The obtained signal amplification revealed well-defined fluorescent foci located in the perinuclear space and RNase treatment revealed the preferential binding to G4-RNA. Furthermore, ligand treatment affected significantly BG4 foci formation in cells. Our work headed to the development of a new imaging approach combining the advantages of immunostaining and G4-recognition by G4 ligands leading to visualization of G4/ligands species in cells with unrivaled precision and sensitivity.G-quadruplexes (G4s) are secondary structures forming in G-rich nucleic acids. G4s are assumed to play critical roles in biology, nonetheless their detection in cells is still challenging. For tracking G4s, synthetic molecules (G4 ligands) can be used as reporters and have found wide application for this purpose through chemical functionalization with a fluorescent tag. However, this approach is limited by a low-labeling degree impeding precise visualization in specific subcellular regions. Herein, we present a new visualization strategy based on the immuno-recognition of 5-bromo-2′-deoxyuridine (5-BrdU) modified G4 ligands, functionalized prior- or post-G4-target binding by CuAAC. Remarkably, recognition of the tag by antibodies leads to the detection of the modified ligands exclusively when bound to a G4 target both   In this regard, bioinformatics studies predict the existence of a large number of G4 structures within both human genome and transcriptome. Genome-wide, G4s were found in promoters, coding regions, introns and untranslated (UTRs) regions of genes and intergenic regions  (G-quadruplexes (G4s) are four-stranded nucleic acid secondary structures adopted by repetitive G-rich DNA and RNA sequences, which fold over upon coordination with monovalent cations ,2. These regions . Importa regions . More re regions . High-th(G4-Seq) . In addi(G4-Seq) ,20.in situ functionalization  salts limits the application of CuAAC in fixed cells, pioneering studies reported its use to functionalize G4 ligands with fluorescent tags allowing identification of G4 targets in both DNA and RNA  is insufficient to detect single complexes formed by tagged-G4 ligands bound to G4 structures unless the latter are present in multiple copies  that capitalizes both on specific recognition by small molecule and antibody signal amplification properties. This unprecedented integrated approach is expected to be more efficient in systematically identifying G4 ligand targets in cells. By using the Pyrido Dicarboxamide (PDC) core as selective G4 ligand, we synthesized four hapten modified PDC conjugates and four PDC CuAAC precursors for ructures . As hapts Figure , right on Figure , left, iThe experiments were carried out on pre-folded G-quadruplex structures: the sequences were heated at 95°C for 10 min in opportune buffer and left to fold in ice for 30 min. Duplex DNA and transfer RNA (tRNA) sequences were heated at 95°C for 10 min and left to cool down to room temperature over a period of 3 h. Buffer conditions are specified below for each G4 sequence.dsDNA competitor at 0, 3 and 10 μM  respectively, or RNA competitor at 0 and 3 μM, in a total volume of 25 μl. Solutions were prepared in buffer containing 10 mM Li.Caco pH 7.2, and 10 mM KCl, completed by 90 mM LiCl for F21T, F21CTAT and 1 mM KCl, completed by 99 mM LiCl for all the others G-quadruplex sequences. The FAM channel was used to collect the fluorescence signal. Stabilization of compounds with G4-structures was monitored as followed: 5 min at 25°C and then increase of 0.5°C every minute until reaching 95°C. The stabilization was evaluated by determining the temperature at half denaturation of the G4  in the absence and in the presence of ligand, duplex DNA, and tRNA competitor, by using Origin 2021.As previously reported, stabilization of ligands on G4-structures was monitored via FRET-melting assay performed in 96-well plates on a real-time PCR apparatus, 7900HT Fast Real-Time PCR system. This assay was carried out with oligonucleotides mimicking biological relevant G4-forming sequences, labeled with FRET partners at each end (5′-FAM (donor) and 3′-TAMRA (acceptor)) in the presence of G4 ligands. To determine ligand G4 versus duplex selectivity, an unlabeled DNA competitor ds26 was used. In order to assess RNA-G4 versus RNA single strand selectivity, an unlabeled tRNA sequence was used. This oligonucleotide consists of an average of 85 nucleobases and was used at ≃3 μM . Each well was filled with: 0.20 μM (1 equiv) labeled G4 DNA or RNA, 1 μM (5 equiv) of ligand, and A series of oligonucleotides covering a range of possible G4 conformations was used.The experiments were carried out on pre-folded G-quadruplex structures: the sequences were heated at 95°C for 10 min (10 mM Li.Caco pH = 7.3 and 100 mM KCl) and left to fold in ice for 30 min. Duplex DNA was heated at 95°C for 10 min  and left to cool down to room temperature over a period of 3 h.exc = 495 nm, 1.0 nm increment, 0.1 s integration time and slit 5 nm). The percentage of TO displacement was calculated as follows: TOD (%) = 100 – (F / F0) × 100, where F0 is the maximum fluorescence emission area of bound TO before addition of ligand, and F is the fluorescence emission area after each ligand addition. The percentage of TO displacement was then plotted as a function of the concentration of added ligand.FID-assay was performed on a Cary Eclipse spectrofluorometer, using 1 ml quartz cell as previously reported. About 0.25 μM of folded DNA prepared in a solution of 10 mM Li.Caco pH = 7.3 and 100 mM KCl was mixed with 0.50 μM Thiazole Orange (TO) for G4-DNA and 0.75 μM for duplex DNA to obtain a total volume of 1 ml. The solution was left to stand in the dark for 30 min and then the titration was started. Each addition of ligand  was followed by a 3 min equilibration time and fluorescence emission spectra were recorded from 510 to 750 nm  and 5-BrdU-click partners (5-BrdU-Alk and 5-BrdU-N3) were prepared in DMSO at 1 mM concentration, and stored in the dark at −20°C. For all the following experiments, copper-catalyst (CuSO4.5H2O), THPTA and sodium ascorbate solutions were freshly prepared in ultrapure H2O at respectively 10 mM for the CuSO4.5H2O solution, 50 mM for the THPTA solution and 40 mM for the sodium ascorbate solution. Oligonucleotide sequences were purchased from Eurogentec as dried samples purified by HPLC-RP. Oligonucleotides were dissolved in MilliQ water at 200 μM concentration and stored at −20°C. Oligonucleotide sequences used for click reaction in situ are the following: c-Myc22 (G14T-G23T): TGAGGGTGGGTAGGGTGGGTAA, 22AG: AGGGTTAGGGTTAGGGTTAGGG and TERRA: AGGGUUAGGGUUAGGGUUAGGG.Tris(3-hydroxypropyltriazolylmethyl)amine (THPTA) and CuSOOligonucleotides were folded and CuAAC reactions were carried out in K10 buffer .P = 0.4 Bar).Solid-phase extraction (SPE) was performed with Clarity® OTX (Phenomenex) columns. To facilitate elution of the fractions, a manifold device as vacuum source was employed  and THPTA  were added. The mixture was stirred and centrifuged to obtain a blue solution. Afterward, sodium ascorbate  was added, and the mixture was stirred and centrifuged again to give a colorless solution. The latter was diluted with K10 buffer (11.6 μl). In a second 1.5 ml Eppendorf tube, the PDC CuAAC derivative  was added to K10 buffer (11.6 μl). The previously prepared copper solution was subsequently added. After stirring and centrifuging, the 5-BrdU partner was added . The reaction mixture was left to stand for 1 h at room temperature without agitation to enable the click reaction to occur. After 1 h, the reaction mixture was treated by SPE.In a 1.5 ml Eppendorf tube, CuSO4.5H2O  and THPTA  were added. The mixture was stirred and centrifuged to obtain a blue solution. Sodium ascorbate  was added and the mixture was stirred and centrifuged again to give a colorless solution. The latter was diluted with K10 buffer (7.85 μl). The copper-containing solution was then added to the G4-containing tube, and the mixture was stirred and centrifuged. Then, 5-BrdU partner was added . The reaction mixture was stirred, centrifuged, and left to stand for 1 h at room temperature without agitation to enable the click reaction to occur. After 1 h, the reaction mixture was treated by SPE.Oligonucleotide solutions (100 μM) were prepared in K10 buffer. The sequences were folded in their G4 structure by heating the oligonucleotide solutions at 95°C for 10 min. The samples were then cooled down in ice for 30 min. In a 1.5 ml Eppendorf tube, the folded G4 , the PDC CuAAC derivative  and K10 buffer (7.9 μl) required for the click reaction were successively added. The mixture was left to stand for 30 min at room temperature without agitation. In another 1.5 ml Eppendorf tube, CuSO2O). The click reaction sample was diluted with 270 μl of H2O to reach a final volume of 300 μl and loaded on the SPE column. Then, small and medium size molecules were eluted by successively washing the SPE column with: 3 × 500 μl of [H2O/MeCN 1:4 (v/v)], 1 × 500 μl of MeCN and finally 1 × 1 ml of MeOH. The loading and elution fractions were collected together in a single 15 ml plastic tube. The solvent was removed under reduced pressure at 30°C for 3 h. The residue was dissolved in 30 μl of ultrapure H2O and analyzed by UHPLC.SPE column was first conditioned as reported by the supplier . The system was equipped with a Phenomenex Luna® OMEGA Polar C18, 1.6 μm, 100 × 2.1 mm column. Elution conditions: column temperature = 40°C, flow rate = 0.4 ml/min, use of a linear A–B gradient  20% B isocratic for 1 min, (b) 20–80% B gradient for 4 min, (c) 80% B isocratic for 1.5 min and (d) 80–20% B gradient for 0.5 min). The chromatograms were recorded at two different wavelengths: 254 and 350 nm.About 3 μl of the reaction solution were injected in an Ultra-High-Performance Liquid Chromatography (UHPLC) apparatus: Thermo UltiMate 3000, equipped with a Thermo UV/visible detector . Enzyme-linked immunosorbent assay was performed with Pierce Streptavidin Coated High Capacity Plates, Clear, 96-Well . For validation of BG4 and binding in competition with G4 ligands, BG4 was used at increasing concentrations  and detected using HRP-conjugated anti-FLAG antibody . For immunofluorescence experiments, BG4 was used at 20 nM and detected using rabbit anti-FLAG  and Alexa Fluor 555-conjugated goat anti-rabbit  antibodies. 5-BrdU was detected using mouse anti-5BrdU  and Alexa Fluor 488-conjugated goat anti-mouse antibodies  or HRP-conjugated goat anti-mouse antibody . Cell membrane was detected using wheat germ agglutinin CF®640R WGA . TMB solutions as chromogenic reagents for peroxidase designed for ELISA techniques were used as suggested by the supplier . Cells were seeded in 8-well chamber slides , and slides were mounted with coverslips using vectashield antifade mounting agent with DAPI .Oligonucleotides were folded in 10 mM Tris.HCl pH 7.4, 100 mM KCl at 5 μM concentration by heating up at 95°C during 10 min and by letting slowly cool down to 25°C. pSANG10–3F-BG4 plasmid encoding the BG4 single chain antibody was a gift from Shankar Balasubramanian , followed by culturing in 2xYT and kanamycin (50 μg/ml) and chloramphenicol (34 μg/ml) at 37°C and 250 RPM. When OD600 reached 0.6–0.8 the temperature was lowered to 20°C, and induction was initiated with 0.5 mM IPTG. After 15 h of induction at 20°C, bacterial culture was harvested by spinning at 4 000 g for 30 min. Pellets were resuspended in 8 ml TES buffer 1  per 100 ml volume expression culture. The mixture was put in ice for 10 min. About 12 ml of TES buffer 2  per 100 ml volume expression culture were added and the mixture was put in ice for 15 min. Cells were centrifuged at 4°C for 10 min at 8000 g. Supernatant was collected and filtered (0.22 μm). The sample is deposited on a column FF Crude His trap  was washed and preequilibrated with PBS. Resin was then washed with 20 CV of washing buffer high salt  and 20 CV buffer low salt . BG4 protein was eluted with elution buffer , buffer-exchanged to PBS and concentrated using 50 ml centrifugal devices with 10 kDa cut-off (PALL Corporation). Protein quality was assessed by running SDS-PAGE (4–20% TGX gel Biorad), and gels were visualized with Coomassie staining. The concentrated protein was purified on HiLoad superdex 75 pg 26/600 Cytiva. Peak protein fractions were confirmed using SDS-PAGE, pooled and concentrated as described above. Final protein concentration was calculated based on absorbance measurements at 280 nm, with an absorbance ratio at 260/280 nm <0.7 indicating no nucleic acid contamination.The BG4-encoding plasmid (Psang10-3f-BG4) was transformed into ELISAs for validation of BG4 binding affinity were performed using a standard method reported in literature, by employing HPLC purified biotinylated sequences purchased from Eurogentec .2PO4), plates were blocked with blocking buffer (3% BSA in ELISA buffer) during 1 h. After blocking, plates were incubated with mouse anti-5BrdU antibody (1/150 dilution) for 1 h. After washing steps with ELISA buffer supplemented with 0.1% Tween 20, goat anti-mouse horseradish peroxidase (HRP)-conjugated antibody (1/5 000) was incubated for 1 h. After washing (ELISA buffer + 0.1% Tween 20), binding was visualized with TMB . Reaction was stopped with 2 N HCl and signal intensity was measured at 450 nm on a FLUOstar microplate reader (BMG Labtech).About 5 μM biotinylated hTelo, c-Myc, TERRA, 22Agmut, TERRAmut, ihTelo and hp were annealed in 10 mM Tris HCl, pH 7.4, 100 mM KCl by slow cooling from 95 to 25°C. Biotinylated oligonucleotides solutions were diluted to 0.1, 1, 5, 10, 20, 50, 75 and 100 nM in 10 mM Tris HCl, pH 7.4, 100 mM KCl and bound to streptavidin-coated plates during 1 h. Immobilized hTelo, c-Myc, TERRA, 22Agmut, TERRAmut, ihTelo and hp were incubated with 250 nM PDC-4,3-BrdU solution in 10 mM Tris HCl, pH 7.4, 100 mM KCl during 1 h. After washing steps with ELISA buffer , plates were blocked with blocking buffer (3% BSA in ELISA buffer) during 1 h. After blocking, plates were incubated with 0, 10, 20, 50, 75 and 100 nM BG4 antibody during 1 h. Washing steps (ELISA buffer + 0.1% Tween 20) were followed by incubation with HRP-conjugated anti-FLAG antibody (1/10 000 dilution) for 1 h. After washing (ELISA buffer + 0.1% Tween 20), plates were incubated with TMB  and reaction blocked as previously described. Signal intensity was measured at 450 nm on a FLUOstar microplate reader (BMG Labtech).ELISAs for binding affinity and specificity were performed using a modified method already reported in literature. About 5 μM biotinylated hTelo and c-Myc were annealed in 10 mM Tris HCl, pH 7.4, 100 mM KCl by slow cooling from 95 to 25°C. Biotinylated oligonucleotides solutions were diluted to 50 nM in 10 mM Tris HCl, pH 7.4, 100 mM KCl and bound to streptavidin-coated plates during 1 h. For competition experiments performed in the presence of G4 ligands , bound biotinylated hTelo and c-Myc were incubated with 250 nM ligand solution in 10 mM Tris HCl, pH 7.4, 100 mM KCl during 1 h. After washing steps with ELISA buffer  and 2 ml of TrypLE Express Enzyme (Life technologies) was added for 10 min at 37°C. Cells were collected in warm medium (addition of 8 ml) by gently pipetting. Add appropriate aliquots of the cell suspension to new culture vessels. Cells were re-suspended in medium and seeded at a density of 0.5–1 million cells in a T75 culturing flask. Cells were counted using a MoxiZ™ mini automated cell counter .A549 (ATCC® CCL-185™) cell lines were purchased from the American Type Culture Collection  and cultured in Dulbecco’s modified Eagle’s medium GlutaMAX™  supplemented with 10% fetal bovine serum  and 1% Penicillin/Streptomycin (Life Technologies) in humidified atmosphere under 5% CO2 in air at 37°C. Cells were split twice a week. Cell splitting: cells were washed with Phosphate Buffer Solution , and 1 ml of TrypLE Express Enzyme (Life technologies) was added for 4 min at 37°C. Cells were collected in warm medium (addition of 9 ml) by gently pipetting. Add appropriate aliquots of the cell suspension to new culture vessels. Cells were re-suspended in medium and seeded at a concentration of 0.2 × 105 cells/ml in a T25 culturing flask. Cells were counted using a MoxiZ™ mini automated cell counter .A2780 cell lines were purchased from the American Type Culture Collection  and cultured in Roswell Park Memorial Institute  supplemented with 10% fetal bovine serum  and 3% of MIX  and 1% Penicillin/Streptomycin (Life Technologies) in humidified atmosphere under 5% CO2 for 24 h, solutions of PDC-derivatives (stock solution at 10 mM in DMSO and solutions at 500 and 2 000 μM prepared in water by subsequent dilutions) were added to the cells to reach final concentrations of 1, 5, 10, 15, 20 and 30 μM. After incubation for 72 h at 37°C, culture medium was removed and cells were washed with PBS. Afterward, 100 μl of TrypLE Express Enzyme was added to each well, followed by 5 min incubation at 37°C until the cells were thoroughly dissociated from the bottom of the plate. About 900 μl of fresh culture medium was added to stop the reaction, and living cells were counted by using a MoxiZ™ mini automated cell counter .A total of 20 000 A549 cells per well in 1 ml of culture medium were seeded in 24-well cell culture plates. After incubation at 37°C in a humidified atmosphere with 5% CO4.5H2O (2.5 μl of a 20 mM solution in water) was mixed with 0.5 mM of THPTA (5 μl of a 50 mM solution in water) and left at room temperature for 1 h. To the previous solution, 5 mM of sodium ascorbate (25 μl of a 100 mM solution in water) was added and left to react for 30 min at room temperature. Subsequently, 5-BrdU derivative was added to a final concentration of 50 μM and the solution added to the cells. Negative controls of CuAAC reaction were obtained preparing the click reaction mixture omitting the addition of CuSO4.5H2O. Slides were washed with PBS and the same immunostaining protocol described above was followed.For immunostaining, 15 000 A549 or A2780 cells were seeded and grown in 8-well chamber slides over 24 h. Cells were treated with 5, 10 and 15 μM of PDC-derivatives. After 4, 8, 16, 24 or 48 h of incubation, cells were washed two times with PBS, fixed with 4% PFA in PBS for 15 min and permeabilized with 0.25% Triton X-100 in PBS for 15 min. For samples containing PDC 5-BrdU functionalized compounds : cells were washed three times with PBS and blocked with blocking buffer (5% BSA in PBS) for 2 h at room temperature, then incubated with mouse anti 5-BrdU (1/250) for 1 h at room temperature in blocking buffer. Slides were then washed three times with PBS and incubated with Alexa Fluor 488 goat anti-mouse (1/500) at room temperature for 40 min in blocking buffer. Slides were washed five times with PBS and mounted. For samples containing PDC CuAAC precursors, cells were incubated with the click reaction mixture in the dark for 1 h. For 500 μl final volume per well in PBS: 0.1 mM of CuSO4.5H2O (2.5 μl of a 20 mM solution in water) was mixed with 0.5 mM of THPTA (5 μl of a 50 mM solution in water) and left at room temperature for 1 h. To the previous solution 5 mM of sodium ascorbate (25 μl of a 100 mM solution in water) was added and left to react for 30 min at room temperature. Subsequently, Cy5-N3  was added to a final concentration of 10 μM. Slides were washed five times with PBS and mounted with coverslips using vectashield antifade mounting agent with DAPI.About 15 000 A549 cells were seeded and grown in 8-well chamber slides  over 24 h. Cells were treated with 5 μM of PDC-4,3-Alk. After 16 h of incubation, cells were washed two times with PBS, fixed with 4% PFA in PBS for 15 min and permeabilized with 0.25% Triton X-100 in PBS for 15 min. Cells were incubated with the click reaction mixture in the dark for 1 h. For 500 μl final volume per well in PBS: 0.1 mM of CuSOSamples were prepared and staining performed as described above. To compare the staining results with the Cy5-functionalized compounds, Cu-catalyzed click reactions were performed in the presence of 10 μM of 5BrdU-derivatives.About 15 000 A549 cells were seeded and grown in 8-well chamber slides over 24 h. Cells were treated with 5 μM of PDC-derivatives. After 16 h of incubation, cells were washed two times with PBS, fixed with 4% PFA in PBS for 15 min and permeabilized with 0.25% Triton X-100 in PBS for 15 min. CuAAC reaction samples were treated as described above to obtained 5BrdU-functionalized compounds. To eliminate RNA-structures, cells were treated with 0.1 mg/ml RNase A in PBS at 37°C for 1 h. For staining of 5-BrdU functionalized compounds, cells were treated as previously described: cells were blocked with blocking buffer (5% BSA in PBS) for 2 h at room temperature, then incubated with mouse anti-BrdU (1/250) for 1 h at room temperature in blocking buffer. Slides were then washed three times with PBS and incubated with Alexa Fluor 488 goat anti-mouse (1/500) at room temperature for 40 min in blocking buffer. Slides were washed five times with PBS and mounted.For immunostaining, 15 000 A549 cells were seeded and grown in 8-well chamber slides over 24 h. Cells were treated with 5 and 15 μM of PDC-4,3-Alk and PDC-4,3-BrdU. After 16 h of incubation, cells were washed two times with PBS and fixed with 4% PFA in PBS for 15 min. Membrane staining was performed by incubation with wheat germ agglutinin CF®640R WGA 5 μg/ml for 10 min at room temperature. Cells were washed two times with PBS and post-fixation with 4% PFA was carried out during 10 min at room temperature. Cells were washed three times with PBS and permeabilized with 0.25% Triton X-100 in PBS for 15 min. Cells were washed three times with PBS and blocked with 10% goat serum in PBS for 2 h at room temperature. Then, they were incubated with BG4 antibody (20 nM) for 2 h at room temperature in 10% goat serum in PBS. Slides were then washed four times with PBS and incubated with rabbit anti-FLAG (1/800) for 1 h in 10% goat serum in PBS. After washing four times with PBS, slides were incubated with Alexa Fluor 555 goat anti-rabbit (1/1 000) in 10% goat serum in PBS at room temperature for 40 min. Slides were washed five times with PBS and mounted.t-tests were performed using GraphPad Prism 7.0. P-values are stated in the figure captions. Significance: ns, P > 0.05, *P < 0.05, **P < 0.01, ***P < 0.001, ****P < 0.0001. For column scatter plots, error bars are the mean ± SD, central horizontal line is the mean.Results are expressed as mean ± SD (unless stated otherwise). Two-tailed non-paired For immunostaining, 15 000 A549 cells were seeded and grown in 8-well chamber slides over 24 h. Cells were treated with 5 and 15 μM of PDC-4,3-Alk and after 16 h of incubation, they were washed two times with PBS and fixed with 4% PFA in PBS for 15 min. Membrane staining was performed as previously described, and after the post-fixation step and washing with PBS, cells were permeabilized with 0.25% Triton X-100 in PBS for 15 min. Cells were washed three times with PBS. Samples containing PDC-4,3-Alk were submitted to CuAAC reaction for 1 h in the dark as previously described. Samples were blocked with 10% goat serum in PBS for 2 h at room temperature. Then, cells were incubated with BG4 antibody (20 nM) for 2 h at room temperature in 10% goat serum in PBS. After 1 h incubation, mouse anti-5BrdU antibody (1/250) was added for the remaining 1 h. Slides were then washed four times with PBS and incubated with rabbit anti-FLAG (1/800) for 1 h in 10% goat serum in PBS. After washing four times with PBS, slides were co-incubated with Alexa Fluor 555 goat anti-rabbit (1/1 000) and Alexa Fluor 488 goat anti-mouse (1/500) in 10% goat serum in PBS at room temperature for 40 min. Slides were washed five times with PBS and mounted.Images were captured using a Leica 3D SIM (Structured Illumination Microscopy) DM6000 upright wide-field microscope at room temperature with a 63×/1.4–0.6 oil  objective using a MetaMorph software. DAPI was excited at 405/60 nm, and fluorescence was measured at 470/40 nm. Alexa Fluor 488 was excited at 470/40 nm, and emission was measured at 525/50 nm. Alexa Fluor 555 (Cy3) was excited at 545/30 nm, and emission was measured at 610/75 nm. CF® 640R (Cy5) was excited at 620/60 nm, and emission was measured at 700/75 nm. Images were acquired with a large field, highly sensitive sCMOS camera Hamamatsu Flash 4.0 v2. Leica 3D SIM DM6000 microscope is equipped with a motorized stage in XYZ and galvanometric stage for fast and precise z-stack acquisitions.1). PDC-4,2-Alk, PDC-4,3-Alk and PDC-4,PEG-N3  reaction in the presence of 4-amino-1-butanol yielding compound 4. Afterward, synthesis of mesylate 5 was achieved by reaction of methanesulfonyl chloride with alcohol 4 at 0°C. Reaction temperature and base molar equivalents play an important role and have to be carefully controlled to avoid elimination reaction. Compound 5 was directly submitted to nucleophilic substitution reaction (SN2) in the presence of sodium azide and final methylation reaction afforded PDC-4,0-N3 with high yield. All PDC CuAAC precursors were successfully obtained by mere precipitation and filtration.Four different PDC CuAAC precursors have been designed and successfully synthesized. These derivatives bear either an alkyne or an azide functional group on the linker connected to the central pyridine of the PDC core. Four linkers, differing by nature and size, were synthesized to enlarge the chemical diversity of the compounds. All derivatives were prepared starting from a chloro PDC precursor Scheme . PDC-4,23 Figure ,42. The O-position of the deoxyribose ring with an azide (5-BrdU-N3) or an alkyne (5-BrdU-Alk) group by following respectively a four-step and a three-step synthesis . In order to achieve the synthesis of 5-BrdU-Alk, the hydroxyl group in position 3′ was deprotonated by using NaH, and the obtained alcoholate was then reacted with propargyl bromide, obtaining 3′-O-propargyl derivative 8. A final deprotection step, carried out in the presence of TBAF, yielded final compound 5-BrdU-Alk. To functionalize the 3′-O-position of the sugar with an azide and obtain 5-BrdU-N3, we used a modified Pummerer rearrangement  as mixture of solvent. Catalytic amounts of copper(I) salts, large excess of reducing agent, and inert atmosphere ensure the click reaction to proceed smoothly. Lastly, we excluded copper ligands from the reaction mixture to facilitate final product purification. Residues of copper ions were disposed by gently shaking the reaction mixture with Chelex resin buffered at pH 7. After elimination of the latter, 5-BrdU immuno-tag PDC ligands PDC-4,2-BrdU, PDC-4,3-BrdU, PDC-4,PEG-BrdU and PDC-4,0-BrdU were precipitated and obtained as pure compounds.5-BrdU PDC conjugates Figure  were obtO-modified 5-BrdU derivatives and 5-BrdU immuno-tag PDC ligands were isolated with high purity, as evidenced by analytical HPLC, NMR spectra and high-resolution mass spectrometry.PDC CuAAC precursors, 3′-T1/2) observed in the presence of the telomeric sequence are strong in K-rich buffer . As anticipated, the presence of the immuno-tag somewhat affects G4 ligand binding properties by decreasing ΔT1/2 values of ∼10–15°C, but this effect is minor and the high values observed still denote the strong G4 binding ability of the conjugates in particular considering the condition of the assay (low concentration in G4 DNA). Importantly, the functionalization borne by the ligands does not affect selectivity toward G4 DNA as compared to duplex DNA  melting assay ,45. For A Figure . A similA Figure : the preTo further confirm G4 structure selective binding, similar experiments were repeated in the presence of two G4 RNA forming sequences: the telomeric repeat-containing RNA sequence (F21RT) and the human NRAS 5′-UTR sequence (F-NRAS-T) ,52. The 50 values . The values obtained for 5-BrdU immuno-tag modified PDCs are slightly increased, mainly while interacting with 22AG, 21CTA and CEB25wt (DC50 range = 0.5–1.4 μM). This behavior can be explained by the interaction of 5-BrdU with the loops. The presence of lateral loops in the 22AG and 21CTA structures as well as the 9 nucleotide long loop of CEB25wt may affect ligand positioning on the external quartets.To gain additional understanding on the binding affinities of the synthesized G4 ligands, G4-FID assay was performed for all compounds in the presence of the aforementioned unlabeled G4 sequences  53). Th. Th53). In the whole the two biophysical assays revealed fully consistent and highlighted the strong affinity and the remarkably high selectivity of all PDC derivatives toward G4 structures of various topologies.4 (500 μM), THPTA (2.5 mM) and sodium ascorbate (2 mM) was prepared. The two solutions were then combined and 5-BrdU click partner (50 μM) was added. A control reaction in the absence of G4 target was carried out in the same conditions to analyze reaction conversions at a defined reaction time. We compared the reactivity after 1 h of incubation, time at which, in the absence of G4 target, the conversion is about 50% for most PDC derivatives  was added. AbI was then detected by HRP-conjugated secondary antibodies (AbII). Finally, an enzymatic substrate was added to allow chromogenic detection of AbII thus revealing the 5-BrdU-functionalized ligand associated to G4 Figure .Since all 5-BrdU immuno-tag PDCs showed similar biophysical properties toward the different prototype G4 structures, we selected PDC-4,3-BrdU as a reference compound and as biotinylated sequences to perform the assay: hTelo, c-Myc and TERRA as examples of G4s, a hairpin (hp) as example of a duplex, 22Agmut and TERRAmut as examples of G-rich sequences not able to fold in a G4 structure and ihTelo as example of a C-rich sequence. Remarkably PDC-4,3-BrdU was successfully detected when bound to the three G4 structures in a clear G4-concentration dependent manner Figure . The ligKd) of PDC-4,3-BrdU for hTelo, c-Myc and TERRA: Kd[hTelo] = 9.6 ± 1.3 nM, Kd[c-Myc] = 3.5 ± 0.9 nM, and Kd[TERRA] = 29.6 ± 7.8 nM respectively. All Kd values are lying in the low nanomolar range confirming the exceptionally high affinity of this ligand toward G4 structures. To be highlighted, this affinity is close to that observed for the BG4 antibody. Importantly this biochemical assay provided the undisputable proof of concept that 5-BrdU immuno-tag PDCs can be recognized and visualized by antibodies, when bound to G4 targets, thereby fully validating our G4-ligand guided immunostaining strategy.Using this adapted ELISA assay, we were able to determine the binding constants . All PDC derivatives were incubated with A549 cells at three subtoxic concentrations  over a period of 16 h to verify the effect produced by ligand accumulation on the detection method. The fluorescence signal of the cells was monitored by using an upright wide-field microscope. Concisely, after ligand treatment, cells were fixed with paraformaldehyde and permeabilized to allow antibody penetration, then immuno-tag PDC treated cells were directly incubated with anti 5-BrdU primary antibody and Alexa Fluor 488 labeled secondary antibody. Differently, PDC CuAAC precursor treated cells were first submitted to click reaction for 1 h in the dark, optimal reaction conditions were identified as followed: 50 μM of 5-BrdU-N3 or 5-BrdU-Alk partner, 100 μM CuSO4·5H2O, 500 μM THPTA and 5 mM sodium ascorbate. Afterward, cells were treated with antibodies as described above. Interestingly cells incubated with 5-BrdU tagged PDC conjugates or PDC CuAAC precursors assembled in situ with BrdU partners displayed a more or less diffuse fluorescent signal in the cytoplasm together with fluorescent spots (foci) of variable intensity within the nucleus  . To confTaken together, these experiments demonstrate that 5-BrdU immuno-tag modified G4 ligands can be efficiently detected in cells by the G4-GIS methodology and led to the selection of PDC-4,3-Alk as the best candidate for the new labeling approach. Of crucial importance, this method allowed identifying RNA and most likely G4 RNA as the main target of PDC-4,3-Alk.To investigate further the ligand distribution specificity with regard to G4 structures, immunofluorescence experiments in the presence of the G4 specific antibody BG4 were performed ,58. At fKd[hTelo] = 7.8 ± 1.5 nM and Kd[c-Myc] = 21.1 ± 4.6 nM (Kd[hTelo] = 1.6 ± 0.1 nM and Kd[c-Myc] = 1.5 ± 0.2 nM) (Kd[hTelo] = 8.0 ± 1.9 nM), and on the contrary, was slightly improved by the presence of the two benchmark G4 ligands  , and PhenDC3 remarkably improved the binding (Kd[c-Myc] = 9.5 ± 3.2 nM)  . BG4 bin) Figure . Differe) Figure .in vitro data suggest that the nature of the G4 ligand used and the topology of the G4 structure may influence BG4 recognition in cells. We hypothesize that at lower concentration PDC-4,3-Alk targets G4 structures for which it has the strongest affinity thus competing with BG4 binding, as suggested by the Kd values determined for its conjugate PDC-4,3-BrdU. Differently, at higher concentration PDC derivatives can bind, a larger panel of G4 structures of high and low affinity and also promote their formation, altogether increasing BG4 foci.These in situ functionalization by CuAAC reaction, in vitro experiments have demonstrated that 5-BrdU functionalized ligands can be recognized and visualized by specific antibodies, once bound to the G4 target. These recognition properties were further verified in A549 cells, as evidenced by the generation of a fluorescent signal detected by microscopy, validating the immunofluorescence methodology. Notably, in our experimental conditions, immunofluorescent experiments showed preferential ligand cytoplasmic accumulation in defined regions (foci) with low ligand nuclear localisation and RNase A treatment highlighted RNA as the main target of the synthesized ligands. The fluorescent signal amplification guaranteed by secondary antibodies allowed to improve spatial precision and sensitivity and clearly detect perinuclear accumulation of PDC-4,3-Alk with formation of fluorescent foci. Additional immunofluorescence experiments, aiming to the detection of BG4 foci, put in evidence that the presence of a G4 ligand bound to a specific G4 topology may affect BG4 recognition, and as a consequence, BG4 foci detection. Furthermore, G4-GIS methodology showed the impossibility to perform colocalization experiments between BG4 and anti 5-BrdU antibody emphasizing on the strong dependency between secondary antibody binding and spatial proximity of the two primary antibody targets. In a broader perspective this study provides the research basis for the application of immunofluorescence methodology and associated signal amplification to visualize immuno-tag modified-compounds bound to their cellular target and improve the spatial detection of small molecules functionalized with fluorescent probes.In this work, we have developed a new approach for guiding detection of G4 ligands in complex with G4 structures in cells. The new G4-ligand guided immunofluorescence staining (G4-GIS) approach is based on the specific recognition of 5-BrdU immuno-tag modified-G4 ligands by a commercially available primary antibody. After validation of G4 ligand binding properties toward G4 structures by biophysical assays and confirmation of gkab1166_Supplemental_FilesClick here for additional data file.


Telomeres are copied and reassembled each cell division cycle through a multistep process called telomere replication. Most telomeric DNA is duplicated semiconservatively during this process, but replication forks frequently pause or stall at telomeres in yeast, mouse and human cells, potentially causing chronic telomere shortening or loss in a single cell cycle. We have investigated the cause of this effect by examining the replication of telomeric templates in vitro. Using a reconstituted assay for eukaryotic DNA replication in which a complete eukaryotic replisome is assembled and activated with purified proteins, we show that budding yeast telomeric DNA is efficiently duplicated in vitro unless the telomere binding protein Rap1 is present. Rap1 acts as a roadblock that prevents replisome progression and leading strand synthesis, but also potently inhibits lagging strand telomere replication behind the fork. Both defects can be mitigated by the Pif1 helicase. Our results suggest that GC-rich sequences do not inhibit DNA replication per se, and that in the absence of accessory factors, telomere binding proteins can inhibit multiple, distinct steps in the replication process. Telomeres are dynamic nucleoprotein structures that protect and maintain the ends of eukaryotic chromosomes and are composed of short GC-rich sequences repeated over hundreds (yeasts) or thousands (humans) of base pairs. Telomeric defects or deletions from even a single chromosome end are associated with loss of terminal DNA sequences, breakage-fusion-bridge cycles and tetraploidisation, driving karyotype changes and genome instability ,2. To prA diverse range of telomeric properties have been proposed to contribute to this effect, including t-loops  telomeriMuch work in this area has focussed on the impact of telomeric DNA itself. Single stranded G-rich sequences from yeast and human telomeres can assemble into stacks of four planar guanosine residues called G-quadruplexes (G4s) . G4s canA second potential barrier likely to be encountered at most telomeric positions is the collection of DNA-bound proteins that coordinate essentially all aspects of telomere function. Elsewhere in the genome, DNA-bound proteins act as replication roadblocks that stall or pause the replisome . GeneticTo examine directly the impact of different telomeric properties on DNA replication, we have analysed budding yeast telomere replication in vitro. Using a replication system in which a complete eukaryotic replisome is reconstituted with purified proteins, we show that both leading and lagging strand synthesis proceed to completion across budding yeast telomeric DNA unless the telomere binding protein Rap1 is present. Rap1 acts as a replication roadblock that inhibits fork progression, but also induces a penetrant defect in lagging strand synthesis behind the fork. Our results suggest that unwinding of budding yeast telomeric sequences by the replisome alone is unlikely to inhibit DNA replication, but that DNA-bound proteins can act as potent inhibitors of multiple steps in the telomere replication process.In vitro replication of control templates lacking telomeric DNA employed construct MD154, which is based on a GC209, a pUC19 vector containing an insert composed of two inverted optimal ORC binding sites spaced 70 bp apart  for 3 h at 37°C. ScaI-HF digested DNA was used for all reactions except those in Figure d-thio-galactopyranoside (IPTG) was added to 1 mM. After 4 h at 25°C, cells were collected by centrifugation, washed once with phosphate buffered saline (PBS), frozen in liquid nitrogen and stored at −80°C. For lysis, a cell pellet from a starting culture volume of 2 l was resuspended in 30 ml lysis buffer  + 500 mM NaCl, supplemented with 0.5 mM AEBSF, 10 μg/ml Leupeptin, 10 μg/ml Pepstatin A. Lysozyme (Sigma) was added to 0.5 mg/ml and the sample incubated on ice for 20 min. After sonication for 5 min , crude lysate was centrifuged at 23 600 g for 30 min, 4°C, and the supernatant mixed with 1 ml packed volume of glutathione sepharose (Sigma) prewashed in lysis buffer + 500 mM NaCl. After 1.5 h incubation at 4°C, beads were washed with ∼50 column volumes (CV) lysis buffer + 500 mM NaCl, resuspended in 1 CV lysis buffer + 500 mM NaCl supplemented with 0.04 mg/ml 3C protease and incubated at room temperature for 30 min. The flow through was collected, and additional cleaved protein was eluted with four consecutive CVs of lysis buffer + 500 mM NaCl. Rap1-containing eluates were pooled, slowly diluted 3.3× with lysis buffer lacking NaCl to a final salt concentration of 150 mM. and applied to a Mono Q 5/50 GL column (Sigma) preequilibrated with lysis buffer + 150 mM NaCl. Bound protein was eluted over a 20 CV gradient from 150 to 1000 mM NaCl, Rap1 containing fractions were pooled, concentrated to 0.5 ml on a 30 kDa cutoff ultra-4 centrifugal filter (Amicon) and loaded onto a Superdex 200 Increase 10/300 GL column (Sigma) preequilibrated with lysis buffer +150 mM NaCl. Rap1 containing fractions were pooled, concentrated, frozen in liquid nitrogen and stored at −80°C prior to use.ORC, Cdc6, Mcm2-7:Cdt1, DDK, Dpb11, GINS, Cdc45, polymerase ϵ, CDK, Ctf4, RPA, TopoI, PCNA, Mcm10, polymerase δ, polymerase α/primase, Sld3/Sld7, Sld2, Csm3/Tof1, Mrc1 and RFC were expressed and purified as described . Fen1, C32P- labelled dCTP, 2.5 nM DNA template, 10 nM ORC, 25 nM Cdc6, 50 nM Mcm2–7:Cdt1, 40 nM DDK, 45 nM Dpb11, 20 nM GINS, 40 nM Cdc45, 30 nM polymerase ϵ, 30 nM CDK, 20 nM Ctf4, 50 nM RPA, 10 nM TopoI, 70 nM PCNA, 2.5 nM Mcm10, 4 nM polymerase δ, 40 nM polymerase α/primase, 25 nM Sld3/Sld7, 40 nM Sld2, 40 nM Csm3/Tof1, 10 nM Mrc1, 20 nM RFC.MCM loading was carried out in reactions containing a final concentration of 25 mM HEPES pH 7.6, 10 mM MgOAc, 0.02% (v/v) NP-40-substitute, 5% glycerol, 100 mM K-glutamate, 1 mM DTT, 5 mM ATP, 5 nM DNA template, 20 nM ORC, 50 nM Cdc6 and 100 nM Mcm2-7:Cdt1, and incubated for 2 min at 24°C prior to the addition of DDK to 80 nM. After 10 further minutes at 24°C, CDK was added to 20 nM. After 5 further minutes at 24°C, replication buffer was added, the temperature increased to 30°C, and a mix of firing and replication factors added. The final concentrations of reaction components was as follows: 25 mM HEPES pH 7.6, 10 mM MgOAc, 0.02% (v/v) NP-40-substitute, 2.5% glycerol, 220 mM K-glutamate, 0.5 mM DTT, 4.5 mM ATP, 30 μM each dATP, dCTP, dGTP, dTTP, 400 μM each UTP, GTP, CTP, 33 nM α-For replication reactions lacking polymerase δ, the final concentration of K-glutamate was reduced to 100 mM. For replication reactions containing Rap1, Rap1 was added after replication buffer and the reaction incubated for 5 min at 30°C prior to firing factor addition. For the pulse chase reaction in Figure 32P- labelled ATP and 20 U polynucleotide kinase (NEB). After incubation at 37°C for 1 h and heat inactivation at 65°C for 20 min, the sample was desalted over a G-50 microspin column (Sigma) and stored at −20°C prior to use.12.5 μg HindIII-digested phage lambda DNA was dephoshorylated in 1x Cutsmart buffer (NEB) with 10 U Quick-CIP phosphatase (NEB) in a final volume of 30 μl for 1 h at 37°C. After heat inactivation at 80°C for 2 mins, DNA was purified through a spin column (Roche) and 1.4 μg was resuspended in 1x PNK buffer (NEB) supplemented with 1.35 mM γ-Escherichia coli Topoisomerase IV (Topogen) added. After incubation at 37°C for 20 min, loading dye (NEB) was added to 1x, and samples loaded onto a 0.6% agarose 1× TAE gel, which was run at 0.75 V/cm for 18 h, and then dried and imaged by phosphorimaging. For the 2D gel analysis in Figure Replication reactions were quenched by adding an equal volume of stop buffer (see ‘Preparation of DNA templates for in vitro replication’) and incubated at 37°C for 15–20 min. An equal volume of Phenol:Chloroform:Isoamyl Alcohol 25:24:1 saturated with 10 mM Tris, pH 8.0, 1 mM EDTA was added, the aqueous phase was collected after centrifugation for 5 min and the sample desalted using a G-50 microspin column before being processed as follows: samples for analysis by denaturing alkaline agarose electrophoresis were supplemented with 20 mM EDTA, 0.5% sucrose and 50 mM NaOH, incubated at room temperature for 10 mins and loaded onto a 0.7% agarose gel supplemented with 2 mM EDTA and 30 mM NaOH, which was run at 0.75 V/cm for 16 h in 2 mM EDTA and 30 mM NaOH. Gels were fixed in 5% trichloroacetic acid for 30–60 min, dried onto filter paper and imaged by phosphorimaging and autoradiography. Samples for Figure Gel densitometry was performed using original .gel files and ImageJ.32P-labelled ATP. PNK was inactivated by heating at 65°C for 2 min and the sample desalted over a G50 spin column . For the EMSA, labelled DNA was incubated at a final concentration of 2.5 nM with the Rap1 and Pif1 concentrations indicated for 20 min in reactions containing 25 mM HEPES pH 7.6, 10 mM MgOAc, 0.02% (v/v) NP-40-substitute, 2.5% glycerol, 220 mM K-glutamate, 0.5 mM DTT, 4.5 mM ATP. Samples were supplemented with 0.5% sucrose and run on a 1.5% agarose 0.5× TBE gel, which was then dried and imaged by phosphorimaging.20 μg plasmid MD155 was digested with BamHI for 2 h at 37°C in a final volume of 100 μl, 1 μl quickCIP (NEB) phosphatase added and after a further 30 min the reaction quenched with 25 mM EDTA, 0.2% SDS and 0.2 mg/ml proteinase K. After 30 min, an equal volume of phenol:chloroform:isoamyl alcohol was added, the sample centrifuged at full speed for 5 min in a benchtop centrifuge and the supernatant supplemented with NaCl to a final concentration of 300 mM prior to ethanol precipitation as above. Precipitated DNA was resuspended in 10–20 μl 1× TE and run on an 8% tris–borate–EDTA (TBE) polyacrylamide gel, which was then stained with sybrsafe dye (Invitrogen). The band corresponding to the telomeric fragment was excised, fragmented and incubated at room temperature with 300 μl 1× TE supplemented with 300 mM NaCl overnight with shaking, prior to ethanol precipitation as above. Precipitated DNA was resuspended in 1× TE to a final concentration of 150 nM, and end labelled using polynucleotide kinase (PNK) for 1 h at 37°C in a reaction containing 1x PNK buffer (NEB), 40 nM DNA, 10 U PNK, 7.5 μM γ-To examine the causes of telomeric replication stress, we set out to study the replication of telomeric templates using a reconstituted assay for eukaryotic DNA replication . In thisin vivo. To determine whether replication was impaired by this sequence, synthesis of radiolabelled nascent strands was followed via denaturing and native agarose electrophoresis. As shown in Figure To analyse the effect of telomeric DNA on the replication process, a complete 325 bp budding yeast telomere sequence  Supplem was inseTelomeric DNA was positioned internally in these experiments. To exclude that this was responsible for the efficient replication we observed, starting plasmids were digested such that the telomeric tract was positioned proximal to the template end  that are indistinguishable from one-another in the experimental approach used above. To examine the lagging strand in our reactions, we therefore added Fen1 and Cdc9, enabling processing and ligation of OFs together with the leading strand of the opposing replication fork ,27 Figu. In contWhilst we did not reproducibly detect lagging strand defects in the analysis above, lane scans occasionally detected a faint shoulder at the position of the telomere, suggesting lagging strand defects may occur in vitro but are infrequent see e.g. , we examOur telomeric insert contains 325 bp of telomeric DNA and a short non-telomeric G-rich sequence originating from the DNA fragment used for cloning  diagram.Figures in vitro replication was carried out for 15 min in the absence of Fen1 and Cdc9 and products analysed by native and denaturing agarose electrophoresis. As shown in Figure The telomeric fragment examined above contains seven consensus Rap1 binding sites  and was Figure Interestingly, the size of the 5 kb fragment indicates that lagging strand telomeric defects may persist even after replication forks have reached the template end (Figure in vivo, factors known to promote telomere replication within cells may prevent or attenuate these effects. Pausing of replication forks at budding yeast telomeres is increased without the Pif-family helicase Rrm3 (Rap1 is therefore inhibitory for two distinct steps in the replication process: replisome progression Figure , and lagase Rrm3 . Howeverase Rrm3 . Previouase Rrm3 . To examase Rrm3 . Replicaase Rrm3 , lane 3;ase Rrm3 , indicatase Rrm3 , we favoIn reactions containing Fen1 and Cdc9, Pif1 also increased the amount of unit length products formed in the presence of Rap1 Figure , even whSemiconservative replication of telomeres ensures that chromosome ends are protected from one cellular generation to the next. Replication forks pause or stall during this process, perhaps helping to preserve telomere integrity during the replication process. Here, we have used an in vitro system for DNA replication to examine the cause of this effect and found that whilst budding yeast telomeric DNA is efficiently replicated in vitro, the telomere binding protein Rap1 inhibits multiple stages of DNA replication and can be partially overcome by Pif1 helicase.Whilst telomeric DNA was replicated to completion in unperturbed reactions, low concentrations of the G4-stabilizing agent Phen-DC3 were sufficient to induce lagging strand defects over the telomere . As PhenOur finding that Rap1 is a potent inhibitor of telomere replication is consistent with previous in vivo work showing that tethering of Rap1 to an internal chromosomal site induces replication fork pausing  and thatin vivo (in vitro, perhaps indicating that a factor or component is missing or that the conditions in the assay are suboptimal. However, since replication pauses at telomeres even in wild type cells (in vivo, indicating that at least a proportion of forks ‘stuck’ behind Rap1 are processed and cleaved (We have found that the roadblock effect of Rap1 can be partially overcome by Pif1 Figure , a membein vivo . Since Pin vivo ,41, we fin vivo . This moin vivo , and thein vivo , a role in vivo . The mecpe cells , an altepe cells  are a pa cleaved .There are significant differences in the sequences and proteins at telomeres in yeast and human cells. However, like yeast Rap1, TRF1 and TRF2 use Myb-type domains to bind human telomeric DNA , and RTEAll data are available in the main figures and supplementary data file.gkab416_Supplemental_FileClick here for additional data file.


Integrated oncology approaches can improve cancer treatment efficacy, and photothermal-chemotherapy drug delivery nanosystems (DDS) based on nanotechnology and biotechnology have piqued the interest of researchers. This study developed an aptamer-modified graphene quantum dots (GQDs)/magnetic chitosan DDS for photothermal-chemotherapy of HCC. The HCC aptamer and the EPR effect of nanoparticles, in particular, enable active and passive targeting of DDS to HCC. GQDs functioned as photosensitizers, effectively moderating photothermal therapy and inhibiting drug release during blood circulation. Magnetic chitosan demonstrated excellent drug encapsulation, acid sensitivity, and tumor imaging capabilities. Proper assembly of the units mentioned above enables precise combined therapy of HCC. This study indicates that DDS can significantly inhibit tumor growth while also extending the survival duration of tumor-bearing mice. The DDS (DOX-Fe The photosensitizer can transform light energy into heat energy, allowing thermal ablation of the tumor site to be performed.7 Compared to traditional therapy, studies reveal that photothermal therapy offers a shorter treatment time, an apparent curative outcome, and less biological toxicity.8 More importantly, photothermal therapy and traditional chemotherapy have a strong synergistic effect and therapeutic transformation potential.9 The heat generated by photosensitizers can boost the uptake rate of drug carriers by tumor cells, shorten the release time of chemotherapy drugs, avoid drug resistance, and improve chemotherapy efficacy.10 However, precisely targeting chemotherapeutic drugs and photosensitizers to the tumor site simultaneously and managing drug release to obtain the maximal therapeutic effect while avoiding side effects is the key to effective HCC treatment.12Because of its high specificity and minimally invasive advantages, photothermal therapy (PTT) has recently been a research hotspot among therapeutic techniques for HCC.13 They were concentrated in tumor tissues because of the high permeability and long retention effect (EPR effect). Hydrophobic drugs that are water-insoluble can be encased in nanoparticles to make them more water-soluble and avoid the negative effects of standard cosolvents.15In recent years, nano-drug loading technology has demonstrated distinct advantages in tumor treatment. Nanocarriers with 10–200 nm diameter enter tumor tissues without being delivered there by lymphatic system reflux.16 Chemotherapy drugs, such as doxorubicin (DOX), create a chemical bond (N-C-) with CS via the aldehyde group, which allows the drug to remain active for an extended period. In vivo, chitosan does not induce allergic reactions or rejection and can be metabolized into harmless monosaccharides absorbed by the human body.18 Many active amino groups in the molecules are easily cleaved in the low pH tumor microenvironment. As a result of its bioadhesive, biodegradable, and pH-responsive nature, CS is a good option for DDS.20Because of their positive charge on the surface and mucosal viscosity, chitosan (CS) drug-loaded materials easily penetrate the cell membrane, making them one of the best nano-drug carriers.2O3, Fe3O4), and polymers can be easily added to the surface of CS. According to studies, small-sized CS-coated magnetic Fe3O4 can absorb more drug molecules and increase the dispersion and stability of nanoparticles.21 Due to its superparamagnetism and good biocompatibility, magnetic nano-sized iron oxide offers significant application potential in constructing tumor-targeted MRI imaging probes and DDS.23Furthermore, nanoparticles such as quantum dots, magnetic iron oxide particles  light, the conjugated π bond of GQDs absorbs photons. It transforms them into heat, promoting rapid heat of the surrounding environment and producing reactive oxygen species (ROS) to accelerate tumor cell death.26 Furthermore, the surface functional groups  of GQDs allow them to interact with a wide range of nanoparticles, nucleic acids, and proteins.27 GQD appears to be a promising photosensitizer.Photosensitizers are essential for achieving effective photothermal therapy. Traditional photosensitizers, such as noble metal nanoparticles and indocyanine green (ICG), have numerous flaws. Metal materials are frequently poisonous, difficult to break down, and harmful to organisms.3O4 nanoparticles were created using the co-precipitation approach. Lin et al. described the method of CS production by coating magnetic iron with CS.28 To preserve pharmacological action, DOX was loaded into CS via the aldehyde group, and GQDs were changed on the surface of magnetic CS via an amide link. Finally, the TLS11a aptamer with a strong affinity for H22 mouse liver cancer cells29 was employed to functionalize the nanoparticles, resulting in DOX-Fe3O4@CS@GQD-Apt (DOX- Fe3O4@CGA). The DDS can particularly target HCC and has pH sensitivity. Experiments in vivo and in vitro revealed that the DDS effectively treats tumors with synergistic photothermal-chemotherapy, establishing a potential nano platform for HCC therapy.GQDs modified magnetic chitosan chemo/photothermal DDS were produced in this study. As illustrated in Abbreviation: CS, chitosan; CG, Chitosan@GQD; CGA, Chitosan@GQD@Aptamer; DOX, Adriamycin; NIR, Near-infrared light;2.2.14 · 7H2O(Cat.No:215422), PEG-2000(Cat.No:8.21037), Chitosan(CS) (Cat.No:448869) 1-ethyl-3-[3-dimethylaminopropyl] carbodiimide hydrochloride (EDC) (Cat.No: 341006), and N-hydroxysuccinimide (NHS) (Cat.No: 8.04518) were purchased from Sinopharm Chemical Reagent Co., Ltd; NH2- modified TLS11a aptamer 3ʹ-NH2–AAAAAAA CAGCATCCCCATGTGAACAATCGCATTGTGATTGTTACGGTTTCCGCCTCATGGACGTGCTG-5ʹ were synthesized by Shanghai Sangon Biotech Co., Ltd. . Carboxy graphene quantum dots(Cat.No: XF090) were purchased from Nanjing Xianfeng nano Biotechnology Co., Ltd; Adriamycin(DOX) (Cat.No: 3010) was purchased from Shanghai Biochempartner Co., Ltd. .Citric acid(Cat.No:251275), FeSO2.22 in Dulbecco’s modified Eagles’s medium supplemented with 10% fetal bovine serum, 100 U/ml penicillin, 100 μg/ml streptomycin.The hepatoma cell line H22 from BALB/c mice was purchased from the China Center for Type Culture Collection (CCTCC). The cells were cultured in a 37°C with 5% CO2.3Female H-2Kd BALB/c nude mice  were provided by the Animal Experimental Center of Zhejiang Chinese Medical University. The experiment was authorized by the Experimental Animal Ethics Committee of Hangzhou Medical College.2.44 · 7H2O was dissolved and 10 mL of 40 g/L PEG-2000 solution was added. The solution was stirred in a constant temperature water bath at 30°C, and 30 mL dilute ammonia water was added dropwise to form dark green Fe(OH)2 by adjusting the pH to 10. Next, 300 μL of H2O2 was added to oxidize part of Fe2+ to Fe3+, stirred for 10 min, and then was placed in 180°C reactors for 10 h. The reaction solution was removed and washed two times with distilled water and three times with ethanol. Magnetic Fe3O4 nanoparticles were isolated and dried at 37°C by magnetic separation. To prepare Fe3O4@CS, 0.2 g of chitosan was dissolved in 20 mL of 2% acetic acid solution. By adding K2CO3 solution at 60°C, the solution pH can be adjusted to 9. The process of producing Fe3O4 was applied to a chitosan acetic acid solution. After four hours of ultrasonic treatment at 60°C, the deposits were separated by magnets and washed numerous times with ethanol and deionized water to obtain Fe3O4@CS. Slowly, 1 mL of DOX-HCI (2 g/L) solution was added to the dispersion. The mixture was stirred at room temperature overnight. The mixture was centrifuged at high speed and washed with PBS to remove the free DOX, and the residue was dried overnight in a vacuum drying oven to obtain DOX-Fe3O4@CS.In 40 mL of secondary distilled water, 2.8 mg FeSO3O4@CS nanoparticles were measured using Uv-vis. DOX concentration was derived from the standard curve; the drug encapsulation and loading efficiency could be determined based on calculation. EE is defined as the actual doxorubicin loading of nanoparticles referred to as the inputted doxorubicin. DLE is defined as the ratio of the mass of the drug encapsulated in nanoparticles to the total mass of drug-loaded nanoparticles.The standard curve of DOX based on the relationship between DOX concentration in solution and UV absorbance was plotted to measure drug encapsulation efficiency (EE) and loading efficiency (DLE). The absorbance values of DOX encapsulated in FeEE (%) = mass of DOX loaded/mass of DOX inputted×100%DLE (%) = mass of DOX loaded/mass of nanoparticles ×100%2.53O4@CS nanoparticles and 2 mg carboxyl GQDs were added in a 2 mL MES (4-morpholineethanesulfonic acid) buffer solution (pH = 6). After shaking for three hours, the particles were crosslinked overnight using EDC (1-ethyl-3-[3-dimethylaminopropyl] carbodiimide hydrochloride)/NHS (N-hydroxysuccinimide). Then, they were centrifuged and rinsed five times with dispersion liquid. Second, 10 mL aptamer (5 mM) was added to the above solution, and the reaction was allowed to run for 2 h. Following that, 1% BSA was added to seal for 30 min, the mixture was centrifuged at high speed and washed three times, and the aptamer coupled nanoprobes (DOX-Fe3O4@CGA) were obtained via magnetic adsorption.The capping of GQDs and the grafting of aptamers were conduceted via the amide bond. First, 10 mg DOX-Fe2.6The morphology and structure of the prepared nanoparticles were examined by transmission electron microscopy . The nanoparticles’ hydrodynamic particle size, polydispersity index (PDI) were evaluated by a Malvern Zetasizer Nano-ZS . The superparamagnetic properties of the magnetite nanoparticles were performed by a vibrating sample magnetometer . Fourier-transform infrared spectra were measured on a Nexus 670 spectrometer (Thermo Fisher Scientific). Thermogravimetric analysis (TGA) was performed with a Perkin Elmer Pyris1 thermogravimetric analyzer.2.7in vitro release of DOX from DOX-Fe3O4@CGA nanoparticles. Briefly, 0.2 mg of DOX-Fe3O4@CGA was dispersed in 2.0 mL of PBS and then transferred to a dialysis bag (molecular cutoff of 3000 DA). The samples were dialyzed in PBS (80 mL) at 37°C with continuous stirring at pH 5.5 or 7.4, and DOX-Fe3O4@CGA was incubated with or without NIR irradiation  at specified time points. At predetermined time intervals, 1 mL of release medium was removed for analysis and replaced with an equivalent volume of fresh PBS; the amount of DOX released was quantified using a UV-vis spectrophotometer; all experiments were conducted three times.The dialysis method was used to investigate the 2.83O4@CS@GQD solution with a Fe concentration of 0 ~ 0.8 mm was produced and aligned. The T2 * scan sequence parameters for MRI were set as follows: TR = 150 ms, TE = 40 ms, slice thickness = 10 mm. T2 values at various concentrations were obtained through signal processing with a magnetic resonance scanner and a postprocessing system. The relaxivity value (r2) was found by fitting the data points to a slope based on solution concentration and relaxation efficiency.The MRI of nanoparticles in aqueous solutions was performed using a clinical MR scanner . In brief, a Fe2.95 cells per well. Following that, 2 mL of a new medium with DOX or DOX-Fe3O4@CGA (at DOX concentrations ranging from 5 to 20 μg/mL) was added to the culture medium. For 5 min, the photothermal group was exposed to a NIR laser at 808 nm at 2 W/cm.2 Following that, the medium was removed. The cells were washed several times with PBS before being fixed by adding 4% paraformaldehyde for 20 min, removing the paraformaldehyde, adding a suitable amount of DAPI staining for 5 min, and washing with anti fluorescence quencher. Finally, confocal fluorescence microscopy was used to examine them.H22 logarithmic growth phase cells were harvested and cultured overnight in six-well plates at a density of 1 × 102.10In vitro cytotoxicity of Fe3O4@CGA was investigated using MTT assay. H22 cells were grown in cell incubators after being injected into 96 well plates. The medium was then removed, and 100 µL of medium containing various concentrations of Fe3O4@CGA was added. The laser group was exposed to a laser  for 5 min. After incubating for 24 h, it was rinsed three times with PBS before mixing uniformly with 100 µL of 0.5 mg/mL MTT solution. To dissolve the nail Zan crystal, 150 µL of DMSO solution was added and vibrated for 10 min. A microplate reader measured the absorbance value to calculate the relative survival rate of cells.2.116 H22 cells were seeded into the right axilla of BALB/c mice (6–8 weeks). To examine biodistribution, DOX-Fe3O4@CGA was labeled with ICG, a commonly used fluorescent dye that can self-assemble and cross-link with CS.30 When tumors reached 300 mm3, mice were randomly assigned to one of four groups (n = 5): (1) PBS+Laser, (2) free ICG, (3) ICG labeled DOX-Fe3O4@CGA, and (4) ICG labeled DOX Fe3O4@CGA + laser. Group 1 was injected with 100 μL PBS and laser-irradiated. Group 2 received ICG injections at a dose of 2 mg/kg ICG in 200 mL of PBS per mouse. Group 3 received the same dose of DOX- Fe3O4@CGA-ICG as Group 2. Group 4 received the same DOX- Fe3O4@CGA-ICG injection as Group 3 and was laser irradiated. Following the intravenous injection, the mice were sedated with 100 ml of 1% 50 mg/kg sodium pentobarbital  for a standard irradiation procedure. The laser groups were irradiated for 5 min every two days with an 808 nm laser (2 W/cm2) under the guidance of fluorescence imaging, beginning on day 1 (after the first injection for 24 h). The Bruker imaging system  was used to capture fluorescence images of groups 2, 3, and 4. During 5-min irradiation, infrared thermometers were used to record the temperature change of the tumor site in groups 1, 3, and 4.To establish a hepatocellular carcinoma-bearing mouse model, 2.0 × 102.123O4@CGA, and DOX, respectively. DOX-Fe3O4@CGA was injected into groups 4 and 5, and groups 2 and 5 were also exposed to NIR light. DOX dosage was set evenly at 8 mg/kg and administered every two days for five treatments. Tumor volume and mouse body weight were monitored every two days during therapy. The tumor volume (V) was determined using the short and long diameters  of tumor tissue (V = d2 × D/2). All animals were slaughtered with 180 mg/kg sodium pentobarbital after 22 days of treatment, and tumor tissues were weighed and H-E stained to assess anti-tumor activity. To evaluate the biological toxicity of treatment, major tissues from PBS and DOX-Fe3O4@CGA+Laser groups were collected for H-E staining, including heart, liver, spleen, lung, and kidneys. The tumor-bearing mice were established to measure mouse survival after treatment. Mice were divided into five groups (n = 8/group) and subjected to the previously described treatment. The survival times of mice in several groups were reported.The tumor-bearing mice were established using the method outlined above. The mice were then divided into five groups (n = 4/group): group 1 injected PBS via the tail vein, groups 2 and 3 received Fe2.13p < .05 were defined as statistically significant differences. Data were presented as mean ± SD.The independent samples t-test was used to determine statistically significant differences, and *3.3.13O4, Fe3O4@CS, and DOX-Fe3O4@CGA. As illustrated in 3O4 was around 9 nm on average and increased to approximately 26 nm following encapsulation in CS. The average particle size increased to 37 nm after loading DOX, GQD, and aptamers. DOX-Fe3O4@CGA magnetic drug-loaded particles had a nearly spherical morphology and a uniform distribution. However, chitosan, adriamycin, aptamer, and GQD encapsulated on the surface of composite nanoparticles were not visible in TEM pictures because they were composed of components with a relatively light atomic mass, such as C, H, O, and N, and TEM was not capable of detecting such elements. The nanoparticle was further examined using DLS, IR spectroscopy, and thermogravimetry to effectively manufacture the composites. Dynamic light scattering (DLS) analysis demonstrated that Fe3O4@CS  after burning are nearly identical to Fe3O4@CS@GQD, indicating that the nanocomposites were synthesized successfully.Fourier transform infrared (FTIR) spectroscopy was used to determine the chemical functional group composition of the prepared nanomaterials. The infrared spectra  revealed3O4 and DOX-Fe3O4@CGA was evaluated. As shown by the magnetization curves  and field cooling (FC) (FC). DOX-Fe3O4@CGA and Fe3O4 have notable peaks at 228 K and 153 K, respectively, as shown in This study aims to investigate the superparamagnetic characteristics of magnetic nanoparticles. First, the saturation magnetization intensity of Fen curves , the magrcentage , showing3.23O4@CGA. As displayed in r2) of Fe3O4@CGA was determined to be 16.70 m/M/S after the images were processed using the MRI data processing system and fitted with a linear function.A 3.0 T clinical MR scanner was used to assess the magnetic resonance imaging capacity of Fe3.33O4@CS nanoparticles solution. As depicted in The encapsulation efficiency was evaluated using a UV-vis spectrophotometer. DOX’s maximum UV absorption peak, as shown in 3O4@CGA at pH 5.5 and 7.4 were evaluated with and without NIR irradiation. The drug release efficiency of drug-laden nanoparticles was lower at pH = 7.4 than at pH = 5.5, as illustrated in To investigate the drug release features of drug-laden nanosystems, the drug release efficiency of DOX-Fe3.43O4@CGA was studied in H22 cells using laser confocal microscopy. After 4-h incubation, the red fluorescence of free DOX was detected in the nucleus, as presented in 3O4@CGA group. When the incubation time was increased to eight hours, the red fluorescence became brighter, but most still did not co-localize with the nucleus. The NIR laser-triggered drug release of DOX from DOX-Fe3O4@CGA in H22 cells was further investigated. The cells were exposed to an 808 nm laser for  before being incubated for four h. In the figure, there was a strong red fluorescence in the nucleus. The results show that DOX-Fe3O4@CGA maintained its perinuclear localization throughout the experiment and that DOX migrated away from the nanoparticles and into the nucleus in response to NIR laser.The cellular uptake of DOX-Fe3.53O4@CGA nanocarriers, the cell viability of Fe3O4@CGA on H22 cells was detected at a range of Fe ion concentrations by the MTT (3-(45)-dimethylthiahiazo (-z-y1)-35-di- phenytetrazoliumromide) technique. 3O4@CGA on H22 cells was low, and cell survival was greater than 70% even at high doses (800 µg/mL), indicating excellent biocompatibility of Fe3O4@CGA. The cell viability in Fe3O4@CGA+Laser group was decreased due to the photothermal effect of nanocarriers, although it remained greater than 65%. The cytotoxic effect of various treatments after loading DOX on nanocarriers was explored further (3O4@CGA group was more effective in killing tumor cells than DOX group. The DOX-Fe3O4@CGA+Laser group outperformed DOX or DOX-Fe3O4@CGA groups in terms of tumor cytotoxicity (p < .05). When DOX-Fe3O4@CGA+Laser group was supplied with a DOX concentration of 20 µg/ml, cell viability dropped to 37%.To evaluate the biocompatibility of Fe further . Because3.63O4@CGA in the H22 tumor-bearing mice, and a statistical graph of the fluorescence intensity at the tumor site is depicted in 3O4@CGA-ICG group. On the other hand, the fluorescence intensity gradually increased at the tumor site. It had the highest fluorescence intensity at 24 h after injection, then decreased significantly at 48 h, most likely due to the degradation of nanomaterials by metabolism in mice. When the fluorescence intensity was coupled with infrared light, it was higher at the tumor site after injection, implying that the photothermal boosted drug-laden particles aggregated at the tumor site, treating the tumor more effectively.In 3.73O4@CGA group without NIR exhibited relatively minor temperature fluctuations during 5 min. When the tumor site was exclusively bombarded with NIR, the temperature rose from 38°C during the first 80 seconds to 41°C. Within the first 50 seconds, the temperature in the photothermal group increased rapidly to roughly 43°C, then gradually increased to 48°C near the tumor site. Even though the tumor cells are killed at this temperature, the distance between the laser probe and the mice was changed to keep the tumor site temperature between 43°C and 45°C. This allowed for gentle photothermal treatment without causing damage to normal tissues.To determine the susceptibility of mice to various treatments. Temperature changes at the tumor location of mice were monitored using an infrared thermometer. As illustrated in in vivo. As illustrated in 3O4@CGA+Laser group than in the other three, demonstrating that photothermal therapy alone was ineffective. This could be because the photothermal therapy applied in this study was moderate. The DOX group developed tumors slowly for the first eight days, similar to DOX-Fe3O4@CGA+Laser and DOX-Fe3O4@CGA groups; the rapid growth in the latter period may be attributed to drug-resistance development in the animals and ineffective treatment targeting. Due to the synergistic therapeutic impact of photothermal materials, DOX-Fe3O4@CGA+Laser group had smaller tumor volumes than DOX-Fe3O4@CGA group during treatment. From the eighth day on, DOX-Fe3O4@CGA group demonstrated an advantage over DOX group, most likely due to tumor targeting by TLS11a aptamer and EPR effect of nanoparticles. 3O4@CGA+Laser group was much less than that of the other groups. Bodyweight is another useful indicator of the health status of mice and can be used to determine the efficacy of photothermal treatment. As shown in 3O4@CGA+Laser and DOX-Fe3O4@CGA groups increased gradually, indicating a gradual return to health following tumor growth control. PBS and DOX groups were in poor condition and lost significant weight as the tumors progressed. HE staining was utilized to examine organ and tumor damage in mice. As illustrated in 3O4@CGA+Laser groups killed only a few cancer cells. Concurrently, DOX-Fe3O4@CGA group eliminated the most cancer cells. Almost all tumor cells were killed in the DOX-Fe3O4@CGA+Laser group, and visible degenerative changes such as nuclear hemorrhage and nuclear lysis can be seen. As illustrated in The tumor volume, tumor weight, mouse body weight, and survival rate during therapy were evaluated to determine the anti-tumor impact of DDS 3O4@CGA+Laser, DOX, DOX-Fe3O4@CGA, and DOX-Fe3O4@CGA+Laser groups were 36, 36, 39, 59, and 76 days, respectively. The median survival time in Fe3O4@CGA+Laser group was comparable to that in the control group, which could be attributed to the favorable photothermal treatment. In comparison, DOX-Fe3O4@CGA+Laser group dramatically increased the median survival time of mice, demonstrating that photothermal from DDS successfully promoted the chemotherapeutic effect. The DOX group had a shorter median survival time than DOX-Fe3O4@CGA and DOX-Fe3O4@CGA+Laser groups. It demonstrated that DDS is less toxic and effective against tumors.Additionally, the survival time of mice was evaluated under various interventions. Kaplan-Meier curves  revealed4.in vivo or in vitro by varying the size of the synthesized material and the intensity of photothermal therapy. Overall, the constructed DDS is biocompatible and multifunctional, and this study may provide a new technique for future research on photothermal chemotherapy paired with HCC.In summary, a unique DDS based on GQDs and CS has been designed successfully. Due to EPR effect and HCC cell-specific aptamers, DDS targeted HCC both actively and passively. Due to superior photothermal capabilities of GQDs and appropriate surface modification, they may efficiently generate heat in the near-infrared to promote the aggregation of magnetic chitosan drug carriers in tumors and acid-sensitive drug release. Meanwhile, it found no evidence of substantial biological toxicity or adverse effects


F5 and F6 were also highly stable to repeated freeze-thaw cycles, with >99% of FGF-2 load remaining after the third cycle. In addition, F5 and F6 were stable to lyophilisation, and the lyophilised products could be stored at −4, 4 or 18 °C for at least 12 months, with less than 1% loss in mean FGF-2 content. Thus, FGF-2 solution is effectively stabilised against both thermal and processing stressors in the presence of MC and alanine (F5), or MC and HSA (F6). The resultant FGF-2 solutions may be applied as medicinal products or further processed into more advanced medicinal products, e.g., scaffolds, for wound healing and tissue regeneration.Basic fibroblast growth factor (FGF-2) is a highly labile protein with strong potential for tissue engineering. The aim of this study was to develop FGF-2 formulations that are stable against physical stressors encountered in pharmaceutical processing and evaluation. Pharmaceutical excipients, alone or in combination, were added to aqueous FGF-2 (770 ng/mL) solution and the stability of the resulting solutions on storage at 4–37 °C was evaluated. Stability of the solutions to repeated freeze-thaw cycles and lyophilisation was also evaluated, as well as the stability of the lyophilised stabilised protein to storage at −4, 4 and 18 °C for up to 12 months. In all of these experiments FGF-2 was quantified by ELISA assay. The as-received FGF-2, when dissolved in water, was highly unstable, retaining only 50% of baseline protein content after 30 min at 37 °C or 1 h at 25 °C. By contrast, FGF-2 solutions prepared with 0.5%  Basic fibroblast growth factor FGF-2) is an endogenous, 18-kDa heparin-binding protein that promotes cellular proliferation, migration and differentiation of a variety of tissues  is an en, with thFGF-2 is very quickly degraded, making it highly challenging to formulate it into acceptable medicinal products ,16. A clOther alternatives, e.g., to reconstitute lyophilised FGF-2 on demand or to dose FGF-2 more frequently, are achievable only with highly compliant patients. Lou et al. found 31.3% of participants in a clinical trial to be noncompliant, their tendency to administer FGF-2 at higher than prescribed doses raising the risk of serious adverse events . There iDifferent approaches have been reported for preserving FGF-2 in aqueous solutions ,23,24. TA customised approach is required for FGF-2 as there is no ‘one size fits all’ strategy to protein stabilisation. To date, a systematic approach to stabiliser selection has not been reported for FGF-2, and most stability studies have been directed at sustaining FGF-2 activity in cell cultures, with a smaller number aiming to develop sustained release FGF-2 formulations for tissue engineering. However, the capacity of stabilised FGF-2 solutions to withstand the stressors of medicinal product manufacture and testing has not been reported, and there is as yet no published data on long term storage stability, a critical predictor of the commercial viability of a medicinal product formulation.The aims of this study were to apply a systematic approach to identify and optimise stabilisation vehicles for FGF-2, and to evaluate the stability of the final stabilised FGF-2 formulations against thermal and processing stressors encountered in medicinal product manufacture and evaluation. The manufacture of therapeutic protein products commonly involves repeat freeze/thaw cycles and lyophilisation that often cause protein denaturation , while pE. coli expression system) was kindly provided by Essex Bio-Pharmaceutical Co. . Low viscosity sodium alginate was purchased from Buchi Labortechnik AG , sodium chloride, glycine and mannitol were purchased from Ajax Finechem , maltodextrin M180 was sourced from the Grain Processing Corporation , D-glucose was purchased from ChemSupply , methylcellulose USP 4000 was purchased from Professional Compounding Chemists of Australia , and human serum albumin, DL-alanine and hydroxypropyl methylcellulose (HPMC) were purchased from Sigma-Aldrich . Deionised water was supplied by a BOSS water system  and used throughout.Lyophilised, recombinant human FGF-2 . FGF-2 content was determined by diluting a freshly reconstituted solution on ice to 100 pg/mL (dry powder weight) and analysing immediately using an ELISA kit  as per the supplied protocol [n = 19) were linear  and reproducible, with no significant differences detected in the slope  and intercept (p = 0.3119) values.Lyophilised FGF-2 powder as received from the manufacturer was reconstituted in water at 1 mg/mL (based on dry powder weight), and stored as aliquots (20 to 100 µL) in 0.1 mL Eppendorfprotocol . The kit® tubes containing water pre-incubated for 2 h at 4° C, 25 °C or 37 °C  to prepare 50 µL of 315 pg/mL FGF-2 test samples. As this was a preliminary evaluation, the assessment of FGF-2 stability was conducted at a sub-therapeutic concentration (315 pg/mL) in order to conserve FGF-2. The samples (n = 4) were incubated at 4 °C, 25 °C or 37 °C for up to 48 h, and their residual FGF-2 content, analysed by ELISA, was expressed as a percent of the baseline FGF-2 content obtained at t = 0. Samples were deemed sufficiently stable if their FGF-2 content did not fall by greater than 10%.Frozen FGF-2 solution was thawed at 4 °C  and serially diluted with water to 1.7 ng/mL (as determined by ELISA). Aliquots were added to 0.1 mL Eppendorfn = 3) were stored at 25 °C for 2 h, and their FGF-2 content as determined by ELISA was compared to baseline level (t = 0). All samples with an initial FGF-2 concentration of 770 ng/mL were assayed neat as well as diluted 1 in 500 and 1 in 1000 with water, to ensure the residual FGF-2 concentration fell within the range of the standard curve provided by the commercial ELISA kit.Excipients falling into the 4 protein stabiliser categories of salt, sugar, polymer and protein/amino acids  were evaBased on this initial excipient testing, glucose, methylcellulose (MC), human serum albumin (HSA) and alanine were chosen for further studies. FGF-2 aliquots were thawed at 4 °C, serially diluted with water to 2.5 µg/mL (as confirmed by ELISA), and mixed with an aqueous stock solution of the specified excipient(s). Test samples contained final excipient concentrations as specified in Test and control samples were incubated for specified periods at 4 °C, 25 °C, and 37 °C, then analysed for FGF-2 content with the ELISA kit. Samples were also exposed to repeated freeze/thaw cycles by incubating the samples at −20 °C for 24 h followed by 30 min at 4 °C before returning the samples to the freezer again, the cycle being repeated a total of 3 times before sample analysis by ELISA. The effects of lyophilisation were studied by freezing the samples at −20 °C for 12 h and lyophilising the frozen samples over 24 h . The lyophilised samples, stored at −20 °C until analysis, were reconstituted with water to their pre-lyophilisation volume immediately prior to analysis by ELISA. Samples were deemed sufficiently stable if their FGF-2 content did not fall by greater than 10% after processing.n = 3) and lyophilised as 50-µL aliquots in 0.1 mL Eppendorf® tubes. Lyophilised samples were stored at −4° C, 4 °C or 18 °C for up to 12 months, with storage temperatures monitored weekly . At defined storage time points , triplicate samples were reconstituted with 50 µL of water. The reconstituted solution was processed according to the scheme outlined in F1, F5 and F6 immediately after preparation were measured for baseline FGF-2 content via ELISA  and a FGF-2 solutions prepared by dissolving the commercial lyophilised powder in water were found to deteriorate very rapidly on storage, even with refrigeration. Samples exposed to 4 °C were stable for only 2 h while those exposed to 25 °C were stable for 30 min. Half-life for the reconstituted FGF-2 solution (315 pg/mL) was 30 min at 37 °C, 1 h at 25 °C and 8 h at 4 °C . As thisA range of excipients from the classes of salt, sugar, polymer, protein and amino acid  were assw/v was the only salt employed. It significantly reduced FGF-2 degradation after 2 h at 25 °C compared with water alone  after 2 h at 25 °C (w/v (11.9% of baseline FGF-2); nor did maltodextrin at 1 and 10% w/v  or mannitol at 5 and 10% w/v , although all the sugar solutions produced greater stabilisation effects on FGF-2 than water (p < 0.0001). Of the polymers, only MC effectively stabilised the FGF-2 for 2 h at 25 °C , the mean FGF-2 content in the HMPC 0.5% vehicle was 87.9%, which did not meet the criteria for sufficient FGF-2 stabilisation. HPMC 2% w/v and sodium alginate 2% w/v also failed to sufficiently stabilise FGF-2 during 2 h incubation at 25 °C. In the protein/amino acids class, HSA 1 mg/mL, alanine 20 mM and alanine 100 mM were able to stabilise FGF-2 at 25 °C for 2 h (p = 0.2686). Glycine at 20 mM and 100 mM also failed to sufficiently stabilise the FGF-2 solution over 2 h of exposure at 25 °C.Sodium chloride at 0.9% at 25 °C B. Glucosat 25 °C C, with c for 2 h D. Increaw/v, MC 0.1 and 0.5% w/v, HPMC 0.5% w/v, HSA 1 mg/mL, and alanine 20 and 100 mM were chosen for further evaluation. Although HPMC 0.5% w/v did not strictly meet the criteria of an effective FGF-2 stabiliser, it was included due to the mean residual FGF-2 content remaining statistically comparable to the MC solutions.On the basis of these results, glucose 10% w/v was not effective at sufficiently stabilising the FGF-2 solution at all three prescribed storage conditions whereas MC 0.1% w/v showed a strong stabilisation effect , but was not successful at maintaining FGF-2 content above 90% after 2 h at 37 °C. The effectiveness of the polymers in stabilising the FGF-2 solution to the prescribed temperature stressors may be ranked in the decreasing order of MC 0.1% w/v > HPMC 0.5% w/v > MC 0.5% w/w. In the protein/amino class, HSA 1 mg/mL was more effective than alanine 20 mM, and alanine in turn was more effective at 20 mM than at 100 mM in stabilising FGF-2 at all three storage conditions (p < 0.0001). None of the excipients were able to sufficiently stabilise the FGF-2 solution for 2 h at 37 °C, although MC 0.5% w/v, HSA 1 mg/mL and alanine 20 mM produced a greater stabilisation effect when compared to water alone (p < 0.0001).Stabilisation of FGF-2 solutions to temperature stressors was evaluated at 4 °C, 25 °C and 37 °C. Glucose 10% n effect , retainiw/v. w/v indeed resulted in improved stability of FGF-2 at 37 °C (p < 0.0001). Compared with MC 0.1% and MC 0.5%, which retained residual FGF-2 contents of 36.9% and 13.4%, respectively, after 2 h at 37 °C, MC 0.05% could retain 81% of baseline FGF-2 content after similar heat exposure.With MC exhibiting concentration-dependent stabilisation effects on FGF-2, being more effective at 0.1% than at 0.5% , the staw/v with either alanine 20 mM, HSA 1 mg/mL or both alanine 20 mM and HSA 1 mg/mL, were evaluated. To facilitate discussion, the blank vehicles and corresponding FGF-2 formulations will henceforth be identified by their respective ID as listed in As no single excipient was able to sufficiently stabilise FGF-2 against thermal degradation at 37 °C, combinations of the excipients were applied to evaluate for synergistic effects ,32,33. CUnlike F2 , F3  and F4 , there were insignificant losses of FGF-2 content in F5, F6 and F7 following incubation for 5 h at 4 °C, 5 h at 25 °C and 2 h at 37 °C . This inp = 0.0929) and F4, while more stable than F1 (p < 0.0001), demonstrating a residual FGF-2 content of 32.5% which was unacceptably low.F2, F5, F6 and F7 were stable against repeated freeze/thaw cycles, with ≥99.8% baseline FGF-2 measured in these samples after the third cycle of processing . By compp = 0.0410). The collective data suggest that having the three excipients in combination (F7) was no more effective in stabilising the FGF-2 solution than the dual combinations of MC with alanine (F5) or MC with HSA (F6). Thus, only F5 and F6 were progressed to further storage stability studies.There was substantial loss of FGF-2 content following lyophilisation of F2, F3 and F4, all of which contain a single stabiliser excipient . By contp = 0.2503), and were 768, 769 and 771 ng/mL, respectively.F5 and F6 were lyophilised and evaluated for storage stability as lyophilised powders and as reconstituted solutions prepared by adding water to the lyophilised powders. F1  served as the control. The lyophilised FGF-2 powders were stored at −4 °C, 4 °C and 18 °C for specified periods of up to 12 months, and were deemed sufficiently stable if the FGF-2 content, as measured with ELISA following reconstitution of the powders with water, did not differ by greater than 10% from baseline FGF-2 content measured in the pre-lyophilised solutions. The mean baseline FGF-2 contents for F1, F5 and F6 were comparable (p = 0.9367). When the storage temperature was increased to 4 °C, the FGF-2 content in F1 was maintained for only 9 months, with further storage leading to undetectable levels of FGF-2 in the lyophilised F1 powder at 12 months. Storage at the higher temperature of 18 °C caused the FGF-2 content of the lyophilised F1 powder to fall below the detectable limit at 3 months. By comparison, F5 and F6 were not only stable to lyophilisation, but the lyophilised F5 and F6 powders were also stable to storage at −4 °C, 4 °C and 18 °C for up to 12 months. The FGF-2 contents in the lyophilised F5 and F6 powders remained above 99% of baseline levels throughout the duration of the prescribed storage conditions.Immediately following lyophilisation, only 7.1% of the baseline FGF-2 content of F1 had remained, representing a significant drop in FGF-2 content . This FGThe lyophilised F1, F5 and F6 powders following storage at −4 °C, 4 °C or 18 °C for specified time points were reconstituted with water and the resultant solutions were stored at either 4 °C or 18 °C for 24 h or 7 days. Solutions prepared by reconstituting the lyophilised F1 powder were not stable to storage at 4 °C or 18 °C for 24 h, the FGF-2 content in all F1 reconstituted solutions falling below the detectible limit of the ELISA assay (data not presented). By comparison, the solutions reconstituted from the lyophilised F5 and F6 remained stable for not only 24 h  but alsoData from this study highlight the need for effective stabilisation strategies to allow FGF-2 to be formulated into viable, safe and effective medicinal products. The commercial lyophilised FGF-2 used in this study was recommended by the manufacturer to be reconstituted with water, however, the FGF-2 reconstituted in water alone (F1) underwent a rapid loss of FGF-2 content, as detected by ELISA, at ambient and refrigerated temperatures, indicating a loss of functionality. Our findings are congruent with published data, which have shown FGF-2 solutions to lose 50% functionality after just 46 min at 25 °C, and the FGF-2 functional half-life to decrease to 37 min when the storage temperature was increased to 37 °C ,35. Intew/v has been shown to negate the stabilising effects of heparin on FGF-2 by disrupting the binding of heparin to FGF-2 [w/v, and this concentration was employed for the stability study. NaCl at 0.9% w/v did not stabilise the FGF-2 solution against thermal degradation, nor did it destabilise FGF-2 compared with the F1 control. In contrast, human prior protein had reduced stability in the presence of low NaCl concentrations, possibly due to the ion-induced destabilisation of salt bridges within the protein [Although no accepted limits have been set for the stability of biological products in any of the current regulatory guidelines, with stability limits instead determined on a molecule-by-molecule basis, this study adopted a cut-off of 90% FGF-2 content remaining as being indicative of sufficient FGF-2 stability for therapeutic use. This cut-off was chosen on the basis of the high quality assurance standards required for pharmaceutical products , as wellto FGF-2 . However protein . The FGF protein , perhapsw/v would have higher solution viscosity and thus will be better able to minimise protein aggregation than glucose 5% w/v.Sugars are well known protein stabilisers, and are widely employed as cryoprotectants. Sugars increase the surface tension and viscosity of solutions to protect proteins against aggregation . They alAmino acids with no net charge, such as alanine and glycine, are also common cryoprotectants of protein formulations, providing stability through weak electrostatic interactions . In thisProteins like HSA and bovine serum albumin are able to stabilise the structure of other proteins through ionic, electrostatic and hydrophobic interactions ,27, and The polymers investigated in this study, MC, HPMC and sodium alginate, are all hydrogels ,51,52 poLyophilisation is a common technique for prolonging the shelf life of commercial protein products which are insufficiently stable to storage as aqueous solutions . The proLyophilisation involves two major stressors, freezing and drying, with both processes capable of damaging a protein structure. Many studies have shown that a stabiliser effective at protecting a protein against freezing may not be effective at stabilising the protein against the process of lyophilisation . This waLong term stability of the FGF-2 solutions was conducted in an incubator set at 18 °C in this study as the ambient temperature in the laboratory fluctuated widely over the course of the study. A temperature of 20–25 °C is more widely accepted as representative of ambient temperature, and future stability studies may be conducted at 25 °C for the stabilised FGF-2 solutions. However, as there was no significant difference in stability following storage of the stabilised FGF-2 solutions for 5 h at 4 °C and 5 h at 25 °C , there iThe focus of this study was the stabilisation of FGF-2 for direct therapeutic use as a medicinal product, and to enable FGF-2 processing into more complex medicinal products. It is recognised that this research also has implications for cell culture and other basic research applications where FGF-2 stability is of concern. However, as the conditions related to these applications, including different applied FGF-2 concentrations and media, were not assessed in this study, further research would be required. Nonetheless, the stabilised FGF-2 solutions and lyophilised powders identified in this study may provide a foundation for further investigation.The quantification of FGF-2 is integral to this stability study. FGF-2 has been detected by high performance liquid chromatography (HPLC) and Western Blot ,58,59,60Pharmaceutics October 2021).Although the ELISA results for stabilised FGF-2 aqueous solutions presented in this study appear indicative of a greater retention of FGF-2 functionality than control, the in vitro efficacy of these formulations must be confirmed prior to the clinical translation of this research. This research is the subject of a follow-up study evaluating the in vitro efficacy of stabilised FGF-2 solutions for wound healing applications  and F6  formulations would be preferred. F5 and F6 are viable formulations of FGF-2 that are applicable as medicinal solutions, or for further processing into more sophisticated medicinal products as they are stable to a range of temperatures, freeze/thaw cycles, lyophilisation, and storage as lyophilised powders and aqueous solutions reconstituted from the lyophilised powders.This study has clearly demonstrated the advantage of combining excipients to stabilise FGF-2 against thermal and processing stressors encountered in medicinal product manufacture and testing, and further provided data on long term storage stability. Although the combination of MC, alanine and HSA (F7) had performed well in the stability studies, the data suggested no advantage to using a combination of all three stabilisers. Considering it is desirable from a toxicity and cost perspective to use the least number and lowest concentrations of excipients to achieve a specified outcome for a medicinal product, the simpler F5 (FGF-2, 0.05% This work is the subject of Patent Application AU 2021902450.