR4 (Supporting Information Figures S18B and S18D), especially at higher DNA concentrations (Supporting Information Figure S18E). A similar pan-specific recognition
capacity was recently observed in a cell-SELEX approach, where a mixture of different strains of S. pneumoniae bacteria was used as target.⁹⁸ These fluorescent binding experiments allowed the evaluation of both the affinity and the specificity of aptamers S9 and S11, but are not compatible with Ru-modified sequence S1, which is difficult to produce on large scale. Hence, we switched to a flow cytometry-based method to evaluate whether Ru-modified sequence S1 was capable of binding to target bacteria. To do so, we adopted a recently developed flow cytometry-based method for the binding determination of chemically modified aptamers⁴⁰ to bacterial rather than protein targets. We first evaluated the binding capacity of the best unmodified binder, i.e., sequence S9 against 10⁸ fixed S. pneumoniae R6 in PBS buffer (Fig. 7A). By application of this method, a K_d value of 55 ± 10 nM could be determined for S9, which is in the range of what was obtained by fluorescent binding assays. Next, we applied a similar strategy to the modified sequence S1, which revealed that this Ru-modified oligonucleotide was capable of binding to S. pneumoniae R6 with slightly lower affinity than the best, unmodified sequence (K_d = 125 ± 91 nM; Fig. 7B). However, the error on the measurement of the K_d value was rather important because we observed the formation of large aggregates (Supporting Information Figure S20), even at concentrations as low as 100 nM (Fig. S20D). The formation of such large aggregates is most likely triggered by the presence of the thirteen ruthenium polypyridyl complexes, which are known to self-assemble into nanoaggregates.¹⁰²⁻¹⁰³

Finally, we strived to compare unmodified sequence S9 and modified aptamer S1 for their
binding capacity as well as the necessity of the presence of the modifications for activity. To
do so, we first incubated 10⁸ fixed S. pneumoniae R6 with 250 nM of S9, modified S1, and
unmodified S1 (where dU<sup>Ru(bpy)</sup>MPs were substituted with canonical dTMPs) and measured
the mean fluorescence intensity of the resulting complexes by flow cytometry (Fig. 7C). This
analysis clearly revealed that i) in the absence of the Ru modifications, sequence S1 nearly
lost all binding activity suggesting that the complexes play are important for binding and ii) that
S9 bound better to the bacterial target than modified sequence S1 as reflected by the
dissociation constants. Nonetheless, when bacteria were pre-incubated with unlabeled S9 and
then allowed to interact with fluorescently tagged S1 and S9, similar fluorescence intensities
were measured in both instances (Fig. 7C). This suggests that both 5'-FAM-labeled S1 and S9
are capable of displacing unlabeled S9.

<figure><img src="image_10.png" /><figcaption>**Fig. 7.** Evaluation of the binding affinity of unmodified aptamer sequence S9 and Ru-modified aptamer S1 against 10⁸ fixed S. pneumoniae R6 in PBS buffer by flow cytometry. A) Binding curve of aptamer S9, which was analyzed using GraphPad Prism with nonlinear regression (curve fitting) based on a one-site total binding model. The calculated K_d values and their 95% confidence intervals were used to determine the standard deviation. B) Binding curve of Ru-modified aptamer S1, which was analyzed using GraphPad Prism with nonlinear regression (curve fitting) based on a one-site total binding model. The calculated K_d values and their 95% confidence intervals were used to determine the standard deviation. C) Competition binding assay of aptamer S9 with Ru-modified aptamer S1 for S. pneumoniae R6. Experiments were carried out in triplicate with different bacterial samples.</figcaption></figure>
Discussion

Beyond their potential as diagnostic tools for detecting and monitoring bacterial infections, aptamers are interesting candidates to combat AMR. Aptamers are indeed capable of guiding drugs to intended locations and, under certain circumstances, can even internalize into cells/bacteria and lead to inhibition of growth.<sup>104-105</sup> Moreover, by partial re-selection or by a new SELEX experiment, aptamers can easily adapt and readjust to changes that the target bacteria may have adopted to elude treatments. Combining these advantages with efficient PDT agents might yield even a better solution to combat the impending threat of AMR. With this aim in mind, we sought to develop aptamers against bacterial targets equipped with multiple Ru polypyridyl complexes, which display excellent photophysical properties for PDT. This approach would not only circumvent the need for post-SELEX modification of aptamers with Ru-based PS agents, which often includes a risk of loss of affinity/specificity but also permits the inclusion of multiple rather than just one PS agent. We also hypothesize that the inclusion of multiple Ru complexes might lead to higher local ROS concentrations and thus more efficient PDT, which in turn would involve injecting less PS agents to patients. However, prior to reaching these aims, a number of challenges had to be met. First, due to the presence of large aromatic ligands, Ru polypyridyl complexes are rather hydrophobic and excellent DNA intercalators even when connected to nucleoside triphosphates. This high propensity at intercalating dsDNA is an important impediment to enzymatic DNA synthesis. Notwithstanding, we identified one nucleoside triphosphate decorated with a [Ru(bpy)₂dppz]²⁺ complex that was compatible with polymerase-mediated synthesis. Second, SELEX with such a heavily modified nucleotide revealed to be challenging due to the moderate substrate tolerance by polymerases and the more complex set-up required when using PEX reactions. Third, while enzymatic production of sequence **S1** is rather straightforward, synthesis at >10 nmoles is a challenging endeavour. The presence of *thirteen* Ru complexes confers unusual properties to oligonucleotides, which are highlighted by the formation of a highly stable structure ($T_m$ value of 84°C). The Ru complexes affixed on the nucleobase can engage DNA in many possible ways such as intercalation, kinking, or even inducing triplex formation.<sup>72-80, 106-107</sup> In the absence of an X-ray structure, the exact nature of the stable structure formed by **S1** is currently difficult to assess. The involvement of the Ru complexes on folding of **S1** and the nature of the stable structure are even more difficult to predict since, to the best of our knowledge, no sequence equipped with thirteen complexes, covalently bound to DNA, has ever been reported yet. Fourth, as for monomeric Ru complexes, formation of aggregates could be observed during binding affinity determination of sequence **S1**. While the $K_d$ value for the bacterial target could still be evaluated, the formation of nanoaggregates could reveal deleterious for Ru-based aptamer selection methods and concentrations should be kept low. Hence, the design of future *in vitro* selections with such nucleotide analogs should consider the possible formation of such nanoaggregates. Interestingly, since we carried out parallel SELEX experiments with and without modifications, we could directly compare the outcome of both *in vitro* selections for the same target. Both the modified aptamer **S1** and unmodified aptamer **S9** displayed $K_d$ values that were in the same range, while sequence **S11** exhibited a lower propensity at binding to the bacterial target. Moreover, all these sequences display comparable binding affinities to previously reported aptamers against *S. pneumoniae* strain 262 (Klein)⁹⁷ but slightly lower than that of pan-binding aptamers isolated against mixtures of *S. pneumoniae* strains.⁹⁸ Altogether, this advocates that aptamers, either as singlets or mixtures, against *S. pneumoniae* strains holds potential as an innovative and rapidly scalable method for diagnostics and serotype isolate specific treatment.

Conclusions

Aptamers are slowly integrating the field of antibacterial agents given their high propensity at binding to targets with high affinity and specificity and due to their ease of selection. Here, we have evaluated the possibility of identifying aptamers modified with highly efficient
photosensitizers against a S. pneumoniae strain. Such chemically modified aptamers could convey specificity to the aPDT treatment modality, increase the efficiency of the PDT treatment, and ultimately lead to broad spectrum antibacterial agents. We have synthesized various Ru-modified nucleotides and evaluated their capacity at serving as substrates for polymerases for enzymatic DNA synthesis. Their compatibility with enzymatic synthesis is impaired by the presence of large aromatic ligands, which lead to efficient intercalation into dsDNA and halt DNA synthesis. We have used one of these modified nucleotides in a SELEX experiment against fixed S. pneumoniae strain R6, which resulted in the identification of a modified aptamer. This modified aptamer displayed similar binding affinities than an unmodified aptamer identified in a parallel SELEX experiment conducted with canonical DNA. During the preparation of sequence S1, we observed the formation of a secondary structure of extremely high thermal stability (Tm of 84°C), while exact Kd value determination was impaired by the formation of nanoaggregates. We envision to produce the modified sequence S1 on larger scale for structural elucidation using our recently developed ligation method¹⁰⁸, which will by-pass the need for polymerases and might facilitate the purification protocol. In addition to structural elucidation, large scale production of sequence S1 will allow further studies to verify whether this aptamer binds to live cells and whether PDT efficiency and specificity are improved by the presence of multiple, covalently bound Ru complexes. In parallel, we will also consider other, maybe less efficient PSs but better tolerated modifications³⁹ to raise aptamers for aPDT applications. Overall, we have identified the first aptamer equipped with such a complex modification pattern and these findings open directions for the identification of chemically modified aptamers to combat AMR via PDT.
**Data availability**

The authors declare that all data supporting the findings of this study are available within the article and the Supplementary Information upon reasonable request from the corresponding author.

**Competing interests**

The authors declare no competing interests.

**Acknowledgements**

M.H. acknowledges funding from Institut Pasteur, the Joe W. and Dorothy Doresett Brown Foundation (exploratory and advanced grants), the French national science foundation ANR (project "ENERGIR" grant number: ANR-21-CE11-0039 and project MolecularXiv grant number: ANR-22-PEXM-0002), and INSERM (project "FusTarG"). L. K. M. thanks the ARC Foundation for cancer research for a postdoctoral Research Fellowship (grant number: S-FB18006). M. F. and G.N. gratefully acknowledge PhD fellowships from the doctoral school MTCI, Université Paris Cité. This work was financially supported by an ERC Consolidator Grant Photo-MedMet to G.G. (GA 681679), has received support under the program "Investissements d'Avenir" launched by the French Government and implemented by the ANR with the reference ANR-10-IDEX-0001-02 PSL (G.G.)

**Supporting Information**

Supporting Information Availability: Detailed materials and methods, ESI-MS, NMR (¹H, ³¹P, ¹³C) spectra, additional gel images, data from the fluorescent binding assays, NGS sequence analysis, and fluorescent cytometry analysis are given in the Supporting Information, available free of charge at http://pubs.acs.org (PDF).

**Author information**

**Corresponding authors**

Marcel Hollenstein: Institut Pasteur, Université Paris Cité, CNRS UMR3523, Department of Structural Biology and Chemistry, Laboratory for Bioorganic Chemistry of Nucleic Acids, 28, rue du Docteur Roux, 75724 Paris Cedex 15, France.; [marcel.hollenstein@pasteur.fr](mailto:marcel.hollenstein@pasteur.fr)

Gilles Gasser:

Chimie ParisTech, PSL University, CNRS, Institute of Chemistry for Life and Health Sciences, Laboratory for Inorganic Chemical Biology, 75005 Paris, France; [gilles.gasser@chimieparistech.psl.eu](mailto:gilles.gasser@chimieparistech.psl.eu)

**Authors:**

Marie Flamme: Institut Pasteur, Université Paris Cité, CNRS UMR3523, Department of Structural Biology and Chemistry, Laboratory for Bioorganic Chemistry of Nucleic Acids, 28, rue du Docteur Roux, 75724 Paris Cedex 15, France
Germain Niogret: Institut Pasteur, Université Paris Cité, CNRS UMR3523, Department of Structural Biology and Chemistry, Laboratory for Bioorganic Chemistry of Nucleic Acids, 28, rue du Docteur Roux, 75724 Paris Cedex 15, France

Luke McKenzie: Institut Pasteur, Université Paris Cité, CNRS UMR3523, Department of Structural Biology and Chemistry, Laboratory for Bioorganic Chemistry of Nucleic Acids, 28, rue du Docteur Roux, 75724 Paris Cedex 15, France

Usman Akhtar: Institut Pasteur, Université Paris Cité, CNRS UMR3523, Department of Structural Biology and Chemistry, Laboratory for Bioorganic Chemistry of Nucleic Acids, 28, rue du Docteur Roux, 75724 Paris Cedex 15, France

Fabienne Levi-Acobas: Institut Pasteur, Université Paris Cité, CNRS UMR3523, Department of Structural Biology and Chemistry, Laboratory for Bioorganic Chemistry of Nucleic Acids, 28, rue du Docteur Roux, 75724 Paris Cedex 15, France

Albert Gandioso: Chimie ParisTech, PSL University, CNRS, Institute of Chemistry for Life and Health Sciences, Laboratory for Inorganic Chemical Biology, 75005 Paris, France

Emma Clarke: Institut Pasteur, Université Paris Cité, CNRS UMR3523, Department of Structural Biology and Chemistry, Laboratory for Bioorganic Chemistry of Nucleic Acids, 28, rue du Docteur Roux, 75724 Paris Cedex 15, France

Pierre Nicolas Bizat: Institut Pasteur, Université Paris Cité, CNRS UMR3523, Department of Structural Biology and Chemistry, Laboratory for Bioorganic Chemistry of Nucleic Acids, 28, rue du Docteur Roux, 75724 Paris Cedex 15, France

Michael Connor: Institut Pasteur, Université Paris Cité, Chromatin and Infection Laboratory, 28, rue du Docteur Roux, 75015, Paris, France

Mélanie A. Hamon: Institut Pasteur, Université Paris Cité, Chromatin and Infection Laboratory, 28, rue du Docteur Roux, 75015, Paris, France

## References

1. Vigueras, G.; Gasser, G.; Ruiz, J., Breaking the deep-red light absorption barrier of iridium(iii)-based photosensitizers. *Dalton Trans.* **2025**, 54 (4), 1320-1328.

2. Kuznetsov, K. M.; Cariou, K.; Gasser, G., Two in one: merging photoactivated chemotherapy and photodynamic therapy to fight cancer. *Chem. Sci.* **2024**, 15 (43), 17760-17780.

3. Zhang, Y.; Doan, B.-T.; Gasser, G., Metal-Based Photosensitizers as Inducers of Regulated Cell Death Mechanisms. *Chem. Rev.* **2023**, 123 (16), 10135-10155.

4. Ho, C. S.; Wong, C. T. H.; Aung, T. T.; Lakshminarayanan, R.; Mehta, J. S.; Rauz, S.; McNally, A.; Kintses, B.; Peacock, S. J.; de la Fuente-Nunez, C.; Hancock, R. E. W.; Ting, D. S. J., Antimicrobial resistance: a concise update. *Lancet Microbe* **2025**, 6 (1), 100947.

5. Jiang, J.; Lv, X.; Cheng, H.; Yang, D.; Xu, W.; Hu, Y.; Song, Y.; Zeng, G., Type I photodynamic antimicrobial therapy: Principles, progress, and future perspectives. *Acta Biomater.* **2024**, 177, 1-19.

6. Nguyen, V.-N.; Zhao, Z.; Tang, B. Z.; Yoon, J., Organic photosensitizers for antimicrobial phototherapy. *Chem. Soc. Rev.* **2022**, 51 (9), 3324-3340.

7. Gourlot, C.; Gosset, A.; Glattard, E.; Aisenbrey, C.; Rangasamy, S.; Rabineau, M.; Ouk, T.-S.; Sol, V.; Lavalle, P.; Gourlaouen, C.; Ventura, B.; Bechinger, B.; Heitz, V., Antibacterial Photodynamic Therapy in the Near-Infrared Region with a Targeting Antimicrobial Peptide Connected to a π-Extended Porphyrin. *ACS Infect. Dis.* **2022**, 8 (8), 1509-1520.

8. Huang, K.; Wang, J.; Zhuang, A.; Liu, Q.; Li, F.; Yuan, K.; Yang, Y.; Liu, Y.; Chang, H.; Liang, Y.; Sun, Y.; Yan, X.; Tang, T.; Stang, P. J.; Yang, S., Metallacage-based enhanced PDT
strategy for bacterial elimination via inhibiting endogenous NO production. *Proc. Natl. Acad. Sci. U.S.A.* **2023**, *120* (29), e2218973120.

9. Chen, H.; Li, S.; Wu, M.; Kenry; Huang, Z.; Lee, C.-S.; Liu, B., Membrane-Anchoring Photosensitizer with Aggregation-Induced Emission Characteristics for Combating Multidrug-Resistant Bacteria. *Angew. Chem. Int. Ed.* **2020**, *59* (2), 632-636.

10. Zhou, C.; Li, Y.; Shen, Y.; Lv, Z.; Feng, J.; Jiang, M.; Du, J.; Guan, W., Self-degradable photosensitizer exhibiting bacterial agglutination and membrane insertion toward safe photodynamic ablation. *Chem. Commun.* **2024**, *60* (55), 7053-7056.

11. Yu, E. Y.; Chau, J. H. C.; Lee, M. M. S.; Koo, T. H.; Lortz, R.; Lam, J. W. Y.; Kwok, R. T. K.; Li, Y.; Tang, B. Z., Recyclable and Environmentally Friendly Magnetic Nanoparticles with Aggregation-Induced Emission Photosensitizer for Sustainable Bacterial Inactivation in Water. *ACS Nano* **2024**, *18* (3), 1907-1920.

12. Liu, S.; Wang, B.; Yu, Y.; Liu, Y.; Zhuang, Z.; Zhao, Z.; Feng, G.; Qin, A.; Tang, B. Z., Cationization-Enhanced Type I and Type II ROS Generation for Photodynamic Treatment of Drug-Resistant Bacteria. *ACS Nano* **2022**, *16* (6), 9130-9141.

13. Demidova, T. N.; Hamblin, M. R., Effect of Cell-Photosensitizer Binding and Cell Density on Microbial Photoinactivation. *Antimicrob. Agents Chemother.* **2005**, *49* (6), 2329-2335.

14. Zhou, W.; Chen, L.; Li, H.; Wu, M.; Liang, M.; Liu, Q.; Wu, W.; Jiang, X.; Zhen, X., Membrane Disruption-Enhanced Photodynamic Therapy against Gram-Negative Bacteria by a Peptide-Photosensitizer Conjugate. *ACS Nano* **2024**, *18* (30), 19771-19782.

15. Galstyan, A.; Schiller, R.; Dobrindt, U., Boronic Acid Functionalized Photosensitizers: A Strategy To Target the Surface of Bacteria and Implement Active Agents in Polymer Coatings. *Angew. Chem. Int. Ed.* **2017**, *56* (35), 10362-10366.

16. Soncin, M.; Fabris, C.; Busetti, A.; Dei, D.; Nistri, D.; Roncucci, G.; Jori, G., Approaches to selectivity in the Zn(ii)-phthalocyanine-photosensitized inactivation of wild-type and antibiotic-resistant Staphylococcus aureus. *Photochem. Photobiol. Sci.* **2002**, *1* (10), 815-819.

17. Grimmeisen, M.; Jessen-Trefzer, C., Increasing the Selectivity of Light-Active Antimicrobial Agents – Or How To Get a Photosensitizer to the Desired Target. *ChemBioChem* **2023**, *24* (18), e202300177.

18. Nestoros, E.; Sharma, A.; Kim, E.; Kim, J. S.; Vendrell, M., Smart molecular designs and applications of activatable organic photosensitizers. *Nat. Rev. Chem.* **2025**, *9* (1), 46-60.

19. Yu, C.; Gao, Y.; Zhang, Y.; Wang, J.; Zhang, Y.; Li, J.; Zhang, X.; Wu, Z.; Zhang, X., A Targeted Photosensitizer Mediated by Visible Light for Efficient Therapy of Bacterial Keratitis. *Biomacromolecules* **2021**, *22* (9), 3704-3717.

20. Ucuncu, M.; Mills, B.; Duncan, S.; Staderini, M.; Dhaliwal, K.; Bradley, M., Polymyxin-based photosensitizer for the potent and selective killing of Gram-negative bacteria. *Chem. Commun.* **2020**, *56* (26), 3757-3760.

21. Dunn, M. R.; Jimenez, R. M.; Chaput, J. C., Analysis of aptamer discovery and technology. *Nat. Rev. Chem.* **2017**, *1* (10), 0076.

22. Dong, Y. H.; Wang, J. P.; Chen, L.; Chen, H. N.; Dang, S. B.; Li, F., Aptamer-based assembly systems for SARS-CoV-2 detection and therapeutics. *Chem. Soc. Rev.* **2024**, *53*, 6830-6859.

23. Yang, L. F.; Ling, M.; Kacherovsky, N.; Pun, S. H., Aptamers 101: aptamer discovery and in vitro applications in biosensors and separations. *Chem. Sci.* **2023**, *14* (19), 4961-4978.

24. Tuerk, C.; Gold, L., Systematic Evolution of Ligands by Exponential Enrichment: RNA Ligands to Bacteriophage T4 DNA Polymerase. *Science* **1990**, *249*, 505-510.

25. Ellington, A. D.; Szostak, J. W., In vitro selection of RNA molecules that bind specific ligands. *Nature* **1990**, *346*, 818-822.

26. Xiao, Y.; Pan, T.; Da, W.; Liu, Y.; Chen, S.; Chen, D.; Liu, K.; Zheng, Y.; Xie, D.; Gao, Y.; Xu, H.; Sun, Y.; Tan, W., Aptamer-drug conjugates-loaded bacteria for pancreatic cancer synergistic therapy. *Sig. Transduct. Target. Ther.* **2024**, *9* (1), 272.

27. Powell Gray, B.; Kelly, L.; Ahrens, D. P.; Barry, A. P.; Kratschmer, C.; Levy, M.; Sullenger, B. A., Tunable cytotoxic aptamer-drug conjugates for the treatment of prostate cancer. *Proc. Natl. Acad. Sci. U.S.A.* **2018**, *115* (18), 4761-4766.

28. Yoon, S.; Huang, K.-W.; Reebye, V.; Spalding, D.; Przytycka, T. M.; Wang, Y.; Swiderski, P.; Li, L.; Armstrong, B.; Reccia, I.; Zacharoulis, D.; Dimas, K.; Kusano, T.; Shively, J.; Habib, N.; Rossi, J. J., Aptamer-Drug Conjugates of Active Metabolites of Nucleoside
Analogs and Cytotoxic Agents Inhibit Pancreatic Tumor Cell Growth. Mol. Ther. Nucleic Acids **2017**, *6*, 80-88.

29. Kelly, L.; Maier, K. E.; Yan, A.; Levy, M., A comparative analysis of cell surface targeting aptamers. *Nat. Commun.* **2021**, *12* (1), 6275.

30. Wang, R.; Zhu, G.; Mei, L.; Xie, Y.; Ma, H.; Ye, M.; Qing, F.-L.; Tan, W., Automated Modular Synthesis of Aptamer-Drug Conjugates for Targeted Drug Delivery. *J. Am. Chem. Soc.* **2014**, *136* (7), 2731-2734.

31. McKenzie, L. K.; Flamme, M.; Felder, P. S.; Karges, J.; Bonhomme, F.; Gandioso, A.; Malosse, C.; Gasser, G.; Hollenstein, M., A ruthenium-oligonucleotide bioconjugated photosensitizing aptamer for cancer cell specific photodynamic therapy. *RSC Chem. Biol.* **2022**, *3* (1), 85-95.

32. Hou, W.; Liu, Y.; Jiang, Y.; Wu, Y.; Cui, C.; Wang, Y.; Zhang, L.; Teng, I. T.; Tan, W., Aptamer-based multifunctional ligand-modified UCNPs for targeted PDT and bioimaging. *Nanoscale* **2018**, *10* (23), 10986-10990.

33. Zhang, P.; Fischer, A.; Ouyang, Y.; Wang, J.; Sohn, Y. S.; Nechushtai, R.; Pikarsky, E.; Fan, C.; Willner, I., Aptamer-modified DNA tetrahedra-gated metal-organic framework nanoparticle carriers for enhanced chemotherapy or photodynamic therapy. *Chem. Sci.* **2021**, *12* (43), 14473-14483.

34. Yan, J.; Gao, T.; Lu, Z.; Yin, J.; Zhang, Y.; Pei, R., Aptamer-Targeted Photodynamic Platforms for Tumor Therapy. *ACS Appl. Mater. Interfaces* **2021**, *13* (24), 27749-27773.

35. Kim, J.; Park, W.; Kim, D.; Lee, E. S.; Lee, D. H.; Jeong, S.; Park, J. M.; Na, K., Tumor-Specific Aptamer-Conjugated Polymeric Photosensitizer for Effective Endo-Laparoscopic Photodynamic Therapy. *Adv. Funct. Mater.* **2019**, *29* (23), 1900084.

36. Zhang, D.; Zheng, A.; Li, J.; Wu, M.; Cai, Z.; Wu, L.; Wei, Z.; Yang, H.; Liu, X.; Liu, J., Tumor Microenvironment Activable Self-Assembled DNA Hybrids for pH and Redox Dual-Responsive Chemotherapy/PDT Treatment of Hepatocellular Carcinoma. *Adv. Sci.* **2017**, *4* (4), 1600460.

37. Shieh, Y.-A.; Yang, S.-J.; Wei, M.-F.; Shieh, M.-J., Aptamer-Based Tumor-Targeted Drug Delivery for Photodynamic Therapy. *ACS Nano* **2010**, *4* (3), 1433-1442.

38. Shen, Y.; Tian, Q.; Sun, Y.; Xu, J.-J.; Ye, D.; Chen, H.-Y., ATP-Activatable Photosensitizer Enables Dual Fluorescence Imaging and Targeted Photodynamic Therapy of Tumor. *Anal. Chem.* **2017**, *89* (24), 13610-13617.

39. Niogret, G.; Chériaux, C.; Bonhomme, F.; Levi-Acobas, F.; Figliola, C.; Ulrich, G.; Gasser, G.; Hollenstein, M., A toolbox for enzymatic modification of nucleic acids with photosensitizers for photodynamic therapy. *RSC Chem. Biol.* **2024**, *5* (9), 841-852.

40. Dahm, G. C.; Akhtar, U.; Bouvier-Müller, A.; Lim, L.; Levi-Acobas, F.; Bizat, P. N.; Niogret, G.; Tanner, J.; Ducongé, F.; Hollenstein, M., Probing three-dimensional cyclooctatetraene as nucleobase modification in aptamer selection. *ChemRxiv* **2025**, doi:10.26434/chemrxiv-2025-cglxw.

41. Ren, X.; Gelinas, A. D.; von Carlowitz, I.; Janjic, N.; Pyle, A. M., Structural basis for IL-1α recognition by a modified DNA aptamer that specifically inhibits IL-1α signaling. *Nat. Commun.* **2017**, *8* (1), 810.

42. Ren, X.; Gelinas, A. D.; Linehan, M.; Iwasaki, A.; Wang, W.; Janjic, N.; Pyle, A. M., Evolving A RIG-I Antagonist: A Modified DNA Aptamer Mimics Viral RNA. *J. Mol. Biol.* **2021**, *433* (21), 167227.

43. Gawande, B. N.; Rohloff, J. C.; Carter, J. D.; von Carlowitz, I.; Zhang, C.; Schneider, D. J.; Janjic, N., Selection of DNA aptamers with two modified bases. *Proc. Natl. Acad. Sci. U.S.A.* **2017**, *114* (11), 2898-2903.

44. Kohn, E. M.; Konovalov, K.; Gomez, C. A.; Hoover, G. N.; Yik, A. K.-h.; Huang, X.; Martell, J. D., Terminal Alkyne-Modified DNA Aptamers with Enhanced Protein Binding Affinities. *ACS Chem. Biol.* **2023**, *18* (9), 1976-1984.

45. Kong, D.; Yeung, W.; Hili, R., In Vitro Selection of Diversely Functionalized Aptamers. *J. Am. Chem. Soc.* **2017**, *139* (40), 13977-13980.

46. Morihiro, K.; Hasegawa, O.; Kasahara, Y.; Mori, S.; Kasai, T.; Kuwahara, M.; Obika, S., Azobenzene-modified DNA aptamers evolved by capillary electrophoresis (CE)-SELEX method. *Bioorg. Med. Chem. Lett.* **2021**, *31*, 127607.

47. Monro, S.; Colón, K. L.; Yin, H.; Roque, J., III; Konda, P.; Gujar, S.; Thummel, R. P.; Lilge, L.; Cameron, C. G.; McFarland, S. A., Transition Metal Complexes and Photodynamic
Therapy from a Tumor-Centered Approach: Challenges, Opportunities, and Highlights from the Development of TLD1433. *Chem. Rev.* **2019**, *119*(2), 797-828.

48. McFarland, S. A.; Mandel, A.; Dumoulin-White, R.; Gasser, G., Metal-based photosensitizers for photodynamic therapy: the future of multimodal oncology? *Curr. Opin. Chem. Biol.* **2020**, *56*, 23-27.

49. Hoskins, J.; Alborn, W. E.; Arnold, J.; Blaszczak, L. C.; Burgett, S.; DeHoff, B. S.; Estrem, S. T.; Fritz, L.; Fu, D.-J.; Fuller, W.; Geringer, C.; Gilmour, R.; Glass, J. S.; Khoja, H.; Kraft, A. R.; Lagace, R. E.; LeBlanc, D. J.; Lee, L. N.; Lefkowitz, E. J.; Lu, J.; Matsushima, P.; McAhren, S. M.; McHenney, M.; McLeaster, K.; Mundy, C. W.; Nicas, T. I.; Norris, F. H.; O'Gara, M.; Peery, R. B.; Robertson, G. T.; Rockey, P.; Sun, P.-M.; Winkler, M. E.; Yang, Y.; Young-Bellido, M.; Zhao, G.; Zook, C. A.; Baltz, R. H.; Jaskunas, S. R.; Rosteck, P. R.; Skatrud, P. L.; Glass, J. I., Genome of the Bacterium *Streptococcus pneumoniae* Strain R6. *J. Bacteriol.* **2001**, *183*(19), 5709-5717.

50. Connor, M. G.; Camarasa, T. M. N.; Patey, E.; Rasid, O.; Barrio, L.; Weight, C. M.; Miller, D. P.; Heyderman, R. S.; Lamont, R. J.; Enninga, J.; Hamon, M. A., The histone demethylase KDM6B fine-tunes the host response to Streptococcus pneumoniae. *Nat. Microbiol.* **2021**, *6*(2), 257-269.

51. Hollenstein, M.; Hipolito, C. J.; Lam, C. H.; Perrin, D. M., A self-cleaving DNA enzyme modified with amines, guanidines and imidazoles operates independently of divalent metal cations (M2). *Nucleic Acids Res.* **2009**, *37*(5), 1638-1649.

52. Levi-Acobas, F.; Katolik, A.; Röthlisberger, P.; Cokelaer, T.; Sarac, I.; Damha, M. J.; Leumann, C. J.; Hollenstein, M., Compatibility of 5-ethynyl-2'F-ANA UTP with in vitro selection for the generation of base-modified, nuclease resistant aptamers. *Org. Biomol. Chem.* **2019**, *17*(35), 8083-8087.

53. Abad-Montero, D.; Gandioso, A.; Izquierdo-García, E.; Chumillas, S.; Rovira, A.; Bosch, M.; Jordà-Redondo, M.; Castaño, D.; Bonelli, J.; Novikov, V. V.; Deyà, A.; Hernández, J. L.; Galino, J.; Alberto, M. E.; Francés-Monerris, A.; Nonell, S.; Gasser, G.; Marchán, V., Ruthenium(II) Polypyridyl Complexes Containing COUBPY Ligands as Potent Photosensitizers for the Efficient Phototherapy of Hypoxic Tumors. *J. Am. Chem. Soc.* **2025**, *147*(9), 7360-7376.

54. Knoll, J. D.; Turro, C., Control and utilization of ruthenium and rhodium metal complex excited states for photoactivated cancer therapy. *Coord. Chem. Rev.* **2015**, *282-283*, 110-126.

55. Karges, J., Clinical Development of Metal Complexes as Photosensitizers for Photodynamic Therapy of Cancer. *Angew. Chem. Int. Ed.* **2022**, *61*(5), e202112236.

56. Heinemann, F.; Karges, J.; Gasser, G., Critical Overview of the Use of Ru(II) Polypyridyl Complexes as Photosensitizers in One-Photon and Two-Photon Photodynamic Therapy. *Acc. Chem. Res.* **2017**, *50*(11), 2727-2736.

57. Mari, C.; Pierroz, V.; Ferrari, S.; Gasser, G., Combination of Ru(ii) complexes and light: new frontiers in cancer therapy. *Chem. Sci.* **2015**, *6*(5), 2660-2686.

58. Friedman, A. E.; Chambron, J. C.; Sauvage, J. P.; Turro, N. J.; Barton, J. K., A molecular light switch for DNA: Ru(bpy)2(dppz)2+. *J. Am. Chem. Soc.* **1990**, *112*(12), 4960-4962.

59. Hess, J.; Huang, H.; Kaiser, A.; Pierroz, V.; Blacque, O.; Chao, H.; Gasser, G., Evaluation of the Medicinal Potential of Two Ruthenium(II) Polypyridine Complexes as One- and Two-Photon Photodynamic Therapy Photosensitizers. *Chem. Eur. J.* **2017**, *23*(41), 9888-9896.

60. Karges, J.; Heinemann, F.; Jakubaszek, M.; Maschietto, F.; Subecz, C.; Dotou, M.; Vinck, R.; Blacque, O.; Tharaud, M.; Goud, B.; Viñuelas Zahínos, E.; Spingler, B.; Ciofini, I.; Gasser, G., Rationally Designed Long-Wavelength Absorbing Ru(II) Polypyridyl Complexes as Photosensitizers for Photodynamic Therapy. *J. Am. Chem. Soc.* **2020**, *142*(14), 6578-6587.

61. Cheung, Y. W.; Röthlisberger, P.; Mechaly, A. E.; Weber, P.; Levi-Acobas, F.; Lo, Y.; Wong, A. W. C.; Kinghorn, A. B.; Haouz, A.; Savage, G. P.; Hollenstein, M.; Tanner, J. A., Evolution of abiotic cubane chemistries in a nucleic acid aptamer allows selective recognition of a malaria biomarker. *Proc. Natl. Acad. Sci. U.S.A.* **2020**, *117*(29), 16790-16798.

62. Niogret, G.; Bouvier-Müller, A.; Figazzolo, C.; Joyce, J. M.; Bonhomme, F.; England, P.; Mayboroda, O.; Pellarin, R.; Gasser, G.; Tucker, J. H. R.; Tanner, J. A.; Savage, G. P.; Hollenstein, M., Interrogating Aptamer Chemical Space Through Modified Nucleotide Substitution Facilitated by Enzymatic DNA Synthesis. *ChemBioChem* **2024**, *25*(1), e202300539.
63. Baccaro, A.; Marx, A., Enzymatic Synthesis of Organic-Polymer-Grafted DNA. *Chem. Eur. J.* **2010**, *16* (1), 218-226.

64. Niogret, G.; Roethlisberger, P.; Levi-Acobas, F.; Bonhomme, F.; Gasser, G.; Hollenstein, M., Facilitated Synthetic Access to Boronic Acid-Modified Nucleoside Triphosphates and Compatibility with Enzymatic DNA Synthesis. *Synlett* **2024**, *35* (06), 677-683.

65. McCafferty, D. G.; Bishop, B. M.; Wall, C. G.; Hughes, S. G.; Mecklenberg, S. L.; Meyer, T. J.; Erickson, B. W., Synthesis of redox derivatives of lysine and their use in solid-phase synthesis of a light-harvesting peptide. *Tetrahedron* **1995**, *51* (4), 1093-1106.

66. Audi, H.; Azar, D. F.; Mahjoub, F.; Farhat, S.; El Masri, Z.; El-Sibai, M.; Abi-Habib, R. J.; Khnayzer, R. S., Cytotoxicity modulation of ruthenium(II) tris-bathophenanthroline complexes with systematically varied charge. *J. Photochem. Photobiol. A Chem.* **2018**, *351*, 59-68.

67. Weizman, H.; Tor, Y., Redox-Active Metal-Containing Nucleotides: Synthesis, Tunability, and Enzymatic Incorporation into DNA. *J. Am. Chem. Soc.* **2002**, *124* (8), 1568-1569.

68. Vrábel, M.; Horáková, P.; Pivoňková, H.; Kalachova, L.; Černocká, H.; Cahová, H.; Pohl, R.; Šebest, P.; Havran, L.; Hocek, M.; Fojta, M., Base-Modified DNA Labeled by [Ru(bpy)3]2+ and [Os(bpy)3]2+ Complexes: Construction by Polymerase Incorporation of Modified Nucleoside Triphosphates, Electrochemical and Luminescent Properties, and Applications. *Chem. Eur. J.* **2009**, *15* (5), 1144-1154.

69. Flamme, M.; Clarke, E.; Gasser, G.; Hollenstein, M., Applications of Ruthenium Complexes Covalently Linked to Nucleic Acid Derivatives. *Molecules* **2018**, *23* (7), 1515.

70. Puckett, C. A.; Barton, J. K., Methods to Explore Cellular Uptake of Ruthenium Complexes. *J. Am. Chem. Soc.* **2007**, *129* (1), 46-47.

71. Soliman, N.; McKenzie, L. K.; Karges, J.; Bertrand, E.; Tharaud, M.; Jakubaszek, M.; Guérineau, V.; Goud, B.; Hollenstein, M.; Gasser, G.; Thomas, C. M., Ruthenium-initiated polymerization of lactide: a route to remarkable cellular uptake for photodynamic therapy of cancer. *Chem. Sci.* **2020**, *11* (10), 2657-2663.

72. Hibino, M.; Aiba, Y.; Watanabe, Y.; Shoji, O., Peptide Nucleic Acid Conjugated with Ruthenium-Complex Stabilizing Double-Duplex Invasion Complex Even under Physiological Conditions. *ChemBioChem* **2018**, *19* (15), 1601-1604.

73. Prieto Otoya, T. D.; McQuaid, K. T.; Paterson, N. G.; Cardin, D. J.; Kellett, A.; Cardin, C. J., Re-pairing DNA: binding of a ruthenium phi complex to a double mismatch. *Chem. Sci.* **2024**, *15* (24), 9096-9103.

74. Prieto Otoya, T. D.; McQuaid, K. T.; Hennessy, J.; Menounou, G.; Gibney, A.; Paterson, N. G.; Cardin, D. J.; Kellett, A.; Cardin, C. J., Probing a Major DNA Weakness: Resolving the Groove and Sequence Selectivity of the Diimine Complex $\Lambda$-[Ru(phen)2phi]2+. *Angew. Chem. Int. Ed.* **2024**, *63* (13), e202318863.

75. Gibney, A.; Kellett, A., Gene Editing with Artificial DNA Scissors. *Chem. Eur. J.* **2024**, *30* (57), e202401621.

76. Prieto Otoya, T. D.; McQuaid, K. T.; Cardin, C. J., Structural insights into G-quadruplex binding by metal complexes: implications for drug design. *Med. Chem. Res.* **2024**, *33* (11), 2001-2019.

77. Hall, J. P.; O'Sullivan, K.; Naseer, A.; Smith, J. A.; Kelly, J. M.; Cardin, C. J., Structure determination of an intercalating ruthenium dipyridophenazine complex which kinks DNA by semiintercalation of a tetraazaphenanthrene ligand. *Proc. Natl. Acad. Sci. U.S.A.* **2011**, *108* (43), 17610-17614.

78. Dupureur, C. M.; Barton, J. K., Use of Selective Deuteration and 1H NMR in Demonstrating Major Groove Binding of. DELTA.-[Ru (phen) 2 dppz] 2+ to d (GTCGAC) 2. *J. Am. Chem. Soc.* **1994**, *116* (22), 10286-10287.

79. Hall, J. P.; Gurung, S. P.; Henle, J.; Poidl, P.; Andersson, J.; Lincoln, P.; Winter, G.; Sorensen, T.; Cardin, D. J.; Brazier, J. A.; Cardin, C. J., Guanine Can Direct Binding Specificity of Ru-dipyridophenazine (dppz) Complexes to DNA through Steric Effects. *Chem. Eur. J.* **2017**, *23* (21), 4981-4985.

80. Niyazi, H.; Hall, J. P.; O'Sullivan, K.; Winter, G.; Sorensen, T.; Kelly, J. M.; Cardin, C. J., Crystal structures of $\Lambda$-[Ru(phen)₂dppz]²⁺ with oligonucleotides containing TA/TA and AT/AT steps show two intercalation modes. *Nat. Chem.* **2012**, *4* (8), 621-8.
81. Kumar, P.; Dasari, S.; Patra, A. K., Ruthenium(II) complexes of saccharin with dipyridoquinoxaline and dipyridophenazine: Structures, biological interactions and photoinduced DNA damage activity. *Eur. J. Med. Chem.* **2017**, *136*, 52-62.

82. Tan, C.; Liu, J.; Li, H.; Zheng, W.; Shi, S.; Chen, L.; Ji, L., Differences in structure, physiological stability, electrochemistry, cytotoxicity, DNA and protein binding properties between two Ru(III) complexes. *J. Inorg. Biochem.* **2008**, *102* (2), 347-358.

83. Wise, D. E.; Gamble, A. J.; Arkawazi, S. W.; Walton, P. H.; Galan, M. C.; O'Hagan, M. P.; Hogg, K. G.; Marrison, J. L.; O'Toole, P. J.; Sparkes, H. A.; Lynam, J. M.; Pringle, P. G., Cytotoxic (cis,cis-1,3,5-triaminocyclohexane)ruthenium(ii)-diphosphine complexes; evidence for covalent binding and intercalation with DNA. *Dalton Trans.* **2020**, *49* (43), 15219-15230.

84. Liu, X.-W.; Chen, Z.-g.; Li, L.; Chen, Y.-D.; Lu, J.-L.; Zhang, D.-S., DNA-binding, photocleavage studies of ruthenium(II) complexes with 2-(2-quinolinyl) imidazo[4,5-f][1,10]phenanthroline. *Spectrochim. Acta A Mol. Biomol. Spectrosc.* **2013**, *102*, 142-149.

85. Waring, M. J., Complex formation between ethidium bromide and nucleic acids. *J. Mol. Biol.* **1965**, *13* (1), 269-282.

86. Boger, D. L.; Fink, B. E.; Brunette, S. R.; Tse, W. C.; Hedrick, M. P., A Simple, High-Resolution Method for Establishing DNA Binding Affinity and Sequence Selectivity. *J. Am. Chem. Soc.* **2001**, *123* (25), 5878-5891.

87. Jenkins, Y.; Friedman, A. E.; Turro, N. J.; Barton, J. K., Characterization of dipyridophenazine complexes of ruthenium(II): the light switch effect as a function of nucleic acid sequence and conformation. *Biochemistry* **1992**, *31* (44), 10809-16.

88. Xu, H.; Liang, Y.; Zhang, P.; Du, F.; Zhou, B.-R.; Wu, J.; Liu, J.-H.; Liu, Z.-G.; Ji, L.-N., Biophysical studies of a ruthenium(II) polypyridyl complex binding to DNA and RNA prove that nucleic acid structure has significant effects on binding behaviors. *J. Biol. Inorg. Chem.* **2005**, *10* (5), 529-538.

89. Srishailam, A.; Kumar, Y. P.; Gabra, N. M. D.; Reddy, P. V.; Deepika, N.; Veerababu, N.; Satyanarayana, S., Synthesis, DNA-binding, Cytotoxicity, Photo Cleavage, Antimicrobial and Docking Studies of Ru(II) Polypyridyl Complexes. *J. Fluoresc.* **2013**, *23* (5), 897-908.

90. Narciso, A. R.; Dookie, R.; Nannapaneni, P.; Normark, S.; Henriques-Normark, B., Streptococcus pneumoniae epidemiology, pathogenesis and control. *Nat. Rev. Microbiol.* **2025**, *23* (4), 256-271.

91. Huddar, S.; Park, C. M.; Kim, H. J.; Jang, S.; Lee, S., Discovery of 4-hydroxy-2-oxo-1,2-dihydroquinolines as potential inhibitors of *Streptococcus pneumoniae*, including drug-resistant strains. *Bioorg. Med. Chem. Lett.* **2020**, *30* (9), 127071.

92. Chaudhuri, R.; Prasanth, T.; Biswas, D.; Mandal, S.; Dash, J., Combating multidrug-resistance in S. pneumoniae: a G-quadruplex binding inhibitor of efflux pump and its bio-orthogonal assembly. *NAR Mol. Med.* **2024**, *1* (2), ugae005.

93. Watkins, E. R.; Kalizang'Oma, A.; Gori, A.; Gupta, S.; Heyderman, R. S., Factors affecting antimicrobial resistance in *Streptococcus pneumoniae* following vaccination introduction. *Trends Microbiol.* **2022**, *30* (12), 1135-1145.

94. Dong, W.; Rasid, O.; Chevalier, C.; Connor, M.; Eldridge, M. J. G.; Hamon, M. A., Streptococcus pneumoniae Infection Promotes Histone H3 Dephosphorylation by Modulating Host PP1 Phosphatase. *Cell Rep.* **2020**, *30* (12), 4016-4026.e4.

95. Asaithamby, A.; Hu, B.; Delgado, O.; Ding, L. H.; Story, M. D.; Minna, J. D.; Shay, J. W.; Chen, D. J., Irreparable complex DNA double-strand breaks induce chromosome breakage in organotypic three-dimensional human lung epithelial cell culture. *Nucleic Acids Res.* **2011**, *39* (13), 5474-88.

96. Renders, M.; Miller, E.; Lam, C. H.; Perrin, D. M., Whole cell-SELEX of aptamers with a tyrosine-like side chain against live bacteria. *Org. Biomol. Chem.* **2017**, *15* (9), 1980-1989.

97. Bayraç, A. T.; Donmez, S. I., Selection of DNA aptamers to Streptococcus pneumonia and fabrication of graphene oxide based fluorescent assay. *Anal. Biochem.* **2018**, *556*, 91-98.

98. Kavruk, M.; Babaie, Z.; Kibar, G.; Çetin, B.; Yeşilkaya, H.; Amrani, Y.; Dursun, A. D.; Özalp, V. C., Aptamer decorated PDA@magnetic silica microparticles for bacteria purification. *Microchim. Acta* **2024**, *191* (5), 285.

99. Oliver, M. B.; van der Linden, M. P. G.; Küntzel, S. A.; Saad, J. S.; Nahm, M. H., Discovery of Streptococcus pneumoniae serotype 6 variants with glycosyltransferases synthesizing two differing repeating units. *J. Biol. Chem.* **2013**, *288* (36), 25976-25985.

100. Tettelin, H.; Nelson, K. E.; Paulsen, I. T.; Eisen, J. A.; Read, T. D.; Peterson, S.; Heidelberg, J.; DeBoy, R. T.; Haft, D. H.; Dodson, R. J.; Durkin, A. S.; Gwinn, M.; Kolonay, J.
F.; Nelson, W. C.; Peterson, J. D.; Umayam, L. A.; White, O.; Salzberg, S. L.; Lewis, M. R.; Radune, D.; Holtzapple, E.; Khouri, H.; Wolf, A. M.; Utterback, T. R.; Hansen, C. L.; McDonald, L. A.; Feldblyum, T. V.; Angiuoli, S.; Dickinson, T.; Hickey, E. K.; Holt, I. E.; Loftus, B. J.; Yang, F.; Smith, H. O.; Venter, J. C.; Dougherty, B. A.; Morrison, D. A.; Hollingshead, S. K.; Fraser, C. M., Complete Genome Sequence of a Virulent Isolate of *Streptococcus pneumoniae*. *Science* **2001**, *293* (5529), 498-506.

101. Jothi, R.; Manikandakumar, K.; Ganesan, K.; Parthasarathy, S., On the analysis of the virulence nature of TIGR4 and R6 strains of *Streptococcus pneumoniae* using genome comparison tools. *J. Chem. Sci.* **2007**, *119* (5), 559-563.

102. Notaro, A.; Gasser, G.; Castonguay, A., Note of Caution for the Aqueous Behaviour of Metal-Based Drug Candidates. *ChemMedChem* **2020**, *15* (4), 345-348.

103. Vinck, R.; Dömötör, O.; Karges, J.; Jakubaszek, M.; Seguin, J.; Tharaud, M.; Guérineau, V.; Cariou, K.; Mignet, N.; Enyedy, É. A.; Gasser, G., In Situ Bioconjugation of a Maleimide-Functionalized Ruthenium-Based Photosensitizer to Albumin for Photodynamic Therapy. *Inorg. Chem.* **2023**, *62* (38), 15510-15526.

104. Gutiérrez-Santana, J. C.; Coria-Jiménez, V. R., Diagnosis and Therapeutic Strategies Based on Nucleic Acid Aptamers Selected against Pseudomonas aeruginosa: The Challenge of Cystic Fibrosis. *ChemMedChem* **2024**, *19* (2), e202300544.

105. Shum, K. T.; Lui, E. L. H.; Wong, S. C. K.; Yeung, P.; Sam, L.; Wang, Y.; Watt, R. M.; Tanner, J. A., Aptamer-Mediated Inhibition of Mycobacterium tuberculosis Polyphosphate Kinase 2. *Biochemistry* **2011**, *50* (15), 3261-3271.

106. Grimm, G. N.; Boutorine, A. S.; Lincoln, P.; Nordén, B.; Hélène, C., Formation of DNA Triple Helices by an Oligonucleotide Conjugated to a Fluorescent Ruthenium Complex. *ChemBioChem* **2002**, *3* (4), 324-331.

107. Dalla Pozza, M.; Abdullrahman, A.; Cardin, C. J.; Gasser, G.; Hall, J. P., Three's a crowd – stabilisation, structure, and applications of DNA triplexes. *Chem. Sci.* **2022**, *13* (35), 10193-10215.

108. Sabat, N.; Stämpfli, A.; Hanlon, S.; Bisagni, S.; Sladojevich, F.; Püntener, K.; Hollenstein, M., Template-dependent DNA ligation for the synthesis of modified oligonucleotides. *Nat. Commun.* **2024**, *15* (1), 8009.
Graphical abstract

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