At the ultrastructural level, several atypical and enlarged nuclei and evident nucleoli were also observed. These transformed keratinocytes were apoptosis resistant, exhibited enhanced clonogenicity capacity, increased DNA damage and inflammation, defective DNA repair ability, and increased expression of the oncogene ΔNp63α and intranuclear β-catenin accumulation , compared to those treated with UVA alone. None of these molecular, morphological, or functional effects were observed in cells treated with HCTZ alone. All these features resemble in part those of preneoplastic lesions and NMSCs and provide evidence of a biological plausibility for the association among exposure to UVA, use of HCTZ, and increased risk of NMSCs. These results are of translational relevance since we used environmentally relevant UVA doses and tested HCTZ at concentrations that reflect the plasma levels of doses used in clinical practice. This study also highlights that drug safety data should be followed by experimental evaluations to clarify the mechanistic aspects of adverse events.Recent studies reported the association between increased risk of nonmelanoma skin cancers (NMSCs) and the use of hydrochlorothiazide (HCTZ), one of the most commonly prescribed diuretic, antihypertensive drug, over the world. Although HCTZ is known to be photosensitizing, the mechanisms involved in its potential prophotocarcinogenic effects remain unclear. Under acute exposure, therapeutically relevant concentrations of HCTZ  amplified UVA-induced double-strand breaks, oxidative DNA, and protein damage in HaCaT human keratinocytes, and this effect was associated to a defective activity of the DNA repair enzyme, OGG1. Oxidative damage to DNA, but not that to proteins, was reversible within few hours. After chronic, combined exposure to HCTZ (70 ng/mL) and UVA (10 J/cm Skin photosensitivity reactions, frequently associated to UVA exposure, were indeed reported in patients taking HCTZ –17. A coHowever, these studies were focused on acute effects and did not investigate the biological pathways linking the photosensitizing/phototoxic properties of HCTZ to its potential prophotocarcinogenic effects.Since NMSCs  arise from chronic, repeated UV exposures , 23, 24,On these basis, we characterized both the acute and the long-term, molecular, morphological, and functional consequences of the combined exposure to HCTZ and UVA in human keratinocytes.μg/mL streptomycin and maintained at 37°C in a humidified atmosphere containing 5% CO2. The cell line was authenticated by BMR Genomics  before manuscript submission.Human keratinocytes  were cultured in Dulbecco's Modified Eagle's Medium , supplemented with 10% fetal bovine serum, l-glutamine (2 mM), 100 unit/mL penicillin, and 100 2 of UVA light with an emission centered at 365 nm, using a UV Bio-Link (BLX) , on cold plates to eliminate UVA-induced thermal effects and without lid. The time of irradiation lasted in 40 minutes. Before irradiating the cells with UVA, the medium was removed, and the cells were covered with a thin layer of phosphate buffer solution (PBS). In the HCTZ+UVA group, cells were preincubated with HCTZ (70-370 ng/mL) for 2 hours (h), exposed to 10 J/cm2 of UVA, and then treated with HCTZ for additional 1 or 6 h.The experimental flow chart for short-term experiments is depicted in 5 cells, corresponding to 14,000 cells/cm2, were seeded in Petri dishes, allowed to grow for 24 h, and then treated as follows: cells maintained in standard culture conditions (naive); cells chronically treated with HCTZ alone (70 ng/mL) for 9 weeks (HCTZ); and cells irradiated with UVA alone (10 J/cm2), twice a week, for 9 weeks (UVA) or in the presence of HCTZ (70 ng/mL) (HCTZ +UVA). Each single exposure lasted for 40 minutes. Fresh HCTZ was added every 2 days. The concentration of 70 ng/mL is similar to that found at the steady state following the administration of the dose of 75 mg of HCTZ in humans [The experimental flow chart summarizing the weekly protocol for long-term experiments is depicted in n humans , and then humans .3 cells/well corresponding to 10,000 cells/cm2) were seeded and after 24 h exposed to HCTZ (70-370 ng/mL) with or without UVA irradiation (10 J/cm2). Cytotoxicity was evaluated after 24, 48, and 72 h by using the MTS reagent  as previously described [HaCaT cells  were seeded and after 24 h exposed to HCTZ (70-370 ng/mL) with or without UVA irradiation (10 J/cm2). Carbonyl residues were determined following the method by Correa-Salde and Albesa [HaCaT cells  were seeded and after 24 h exposed to HCTZ (70-370 ng/mL) with or without UVA irradiation (10 J/cm2). Total DNA was extracted using the DNeasy Mini Kit , denatured at 95°C for 5 minutes, and digested with nuclease P1 and alkaline phosphatase at 37°C [HaCaT cells  were seeded and after 24 h exposed to HCTZ (70-370 ng/mL) with or without UVA irradiation (10 J/cm2). The activity of OGG1 was determined by a protocol set-up in our laboratory on the basis of Hamann and Schwerdtle [HaCaT cells . Total protein (1 μg) or 2 units of recombinant hOGG1, used as positive control , were incubated in a reaction mixture containing 1X NEBuffer 2, 40 pmol of synthetic oligonucleotide, and 100 μg/mL BSA, at 37°C for 30 min and then at 95°C for 5 minutes to stop the reaction. A negative control, containing only the reaction mixture was also used. Samples were analyzed by electrophoresis in 3% agarose gel, in TBE 0.5X. The image of residual and intact (negative control) bands was acquired and analyzed through the software Quantity One (Bio-Rad). The percentage of residual oligo was calculated on the basis of the following formula: intact oligonucleotide : 100 = cleaved oligonucleotide : x. The OGG1 glycosylase activity was calculated as follows: 100–x.5′CAATAATAACACGCXCGACCAGTCCTGCTTTTGCAGGACTGGTCGCGCGTGTTATTATTG-3′, 5 cells/well corresponding to 75,000 cells/cm2) were seeded and after 24 h exposed to HCTZ (70-370 ng/mL) with or without UVA irradiation (10 J/cm2). RNA was extracted with TRIzol  and reverse-transcribed using the RevertAid RT kit . Primers were designed on the basis of the human GenBank sequences, and GAPDH was coamplified as the reference [ΔNP63α F: TGCCCAGACTCAATTTAGTGAG, R: TCTGGATGGGGCATGTCTTTGC; OGG1, F: TATCGTGCCCGTTACGTGAG R: CACTGAACAGCACCGCTTG; COX-2 F: TTGCCCGACTCCCTTGGGTGT, R: CCTCCTGCCCCACAGCAAACC; and APE1 F: AGATCTCGCGAGTAGGGCAAC, R: TCCGAAGGAGCTGACCAGTA.HaCaT cells  were seeded and after 24 h exposed to HCTZ (70-370 ng/mL) with or without UVA irradiation (10 J/cm2). PGE2 levels were measured in the supernatants using a commercial kit  according to the manufacturer's specifications.HaCaT cells  were seeded. For the determination of γ-H2AX foci, serum starved, HaCaT cells at confluence, were used. Cells were fixed in 4% paraformaldehyde for 15 minutes, dehydrated in alcohol and paraffin embedded. Immunochemistry was performed on histological slices (4 μm thick) as described by Cinci et al. [β-catenin ; polyclonal rabbit anti Bcl-2 ; and phosphohistone H2A.X (Ser 139) rabbit antibody ; and secondary antibodies are as follows: Alexa Fluor 488 goat anti-mouse or Alexa Fluor 594 goat anti-rabbit  1.200. The analysis of γ-H2AX foci was performed after 1 h since this was previously identified as the time for maximal foci induction in HaCaT cells [γ-H2AX foci if they displayed more than 10 discrete dots of brightness [HaCaT cells  and treated with HCTZ at the final concentration of 70, 140, and 370 ng/mL. After two hours, cells were irradiated with UVA (10 J/cm2) in 100 μL of PBS. After irradiation, PBS was replaced with standard culture medium and incubated at 37°C for 24 h. Cells were washed in PBS and fixed in Bouin's liquid , for 20 minutes, and then stained with Feulgen's reaction, specific for DNA detection. Briefly, cells were incubated in 1 N HCL for 22 minutes at 60°C, with the Schiff reactive for 60 minutes at room temperature and then washed in 0.05 N HCL and 5% NaHSO3. Successively, nuclei were counterstained for 30 seconds with 0.5% fast green in alcoholic solution, dehydrated in ethanol, washed in xylene, and mounted [For acute experiments, HaCaT cells were seeded on histological slide .Microscopic analyses were performed with a fluorescence microscopy  connected to a CCD camera. Ten photomicrographs were randomly taken for each sample at ×400 magnification. The measurements were made by two independent, blinded investigators, using the ImageJ 1.33 image analysis software (μm thick) were stained with hematoxylin-eosin for morphological analysis.Cells were fixed in cold 4% paraformaldehyde for 15 minutes, dehydrated in alcohol, and then paraffin embedded. Sections  in 0.1 mol/L cacodylate buffer, pH 7.4, was directly added to the cultures for 15 minutes at RT: then, the cells were scraped, harvested by centrifugation at 3000 rpm for 10 minutes, and maintained in fixative overnight. The cells were postfixed in osmium tetroxide (1%), dehydrated, and embedded in epoxy resin. Sections (70 nm thick) were examined by a JEOL 1010 transmission electron microscope  at 80 kV, after contrast staining with uranyl acetate and lead citrate.3 cells/well corresponding to 10,000 cells/cm2) were treated with doxorubicin   for 72 h. Cytotoxicity was evaluated by using the MTS reagent as previously described [Cells (5 × 10escribed .4 cell/well corresponding to 4500 cells/cm2) were suspended in 1 mL DMEM with 20% FBS and 0.3% agar solution and then laid on top of the hardened agar medium. Cells were then incubated at 37°C in a 5% CO2 atmosphere for 21 days. Subsequently, the cells were stained with nitroblue tetrazolium chloride solution and incubated overnight at 37°C. Colonies were counted by two blinded independent observers.Anchorage-independent growth assay was performed as described by Borowicz et al. . Brieflyp < 0.05 was considered significant. Statistical analysis was performed with GraphPad Prism 5 .Results are presented as mean ± SEM of at least three independent experiments. Multiple comparisons were performed using 1-way ANOVA followed by Bonferroni's post hoc test. Difference with p < 0.01, posttest for linear trend), concentration-dependently, reaching up to 50% increase at 370 ng/mL (p < 0.05) (p < 0.05) (p < 0.001) and OGG1 activity were significantly increased in cells treated with UVA and HCTZ (370 ng/mL) compared to UVA alone (p < 0.05) , compared to UVA alone ( < 0.05) . At the  < 0.05) . However < 0.05) , and bot < 0.05) .p < 0.05 at 140 ng/mL vs. UVA; p < 0.001 at 370 ng/mL vs. UVA), and their levels remained elevated also after 6 h  and further enhanced in UVA+HCTZ compared to UVA alone (p < 0.01 at 370 ng/mL vs. UVA) (γ-H2AX foci in cells treated with UVA+HCTZ compared to control cells is shown in The percentage of cells positive for vs. UVA) . The clep < 0.05; -40% after 48 h, p < 0.001; and -50% after 72 h, p < 0.001). HCTZ alone was not cytotoxic nor did it exert additive cytotoxic effects in the presence of UVA , no significant differences were observed compared to UVA alone  (p < 0.001). In response to UVA alone or to UVA+HCTZ, the expression of APE1 was induced, but the difference was not statistically significant (0.53 ± 0.05 in naive compared to 0.89 ± 0.14 in UVA-treated cells and 1.06 ± 0.13 in UVA+HCTZ cells). The double-strand breaks marker γ-H2AX was significantly increased in cells treated with UVA compared to naive (p < 0.05) and further enhanced in UVA+HCTZ compared to UVA alone (p < 0.01) . No effect of HCTZ alone was observed.Long-term treatment with UVA+HCTZ slightly increased the levels of 8-OHdG compared to UVA alone  but dras< 0.001)  and tran Figures . UVA+HCT Figures  and sign Figures  , markedly enhanced in UVA+HCTZ cells (p < 0.001 vs. UVA) and similar to that of naive cells when HCTZ was administered alone . Naive and cells treated with HCTZ alone showed similar IC50 values for doxorubicin, 0.8 ± 0.13 μM and 0.7 ± 0.11 μM, respectively.The percentage of apoptotic cells upon long-term treatment with either UVA alone (3.47%) or UVA+HCTZ (1.44%)  was loweWe used the soft agar colony formation assay, a well-known functional test based on the capability of transformed cells to grow independently of a solid surface and considered a hallmark of in vitro carcinogenesis .p < 0.001 vs. UVA, Figures Naive and HaCaT cells treated with HCTZ alone did not form any colony in anchorage-independent (soft agar) clonogenicity assay. As expected for not fully transformed cells, on UVA+HCTZ-treated cells, we were able to detect only few colonies, but compared to UVA alone, they exhibited enhanced clonogenic potential, further suggesting their early-stage oncogenic transformation. On average, HaCaT cells treated with UVA alone formed only 1 colony/well, while the number of colonies formed by UVA+HCTZ cells was markedly enhanced compared to UVA alone  or type II (singlet molecular oxygen) mechanisms and undergoes photodehalogenation, yielding a reactive form, which can damage DNA, lipids, and proteins –40. Setoγ-H2AX, a sensitive marker of double-strand breaks and photogenotoxicity [γ-H2AX foci were due to the replication process, they are more likely attributable to a direct DNA damage since we used serum-starved, confluent cells. Moreover, the presence of discrete γ-H2AX foci, rather than a pan-nuclear staining, supports a mechanism independent on repair [As early as 1 h after acute exposure, HCTZ enhances the UVA-induced formation of 8-OHdG, a major mutagenic oxidative DNA lesion , and of toxicity , 43. Doutoxicity ; althougn repair , 45.To ensure genomic integrity, 8-OHdG is quickly removed by OGG1; whether immediate or slightly delayed, the induction of OGG1 repair activity mitigates the accumulation of 8-OHdG, consistently with the high efficiency of base excision repair enzymes . OGG1, aIn a mice model defective in nucleotide excision repair, Kunisada et al.  demonstrAt the molecular level, the presence of double-strand breaks and 8-OHdG lesions coupled with a defective DNA repair activity may lead to genomic mutations; this seems particularly relevant in the basal layer of human epidermis where keratinocytes express less OGG1 compared to the superficial layer , 52. Morβ-catenin localization [In addition to genomic insults, cotreatment with UVA and HCTZ activates Wnt, one of the oncogenic pathways involved in skin tumorigenesis , and inclization , 62.ΔNp63α [β-catenin [Human preneoplastic lesions and NMSCs also express ΔNp63α , an oncoΔNp63α  that may-catenin , 69.ΔNp63α and Bcl-2. To explore whether chronic treatment with HCTZ and UVA caused the acquisition of resistance to proapoptotic insults, cells were treated with the known proapoptotic drug doxorubicin, observing a significant increase of its IC50 value. He et al. [ΔNp63α inhibited doxorubicin-induced apoptosis in HaCaT keratinocytes, independently of wild-type p53 [It is interesting to note that, after chronic treatment, the level of apoptosis was lower than that observed after acute treatment in cells treated with UVA in the presence of HCTZ, suggesting the acquisition of apoptosis resistance, also supported by the enhanced expression of e et al.  reportedtype p53  and decrtype p53  probablytype p53 .We did not provide in vivo data in support of our findings in HaCaT cells, and no in vitro model system can perfectly recapitulate the in vivo complexity of a disease in terms of immunological response and microenvironmental interactionsWe used a single cell line; however, HaCaT cells are widely used to study keratinocytes transformation upon UVA and UVB radiation  and are We used only UVA rays; despite representing the vast majority of UV received on Earth, the combination of UVA and UVB better simulates the solar radiation. However, no significant increase in the number of UVB-induced skin tumors in mice treated with HCTZ was reported .There are some limitations of our study that should be recognized:We tested HCTZ in a range of concentrations reflecting the plasma levels of doses used in clinical practice , 26; des2 that mimics a human exposure of approximately 1 hour to midsummer sun in Paris [We applied a cumulative weekly UVA dose of 20 J/cmin Paris , and it in Paris ; moreoveThe morphological and molecular features described resemble some aspects of dysplastic lesions described by Coussens and Hanahan  in a mouThe strengths of the study, making it translationally relevant, are as follows:Our results demonstrate that the combined chronic exposure to HCTZ and UVA radiation is more prophotocarcinogenic than it would be expected from UVA alone and highlight the relevance of associating drug safety data with experimental evaluations to clarify the molecular mechanisms underlining adverse drug effects. In fact, HCTZ alone does not possess any direct damaging effect but rather potentiates that of UVA, in accordance with its photosensitizing properties. This further supports the recommendation made by the regulatory agencies for an accurate photoprotection especially while taking HCTZ.


Although PHD3 has been reported to influence cancer cell metabolism and liver insulin sensitivity, relatively little is known about the effects of this highly conserved enzyme in insulin-secreting β cells in vivo. Here, we show that the deletion of PHD3 specifically in β cells (βPHD3KO) was associated with impaired glucose homeostasis in mice fed a high-fat diet. In the early stages of dietary fat excess, βPHD3KO islets energetically rewired, leading to defects in the management of pyruvate fate and a shift from glycolysis to increased fatty acid oxidation (FAO). However, under more prolonged metabolic stress, this switch to preferential FAO in βPHD3KO islets was associated with impaired glucose-stimulated ATP/ADP rises, Ca2+ fluxes, and insulin secretion. Thus, PHD3 might be a pivotal component of the β cell glucose metabolism machinery in mice by suppressing the use of fatty acids as a primary fuel source during the early phases of metabolic stress.The   The prolyl-hydroxylase domain proteins PHD1–3) encoded for by the Egl-9 homolog genes are α-ketoglutarate–dependent dioxygenases, which regulate cell function by catalyzing hydroxylation of peptidyl prolyl residues within various substrates using molecular oxygen –3 encode. There aPHD3 is unusual among the PHDs: it is transcriptionally regulated by HIF1 during hypoxia , althoug+ channels  by crossing the r strain  with anin  to induce obesity and metabolic stress .Egln3 was moderately upregulated in βPHD3CON islets . These data raise the possibility that nutrient-sensing and utilization might have been altered in βPHD3KO islets.14C glucose. Although glucose oxidation was not altered at low or high glucose in islets from βPHD3KO mice after 4 weeks HFD feeding . Analysis of mass isotopomer distribution showed no differences in 13C incorporation into aspartate, glutamate, malate, fumarate, or citrate in βPHD3CON and βPHD3KO islets on either standard chow (SC) or 4 weeks HFD , the rate-limiting step in fatty acid oxidation , 44. IndCpt1a , which inhibits CPT1 through generation of mitochondrial malonyl-CoA, thus suppressing use of fatty acids via β-oxidation  conditions  SC βPHD3CON and βPHD3KO islets (Bnip3 and Car9 were upregulated to similar levels in hypoxic (1% O2) SC βPHD3CON and βPHD3KO islets, whereas Gls did not reliably increase . It will also be interesting to understand how PHD3 activity changes in other models of metabolic stress, such as 2+ fluxes, a sensitive readout of changes in oxygen-dependent regulation  specific PHD3 inhibitors are warranted. Second, interactions between PHD3 and ACC2 are inferred from our metabolic workup and known biochemistry. Identifying hydroxylation sites and creating corresponding ACC1/ACC2 mutants is needed, but current mass spectrometry analysis is challenging due to the assignment of false positives, as mentioned above. Third, we focused our studies on 4 and 8 weeks of HFD, and it is unclear whether the switch toward increased fatty acid utilization might be maladaptive or protective in βPHD3KO mice during longer periods of HFD feeding. Fourth, HFD studies were restricted to male animals, and further studies should be extended to female animals. Although sex differences in phenotype were not observed under standard diet, we cannot exclude a sexually dimorphic effect of HFD.We acknowledge a number of limitations with the present studies. First, workup was limited to rodents, and it will be important to confirm whether or not results translate to human islets. Although our attempts at silencing PHD3 using In summary, PHD3 possesses a conserved role in gating nutrient preference toward glucose and glycolysis during both cell transformation  and metaNo data were excluded unless the cells displayed a nonphysiological state . All individual data points are reported. The measurement unit is animal or batch of islets, with experiments replicated independently. Animals and islets were randomly allocated to treatment groups to ensure that all states were represented in the different experiment arms.LoxP system. Ins1Cre mice (JAX stock 026801), with Cre recombinase knocked into the Ins1 gene locus, were crossbred to mice carrying floxed alleles for PHD3  and their controls  were used from 8 to 20 weeks of age under both standard diet and HFD conditions. No extrapancreatic recombination has been observed in Ins1Cre mice, and possession of a Cre allele is not associated with any changes in glucose homeostasis in our hands  DREADD reporter strain. Animals were maintained on a C57BL/6J background and backcrossed for at least 6 generations following rederivation into the animal facility. Lines were regularly refreshed by crossing to bought-in C57BL/6J (Charles River Laboratories). WT male CD1 mice aged 8–12 weeks (Charles River Laboratories) were used for confirmation of gene expression under hypoxic (1% O2) conditions. βPHD3CON and βPHD3KO mice were fed SC and/or an HFD containing 60% fat , with body weight checked weekly until 18–20 weeks of age. Animals were maintained in a specific pathogen–free facility, with free access to food and water.βPHD3KO mice were generated using the Cre-ur hands , 36. RecMice were fasted for 4 to 6 hours, before i.p. injection of glucose. Animals on SC received 2 g/kg body weight glucose, whereas those on HFD received a lower dose of 1 g/kg body weight. In our hands, this allows the measurement of blood glucose concentration without the need to dilute samples and decreases adverse reactions associated with profound hyperglycemia. Blood samples for glucose measurement were taken from the tail vein 0, 15, 30, 60, 90, and 120 minutes after the glucose challenge. Glucose was measured using a Contour XT glucometer (Bayer). For mice on SC, IPGTT was performed every 2 to 4 weeks, between 8 and 20 weeks of age. HFD-fed mice underwent IPGTT after 72 hours, 4 weeks, and 8 weeks of HFD feeding. OGTT was performed as for IPGTT, except that glucose was delivered using an oral gavage tube .Blood samples were collected after i.p. glucose injection (1 g/kg body weight). Serum was separated by centrifugation, before assaying using the HTRF Mouse Serum Insulin Assay kit (Cisbio). Due to NC3R limits on blood sample volumes, insulin was measured only 0, 15, and 30 minutes after glucose injection.Mice fasted for 4 to 6 hours (SC and 4 weeks HFD cohorts) or overnight (8 weeks HFD cohort) received 0.75 U/kg body weight insulin  given by i.p. injection. Blood glucose was measured 0, 15, 30, 60, 90, and 120 minutes after insulin injection.Male βPHD3CON and βPHD3KO mice fed HFD for 4 and 8 weeks were weighed and sacrificed by cervical dislocation. The following tissues were harvested and weighed immediately postmortem: visceral fat , subcutaneous fat, brown adipose tissue, liver, and muscle (quadriceps femoris). Tissue contribution to body composition was expressed as percentage of body weight.2, 37°C. For experiments under hypoxia, islets were incubated in a Don Whitely H35 Hypoxystation, allowing oxygen tension to be finely regulated at either 1% or 21%.Islets were isolated after bile duct injection with NB8 1 mg/mL collagenase (Serva) and Histopaque/Ficoll gradient separation (MilliporeSigma). Islets were cultured in RPMI medium containing 10% FCS, 100 U/mL penicillin, and 100 μg/mL streptomycin (MilliporeSigma) at 5% CO–ΔΔCt method, expressed as fold change in gene expression. The sequences of the forward and reverse primers used in the study can be found in TRIzol (Thermo Fisher Scientific) purification was used for mRNA extraction, whereas cDNA was synthesized by reverse transcription. Gene expression was detected by quantitative real-time PCR, using PowerUp SYBR Green Master Mix (Thermo Fisher Scientific), and quantification was based on the 2Pancreata were isolated, fixed in 10% formalin, and embedded in paraffin. Paraffin slides were deparaffinized and rehydrated, before antigen retrieval using citrate buffer. Sections stained for PHD3 were incubated overnight at 4°C with guinea pig anti-insulin 1:100  and rabbit anti-PHD3 1:100 , followed by washing and 2 hours incubation at room temperature with anti–guinea pig Alexa Fluor 568 1:300  and anti-rabbit Alexa Fluor 488 1:1000 . PCNA staining was carried out using rabbit anti-insulin 1:500  and mouse anti-PCNA 1:500  as primary antibodies. Secondary antibodies used were anti-rabbit Alexa Fluor 568 1:500  and anti-mouse Alexa Fluor 488 . VECTASHIELD HardSet mounting medium with DAPI (Vector Laboratories) was used to mount coverslips on the sections.Images were taken using a Zeiss LSM780 meta-confocal microscope equipped with highly sensitive GaAsP PMT detectors. Excitation was delivered at λ = 405 nm, λ = 488 nm, and λ = 561 nm for DAPI, Alexa Fluor 488, and Alexa Fluor 568, respectively. For PHD3 staining, the emitted signals were detected at λ = 410 to 472 nm, λ = 507 to 596 nm, and λ = 570 to 694 nm, for DAPI, Alexa Fluor 488, and Alexa Fluor 568, respectively. For PCNA staining, emitted signals were detected at λ = 418 to 507 nm, λ = 507 to 552 nm, and λ = 579 to 641 nm for DAPI, Alexa Fluor 488, and Alexa Fluor 568, respectively.+ staining (fluorescein-12-dUTP)/area of insulin+ staining (as described above). The α cell/β cell ratio was calculated after staining with rabbit antibodies against insulin (as described above) and glucagon . Images were captured as described above. Excitation was delivered at λ = 405 nm, λ = 488 nm, and λ = 633 nm for DAPI, fluorescein-12-dUTP/Alexa Fluor 488, and Alexa Fluor 647, respectively. Emitted signals were detected at λ = 428 to 533 nm, λ = 498 to 559 nm, and λ = 643 to 735 nm for DAPI, fluorescein-12-dUTP/Alexa Fluor 488, and Alexa Fluor 633, respectively. For β cell mass analysis, sections were incubated with rabbit anti-insulin 1:500  and mouse anti-glucagon 1:2000 , followed by washing and application of goat anti-rabbit Alexa Fluor 647 1:500  and goat anti-mouse DyLight 488 1:500 . Coverslips were mounted using VECTASHIELD HardSet with DAPI and 425 images per section captured using a Zeiss Axio Scan.Z1 automated slide scanner equipped with a 20×/0.8 NA objective. β Cell mass (%) was calculated as the area of insulin+ staining/area of the pancreas. Excitation was delivered at λ = 330 to 375 nm and λ = 590 to 650 nm for DAPI and Alexa Fluor 647, respectively. Emitted signals were detected using an Orca Flash 4.0 at λ = 430 to 470 nm and λ = 663 to 738 nm for DAPI and Alexa Fluor 647, respectively.TUNEL staining was performed using the DeadEnd Fluorometric TUNEL System (Promega), as previously described . The pro3, 0.5 Na2HPO4, 5 HEPES, 2.5 CaCl2, 1.2 MgCl2; MilliporeSigma) supplemented with 0.1% BSA at 37°C. Insulin content was extracted using acid ethanol. Insulin concentration (ng/mL) was measured using an HTRF Insulin Ultra-Sensitive Assay kit . For experiments with exogenous lipids, islets were treated with either 0.75% BSA control or 150 μM sodium palmitate dissolved in 0.75% BSA for 48 to 72 hours before the secretion assay. This concentration and timing do not induce profound lipotoxicity in our hands, allowing the study of metabolic phenotype in the absence of β cell failure.Islets (10 to 15 size matched) were stimulated with 3 mM glucose, 16.7 mM glucose, and 16.7 mM glucose plus 20 nM Exendin-4 in HEPES-bicarbonate buffer  or Fura2 , before imaging using a Crest X-Light spinning disk microscope coupled to a Nikon Ti-E base with 10 × 0.4 NA and 20 × 0.8 NA objectives. For Fluo8 imaging, excitation was delivered at λ = 458 to 482 nm using a Lumencor Spectra X light engine. Emission was captured at λ = 500 to 550 nm using a highly sensitive Photometrics Delta Evolve EM-CCD. For experiments with the ratiometric Ca2+ indicator, Fura2, excitation was delivered at λ = 340 nm and λ = 385 nm using Cairn Research Fura LEDs in widefield mode, with emitted signals detected at λ = 470 to 550 nm.Islets were loaded with the CaFor ATP/ADP imaging, islets were transduced with the ATP/ADP sensor Perceval , with glucose and drugs  being added at the indicated concentrations and time points. Fura2 and Epac2-camps traces were normalized as the ratio of 340:385 or Cerulean/Citrine, respectively. Data were presented as raw or F/Fmin where F = fluorescence at any time point and Fmin = minimum fluorescence, or R/R0 where R = fluorescence at any time point and R0 = fluorescence at 0 minutes.In all cases, HEPES-bicarbonate buffer was used  oxidation and incorporation into lipids by scintillation spectrometry, as previously described  for 24 hours . Polar fractions were separated by centrifugation, vacuum dried, and solubilized in 2% methoxyamine hydrochloric acid in pyridine (Thermo Fisher Scientific). Samples were derivatized using N-tertbutyldimethylsilyl-N-methyltrifluoroacetamide with 1% (w/v) tertbutyldimethyl-chlorosilane (both from MilliporeSigma), before analysis on an Agilent 7890B gas chromatograph mass spectrometer, equipped with a medium polar range polydimethylsiloxane GC column (DB35-MS). Mass isotopomer distributions were determined based upon spectra corrected for natural isotope abundance. Data were analyzed using MetaboliteDetector software , and dissolved in RPMI supplemented with 10% FBS, 100 U/mL penicillin, 100 μg/mL streptomycin, and 10% BSA. At 2 hours and 16 hours after incubation, 70 islets per genotype were collected in 250 μL of PBS and lysed prior to DNA quantification and freezing at –20°C. A 200 μL aliquot of D31-palmitate–labeled solution was also collected and stored at –20°C. Similarly, upon overnight incubation, the remaining islets were collected in PBS and lysed and the DNA was quantified. The leftover labeling solution was also collected and frozen at –20°C for measures of background signal.For D31-palmitate tracing, 140 islets per genotype were cultured at 5% COm/z) of M+0 and M+31 were determined by selected ion monitoring. As a marker of fatty acid oxidation, we measured the appearance of 2H2O derived from D31-palmitate in cell media using a Finnigan GasBench-II  data files were downloaded  and loaded onto the local open source University of California Santa Cruz Genome Browser  Act 1986 of the United Kingdom , and approval was granted by the University of Birmingham’s Animal Welfare and Ethical Review Body.DN, FC, KV, RBB, RLW, JPB, MH, JWJ, JC, and DJH performed experiments and analyzed data. FC, AT, EMJ, GGL, LH, and DAT ran and analyzed samples for GC-MS. IA analyzed genomic data. DJH and DAT conceived and designed the studies. DJH supervised the studies. DJH, DN, FC, and DAT wrote the paper with input from all authors.


Stem cell (SC)-basedtherapies hold the potential to revolutionizetherapeutics by enhancing the body’s natural repair processes.Currently, there are only three SC therapies with marketing authorizationwithin the European Union. To optimize outcomes, it is important tounderstand the biodistribution and behavior of transplanted SCs invivo. A variety of imaging agents have been developed to trace SCs;however, they mostly lack the ability to simultaneously monitor theSC function and biodistribution at high resolutions. Here, we reportthe synthesis and application of a nanoparticle (NP) construct consistingof a gold NP core coated with rhodamine B isothiocyanate (RITC)-dopedmesoporous silica (AuMS). The MS layer further contained a thiol-modifiedinternal surface and an amine-modified external surface for dye conjugation.Highly fluorescent AuMS of three different sizes were successfullysynthesized. The NPs were non-toxic and efficiently taken up by limbalepithelial SCs (LESCs). We further showed that we can functionalizeAuMS with a reactive oxygen species (ROS)-sensitive fluorescent dyeusing two methods, loading the probe into the mesopores, with or withoutadditional capping by a lipid bilayer, and by covalent attachmentto surface and/or mesoporous-functionalized thiol groups. All fourformulations displayed a ROS concentration-dependent increase in fluorescence.Further, in an ex vivo SC transplantation model, a combination ofoptical coherence tomography and fluorescence microscopy was usedto synergistically identify AuMS-labeled LESC distribution at micrometerresolution. Our AuMS constructs allow for multimodal imaging and simultaneousROS sensing of SCs and represent a promising tool for in vivo SC tracing. The lack of knowledge of in vivo SC fateafter transplantation represents a significant bottleneck in theirclinical translation. Such knowledge would enable the optimizationof SC processing and transplantation strategies, in turn enablingfaster clinical translation of SC therapies with a higher successrate. Several methods have been reported that enable SC tracing aftertransplantation.3 For example, the transductionof SCs to express fluorescent proteins is a popular method; however,this is largely unsuitable for in vivo applications due to the necessityof fluorescence imaging techniques. Additionally, the genetic modificationof SCs can lead to harmful off-target effects and the produced fluorescentproteins could trigger an immunogenic response.Stemcells (SCs) have immense therapeutic potential due to theirinherent ability to differentiate into different cell types and theircapacity for self-renewal. As such, SCs offer potential new treatmentoptions for many prevalent chronic and degenerative disorders suchas rheumatoid arthritis or Parkinson’s disease.5 Forexample, in metal NPs such as gold and silver, strong optical propertiesare observed by the surface plasmon resonance (SPR) effect which canbe adjusted with morphological manipulation.6 Gold NPs (AuNPs), in particular, show localized SPR frequency, X-rayattenuation, conductivity, and biocompatibility as a function of particlesize.8 Therefore, AuNPs can be imaged by a multitudeof techniques such as optical coherence tomography (OCT), surface-enhancedRaman spectroscopy (SERS), computed tomography (CT), photoacoustic(PA) imaging, and X-ray imaging.10Nanoparticles(NPs) have attracted attention as tracing agentsdue to the limitless variability of their physical and chemical propertiesarising from quantum effects at the nanoscale.11 For example, coating AuNPs with mesoporous silicaenhances the biocompatibility while extending the functionalizationpossibilities for theranostic applications.12 Additionally, by incorporation of fluorescent, optical, or responsiveprobes into mesoporous silica, the imaging capabilities can be expanded,creating composite materials that can be detected using multiple imagingmodalities.13 Multimodal NPs can overcomethe limitations of individual imaging modalities such as low penetrationdepth or low resolution by synergetic image reconstruction.16Bare AuNPs mayalso be coated with polymers and proteins to increasetheir biocompatibility, cellular uptake, and functionality.17 Generally, across cell types, high ROS levelsare associated with decreased proliferation and apoptosis.18 In SCs, high ROS levels have also been shownto be inversely proportional to SC potency, for example, a study donein corneal epithelial SCs has shown that more efficient ROS homeostasiscan be correlated with a higher cell potency.20Although the use of NPs for SC tracing have provided importantinformation on SC biodistribution, they have not provided any functionalinformation of the transplanted SCs. Such information is vital toimprove our understanding of how SCs mediate tissue regeneration,such as whether SCs migrate and how viable they remain throughoutthe tissue regeneration process. One way of obtaining more informationabout the SC function is by monitoring the redox potential by detectinglevels of reactive oxygen species (ROS) or antioxidants. ROS are knownto play a significant role in many cell signaling pathways.2O2, HO•, and ROO• and canthus act as a general oxidative stress marker to monitor the SC redoxpotential.23 The DCF probe was incorporated in the AuMS usingfour different strategies. For the first construct, DCF was loadedinto the mesopores by diffusion (AuMSL–DCFL). Then, DCF release after an intracellular uptake of the NP wouldallow intracellular ROS detection. However, because no gating systemis used, early release of DCF could hinder the long-term applicabilityof the probe. As such, we included a second construct using a lipidcoating to physically entrap the DCF dye in the NP, slowing down DCFrelease (AuMSL–DCFL(LIP)). In the thirdstrategy, we conjugated the DCF to the thiol groups present on theMS (AuMSL–DCFc). To investigate whetherincreasing the amount of reactive groups on the NP surface would alsolead to increased sensing abilities of the NPs, we included a fourthconstruct where the AuMS particles were first postgrafted (PG) withthiol groups prior to the conjugation of DCFDA (AuMSL(PG)–DCFc).Here, we have developed multimodal NPs capable of being detectedby OCT and fluorescence microscopy that can simultaneously be usedto image ROS levels in SCs. These multimodal imaging probes consistof 60 nm AuNP coated with mesoporous silica (AuMS). Using the co-condensationapproach, the mesoporous silica network was doped with a red fluorescentprobe rhodamine-B isothiocyanate (RITC) to allow for particle imagingand to serve as an internal standard for ROS imaging. At the sametime, the internal surface (mesopores) and the external surface werefunctionalized with thiol groups and amine groups, respectively, toallow orthogonal postfunctionalization of the AuMS. Then, for intracellularROS detection, 2′,7′-dichlorodihydrofluorescein diacetate(DCFDA) was used. DCFDA is a ROS-sensitive fluorescent probe susceptibleto a number of ROS including but not limited to H25 we chose todevelop three different AuMS sizes.Given that the NP size influencesthe cell uptake and biocompatibilityand this is cell type-dependent,2O2. They were further tested exvivo in the cornea to demonstrate they can be visualized using OCTand fluorescent microscopy, which are modalities extensively usedin clinical ophthalmology.28 Finally, AuMS were tested ina model of limbal epithelial SC (LESC) transplantation, cultivatedautologous LESC transplantation (LSCT). LSCT is a SC therapy thatuses LESCs to regenerate the corneal epithelium in patients with LESCdeficiency (LSCD) because it has proven efficacy and the cornea iseasily accessible for light imaging due to its unique refractive propertiesas an avascular tissue.29 Further, ourAuMS–DCF constructs are especially applicable for LESC monitoringas it has been shown that H2O2 levels are importantin maintaining LESC health and potency.30 In this work,we show that multimodal NPs based on gold, mesoporous silica, andDCF are promising biodistribution and redox level tracking probesusing OCT and fluorescence microscopy and can effectively be appliedas LESC monitoring tools in a model of LSCT.The developed AuMS weretested in vitro in limbal epithelial SCsimmortalized with human telomerase reverse transcriptase (h-TERTs)for their biocompatibility and ability to detect ROS radicals, inthis case H2–1 . HAuCl4·3H2O (chloroauric acid), tetraethylorthosilicate (TEOS), 3-aminopropyl triethoxysilane (APTES), triethoxyphensylsilane(PTES), 3-mercaptopropyl triethylsilane (MPTES), cetyltrimethylammoniumbromide (CTAB), hydrochloric acid , ammonium nitrate (NH4NO3), RITC, hydroquinone, 2′,7′-dichlorofluoresceindiacetate (DCFDA), 1,2-dioleoyl-sn-glycero-3-phosphocholine(DOPC), 25% NH3, phenazine methosulfate (PMS), calciumchloride dihydrate (CaCl2·2H2O), triiodothyronine,cholera toxin, and collagenase were purchased from Sigma-Aldrich.ATTO 647-maleimide, ATTO 488-maleimide, and ATTO-MB2 maleimide werepurchased from ATTO-TEC GmbH. Absolute ethanol, sodium citrate dihydrate(citric acid), nitric acid 60% (HNO3), paraformaldehyde(PFA), Triton X-100, bovine serum albumin (BSA), gold standard solutionARISTAR for inductively coupled plasma mass spectroscopy (ICP–MS)(10 mg/L), ruthenium standard solution ARISTAR for ICP–MS (1g/L), and hydrogen peroxide 30% (H2O2) werepurchased from VWR. Gibco keratinocyte-SFM with l-glutamine,Gibco Dulbecco’s modified Eagle medium (DMEM)/F12 HEPES nophenol red, DMEM, F12 nutrient mixture, fetal bovine serum (FBS), l-glutamine, BSA, 0.05% trypsin-0.01% EDTA, penicillin streptomycin(1000 μg/mL), CM-H2DCFDA (CM-DCFDA), Slide-A-Lyzer MINI DialysisDevice, 10K MWCO, 0.5 mL, Lab-Tek II Chamber Slide System, F96 MicroWellBlack Polystyrene Plate, Hoechst 33342, trihydrochloride, trihydrate(10 mg/mL), CellMask deep red plasma membrane stain, Gibco amphotericinB, phalloidin Alexa Fluor 647, and goat anti-mouse-Alexa Fluor 488were purchased from Thermo Fisher Scientific. Adenine and hydrocortisonewere purchased from Merck Millipore. Insulin was purchased from LillyMedical . Epidermal growth factor (EGF) was purchased fromAMSBIO (The Netherlands). Accutase was purchased from STEMCELL Technologies.CellTiter 96 AQueous MTS Reagent Powder was purchased from Promega(the Netherlands). The Tissue-Tek O.C.T. compound was purchased fromSakura (USA). Anti-human nuclear antigen [235-1] was purchased fromAbcam.All water was purifiedby the Milli-Q system used with a conductivityof 18.2 MΩ cm2.132 First, 22nm seeds were synthesized by adding a citric acid solution (3 mL)to chloroauric acid  at 100 °C under reflux. Thereaction was continued for 15 min until a deep ruby red color wasachieved. To grow 60 nm AuNPs, the number of seeds in solution wasfirst found by UV–vis spectroscopy. The concentration of AuNPswas found by a method reported previously; the absorbance at 450 wasdivided by the extinction coefficient of 18 nm AuNP (ε450 =3.87 × 108), the total number of moles could thenbe calculated with the consideration of the total volume.33 Then, 9.5 × 1011 AuNP seedswere added to chloroauric acid solution (2.5 × 10–5 M) under rapid stirring. Immediately afterward, citric acid  and hydroquinone  were addedunder continuous stirring, and the solution was left to react for1 h. To change the surface ligand, CTAB  was added tothe solution and left to stir at room temperature overnight. The nextday, particles were collected by centrifugation, washed, and redispersedin H2O (100 mL) to remove excess CTAB. The size of CTAB-stabilized60 nm AuNPs was characterized by dynamic light scattering (DLS)  and UV–vis spectroscopy (Agilent) in order tocalculate the total moles of 60 nm AuNPs by the same method as inthe seeds .33 The number of moles of AuNPs wascalculated so that a standardized amount was always used in the silica-coatingstep. The produced CTAB-stabilized 60 nm AuNPs could be stored at4 °C for up to 1 month.AuNPs of 60 nm size were synthesized byadapting previously reported methods.13 First, 6.5 ×10–6 mol of CTAB-stabilized 60 nm AuNPs, as determinedby UV–vis and DLS, was concentrated by centrifugation  and redispersed in 5 mL of H2O with 5 min of sonication to separate the agglomerates. Then, CTAB  was dissolved in a mix of 75 mL of absoluteethanol and 170 mL of H2O and stirred at 35 °C. Oncethe solution was transparent, NH3 was added and stirred for 5 min. Then, the concentrated AuNPs wereadded and left to stir for a further 5 min. To coat AuNPs with mesoporoussilica in different thicknesses, the molar ratio of the AuNP/silicaprecursor was varied; AuMSS = 1:21, AuMSM =1:25, and AuMSL = 1:29. The conjugation of RITC and APTES(RITC–APTES) was carried out through the adaption of a previouslypublished procedure.34 Briefly, RITC (5mg) was reacted with 44 μL of APTES (molar ratio RITC–APTES= 1:10) in 1 mL of ethanol and stirred overnight in the dark. TheRITC–APTES, MPTES, and APTES ratios were kept constant at 0.4,4 and 10 mol % of total silica precursor. For example, for AuMSS, a mixture of MPTES  and TEOS were added dropwise and the temperaturewas increased to 60 °C. After 20 min, TEOS was added  followed by RITC–APTES  in 2 equal increments 3 min apart, and the mixture wasleft to stir for another 30 min. Then, for −NH2 surfacefunctionalization, a mixture of TEOS and APTES  were added to the mixtureand left to stir overnight. Additionally, non-functionalized controlAuMS were formed by the one-step dropwise addition of 175 μLof TEOS directly to the AuNP solutions followed by overnight stirring.The particles were collected by centrifugation  and washed twice with ethanol. For CTAB removal, the ion-exchangemethod we reported without acid extraction was followed.6 AuMS were stored in ethanol at −20 °C.AuMS were briefly sonicated to thoroughly dispersed before use.The synthesis of AuNPs coated withmesoporous silica (AuMS) was conducted following a reported protocolwith modification.2.22O at a concentration of 0.3 μg/mL. Optical extinctionspectra of AuNP and AuMS were recorded using a Cary 60 UV–visspectrometer (Agilent) at a particle concentration of 100 μg/mL.To confirm thiol mesopore functionalization and subsequent dual fluorescentproperties, AuMSs were labeled with the fluorescent dyes ATTO-488N,ATTO-647N, and ATTO-MB2. Per reaction, 0.5 μL of ATTO dye solution(5 mg mL–1 in DMF) were used to label per 1 mg ofAuMS. Coupling reactions of the dyes with AuMS were performed in absoluteethanol during overnight stirring. AuMS were collected by centrifugation and washed three times with ethanol.To assess the RITC concentration of AuMS, a RITC standard curve from0 to 10 μM was measured followed by AuMS particles at concentrationsof 1 mg/mL. Fluorescence quantifications were performed using a CLARIOstarspectrophotometer equipped with MARS data analysis software . Fluorescence spectra were read on a Cary Eclipse fluorescencespectrometer (Agilent). The fluorescent signal for RITC was detectedat λex = 570 nm and λem = 595 nm,ATTO-488 at λex = 488 nm and λem = 521 nm, ATTO-647 at λex = 647 nm and λem = 667 nm, and ATTO-MB2 at λex = 668 nmand λem = 686 nm.Morphologicalcharacterization was performed by transmission electron microscopy(TEM) using a FEI Tecnai electron microscope. For imaging, AuMS suspensions were spotted on a 200 mesh carbon grid andimaged after air drying at RT overnight. For AuMS size analysis, theparticle analysis function on ImageJ was used. Hydrodynamic size andelectrokinetic potential  were measuredusing the Malvern Zetasizer Nano  at 25 °Cat an angle of 90°. For the analysis, AuNP and AuMS were suspendedin H2.335 For medium preparation, keratinocyte-SFM(with l-glutamine) was supplemented to achieve 0.1 mg/mLpenicillin streptomycin, 0.4 mM CaCl2, 0.2 ng/mL EGF, and25 μg/mL bovine pituitary extract.h-TERT cells were obtainedas a gift from the lab of Prof. D. Aberdam .Limbal h-TERT were developed by Rheinwald et al. and cultured as previouslydescribed.3T3-J2 cells were purchasedfrom Kerafast . 3T3-J2 cells are a subclone ofa mouse embryonic fibroblast line. For 3T3-J2 culture, DMEM was supplementedto achieve 10% FBS. 3T3-J2 cells were used as a feeder layer for thegrowth of primary LESC cultures and were irradiated before use. First,3T3-J2 cells were detached with 0.05% trypsin 0.01% EDTA, resuspendedin media, and counted. For irradiation, the cells were dispersed ina 50 mL tube containing a minimum of 20 mL of supplemented DMEM witha maximum of 20 million cells. The cells were irradiated using a MU15Firradiator  operated at a maximum of 60.00Gy, 225 kV, and 10 mA, with the dose measured using a PTW Unidos dosemeter.Irradiated 3T3-J2 cells were either used immediately or frozen at1 million cells/mL in 50% supplemented DMEM, 40% FBS, and 10% DMSO.l-glutamine,0.4 μg/mL hydrocortisone, 1.36 ng/mL triiodothyronine, 8.47ng/mL cholera toxin, 10 ng/mL EGF, and 5 μg/mL insulin. Themedium was filtered using a bottle top vacuum 0.2 μm PES filtrationsystem (VWR) before use. For LESC cultures, 3T3-J2 cells were firstseeded either from culture at a density of 40,000 cells/cm2 or after thawing at 60,000 cells/cm2. They were leftto adhere for 3 h. Primary LESCs were isolated from the limbus ofthe cornea of an 80 year old male donor. Consent was obtained fromthe donors next of kin and consent forms were issued according tothe guidelines of the CNT . LESCs wereseeded on the feeder layer at 30,000 cells/cm2. All celltypes were cultured in a 5% CO2 incubator at 37 °C.Culture medium was changed every 2 to 3 days.For LESC primary cultures, DMEM medium was supplemented to achieve30% DMEM: F-12, 10% FBS, 25 μg/mL adenine, 4 mM For imaging andMTS/ROS assays, Gibco DMEM/F12 HEPES no phenolred was used instead of keratinocyte-SFM and supplemented by the samemethod as for h-TERT cells.2.42 incubator at 37 °C.Absorbance was read at 490 nm. The average absorbance of the controlwells were set to 1, and metabolic activity (MTS) was calculated aspercentage cell viability relative to this number. For the ROS assay,99 μL of imaging medium and 1 μL of freshly prepared DCFDA were added and the plate wasincubated for 30 min at 37 °C in the dark. The fluorescence wasread at λex = 488 nm and λem = 535nm. The average of the control was set to 100% and fluorescence wascalculated as a percentage of this value.MTS and ROS assays were performed to assesscell metabolism andtoxicity after AuMS labeling. H-TERT cells were exposed to S, M, andL-AuMS for 24 h in concentrations 0–200 μg/mL at 60–80%confluency in triplicate. 15 cell-only control wells were includedper 96 well plate. For both assays, the medium was aspirated and thecells were washed twice with PBS before adding assay reagents. Forthe MTS assay, 80 μL of fresh imaging medium as well as 20 μLof MTS/PMS solution  was added to each welland incubated for 3 h in a 5% CO2.5S, AuMSM, or AuMSL at a concentration of 100 μg/mL in triplicate. Cells wereseeded in 12-well plates and AuMS were added upon 70–80% confluency.Flow cytometry was performed 5 and 24 h after AuMS exposure. All cellsuspensions were counted and the concentration was noted for laterICP–MS. Flow cytometry was also used to assess intracellularretention of AuMS. Cells were seeded in 6-well plates. H-TERTs wereexposed to AuMSS, AuMSM, and AuMSL after seeding for 24 h. Flow cytometry was performed at 5 h, 2 days,4 days, 7 days, and 14 days after AuMS labeling. For the timepointat 14 day, cells were passaged at 7 days. To prepare all samples,cells were washed with PBS dissociated with Accutase and redispersedin 200 μL of PBS. Flow cytometry was carried out using a BDAccuri C6 flow cytometer. For each measurement, 10,000 cells werecollected. FlowJo  was used for data analysis.Flowcytometry and ICP–MS were performed to quantitatively assessthe cellular uptake of different sized AuMS. For all analysis, H-TERTswere exposed to AuMS3 60%) was added to each labeledcell suspension for a v/v; 50/950 μL, cell suspension/aqua regia.The samples were disintegrated overnight at 40 °C using an ultrasonicbath and further homogenized by microwaving until the solutions were transparent and free of particulates. Next,freshly prepared 1% HCl was functionalized with 20 ppb of rutheniumto form the matrix solution. All the homogenized cell samples werediluted 1:10 in the as-prepared matrix .Additionally, a gold standard curve ranging from 1 to 100 μg/Lwas made by the dilution of a gold stock solution in the as-preparedmatrix. ICP–MS was measured using a iCAP RQ ICP–MS (ThermoScientific). With a distinct mass at 197, gold did not interfere withother ions. Gold content in 10,000 cells was calculated with accountof the concentration of each cell suspension and normalized to cellswithout added AuMS.For ICP–MS analysis of cell uptake, freshly prepared aquaregia (HCl 30% and HNOL (100 μg/mL).After 24 h, AuMS exposed cells were directly fixed with 4% PFA inPBS for 15 min and then washed twice with PBS. The cells were permeabilizedwith 0.1% Triton X-100 for 5 min and washed twice with PBS. The cellswere stained against actin  and DNA . The samples were mountedwith Mowiol medium and stored at 4 °C until use. Epifluorescenceimages were taken using an 60× oil objective fluorescence microscope(Nikon Eclipse TI-E). Confocal microscopy images were taken with aLeica TCS SP8 STED confocal microscope, equipped with a white lightlaser tuned to excitation wavelengths of 390 nm (DAPI), 561 nm (AuMSL), and 647 nm . Acquired images were processedand orthogonal sectioning was done with FIJI.36For fluorescence microscopy experiments,h-TERTs were seeded onchamber coverslips and exposed to AuMS2.6L werefunctionalized with DCFDA dye by four different approaches.These were AuMSL–DCFL, AuMSL–DCFL(LIP), AuMSL–DCFc, and AuMSL(PG)–DCFc. In AuMSL–DCFL, DCFDA was loaded into the mesopores andin AuMSL–DCFL(LIP), a lipid bilayer wasadded around the surface of AuMSL–DCFL. To form the lipid solution for creating a lipid bilayer, a DOPClipid solution was prepared by dissolving DOPC (2.5 mg) in chloroform(4 mL) and evaporating and drying overnight (40 °C) to createa lipid film. The film was then dissolved in an ethanol/water mixture to obtain a 2.5 mg/mL lipid solution. For AuMSL–DCFc, CM–DCFDA was conjugated to thiolgroups in the mesopores and in AuMSL(PG)–DCFc, AuMSL was initially postgrafted with thiol groupsusing MPTES then functionalized with CM–DCFDA. For postgrafting,AuMSL  in ethanol was stirred underreflux at 80 °C for 20 min. MPTES was added dropwise to the colloid and reaction was continued for 5h. AuMSL(PG) were collected by centrifugation and washedthree times with ethanol. To confirm postgrafting, the surface potentialthen fluorescence readout of AuMSL(PG) after ATTO647-malconjugation was analyzed and compared to AuMSL. The methodof conjugation and measurement was the same as in the “AuMSL–DCFL) were obtained by stirringAuMSL  overnight with DCFDA in ethanol. Loaded AuMSL were split into twovials  and washed twice with ethanol undercentrifugation . For AuMSL–DCFL(LIP), AuMSL–DCFL were dispersed in 50 μL of DOPC lipid solution and incubatedfor 10 min before adding 350 μL of water to induce lipid bilayerformation. Conjugated particles were obtained by stirring 500 μgof AuMSL and AuMSL(PG) in CM–DCFDA (50μg/mL) overnight for the formation of AuMSL–DCFc and AuMSL(PG)DCFc, respectively. Theparticles were purified by centrifugation  and washed twice with ethanol. To confirm functionalization,the surface potential of AuMSL–DCFL,AuMSL–DCFL(LIP), AuMSL–DCFc, and AuMSL(PG)–DCFc was analyzedand compared with AuMSL. The method of measurement wasthe same as in the “Then AuMS were functionalized with DCFDA. First, loaded particles in a black bottom 96 well plate in triplicate, fluorescenceat λex 495 and λem 520 nm in eachwell was read from above immediately and after 120 min.To validate DCFDA functionalization,the fluorescence of NPs afterincubation with the ROS molecule HL–DCFL and AuMSL–DCFL(LIP), 100 μgof particles were suspended in 500 μL of imaging media supplementedwith 20 μM of H2O2. The solvated particleswere placed in a mini dialysis device, which capped a UV cuvette filledwith the supplemented imaging media and equipped with a stirring flea.The cuvette was closed with parafilm to prevent evaporation. The fluorescenceover time at 37 °C of the media in the cuvette was read every5 min for 6000 min. Fluorescence spectrums were read on a Cary Eclipsefluorescence spectrometer as in “To determinethe DCFDA release profiles of AuMS2.72O2-inducedcells was analyzed by a plate reader and flow cytometry analysis.For the plate reader and flow cytometry analysis, cells were seededin a 96 glass bottom and 12 well plate, respectively. H-TERT cellsat 60% confluence were incubated with AuMSL–DCFL, AuMSL–DCFL(LIP), AuMSL–DCFc, and AuMSL(PG)–DCFc at 100 μg/mL in triplicate for 24 h. For plate readeranalysis, 16 control wells were included; 8 of cells and 8 of cellsand AuMSL. After 24 h, the medium was aspirated, cellswere washed twice with PBS and refreshed with imaging medium. H2O2 was added in increasing concentrations of 0,50, 100, and 200 μM. The fluorescence of DCFDA (λex = 495 and λem = 520 nm) and RITC (λex = 555 nm and λem = 580 nm) of each wellwas read before and immediately after adding H2O2 and at 2 min intervals for 120 min at 37 °C. For flow cytometryanalysis, 6 control wells were included; 3 of cells and 3 of cellsand AuMSL. After 24 h, 100 μM H2O2 was added and incubated for 120 min, then the medium wasaspirated and cells were washed with PBS, dissociated with Accutase,and redispersed in 200 μL of PBS. The fluorescence of DCFDA(λex = 495 and λem = 520 nm) andRITC (λex = 555 nm and λem = 595nm) was read. Flow cytometry was carried out using a BD Accuri C6flow cytometer. For each measurement, 10,000 cells were collected.FlowJo  was used for data analysis.The abilityof the NP types to sense in ROS in H2O2 concentration on h-TERT viability was analyzed by an MTSassay. H-TERT cells were seeded in a clear 96 well plate and allowedone to reach 80% confluence. For each H2O2 concentration;5, 10, 20, 50, 100, 200, 500, and 1000 μM four replicates wereincluded as well as 8 control wells (0 μM H2O2). The MTS reagent was added 1 h after H2O2 was added. The MTS assay was performed the same as in the“To relatefluorescence values to h-TERT health, the effect of H2) wasused for image acquisition. Excitation was set to 390 nm (Hoescht),488 nm (DCF), 561 nm (AuMS), and 647 nm (CellMask). Fluorescent imageswere taken before H2O2 (100 μM) additionand every 5 min for 60 min after an incubation period of 15 min.For live imaging, cells wereincubated with 100 μg/mL ofeach NP type for 24 h. The medium was aspirated, washed twice withPBS, and membrane stained with CellMask deep red plasma stain (5 μg/mL)for 10 min. The medium was aspirated again, washed with PBS, and thenuclei were stained with Hoescht (1 μg/mL) for 10 min, and thenwashed twice with PBS. For imaging, cells in each well were incubatedwith 100 μL of imaging medium. An automated inverted fluorescencemicroscope (Nikon Ti-E), equipped with a Lumencor Spectra X lightsource, Photometrics Prime 95B sCMOS camera, an MCL NANO Z500-N TIz-stage, and a Okolab incubator  was performed, after which cells were thresholdedbased on the CellMask signal and segmented using “separateobjects”. To prevent detection of cell remnants, cells wereonly included for analysis of DCHF signal intensity if the cells containeda single nucleus, which was thresholded separately based on DAPI.Cells touching the border of the frame were excluded from analysis.Subsequently, mean DCHF signal intensity was measured in the individualcells per image for each time point. To track DCFH signal intensityover time, the 2D tracking module was used to consequently measurethe same individual cells.2.8L (100 μg/mL) were added at day 3 afterseeding. At 7 days postconfluence, LESC were ready for transplantation.Before transplantation the corneolimbal buttons were defrosted at4 °C and prepped for culture. Corneoscleral buttons were seton a support made from the bottom of a 50 mL Falcon tube to retainthe curvature of the cornea. Corneolimbal buttons were cultured inKM+ media with 0.25 μg/mL amphotericin B, where media coveredthe limbus but the central cornea area was left exposed to air. LESCswere treated with 0.5 mg/mL collagenase to release the monolayer fortransplantation. Handling carefully, the monolayer was transplantedonto the corneolimbal button and cultured for 3 days. The media werechanged every day and the button was fixed with 4% PFA.Rabbit eyes were obtained from an abattoir.The corneal epitheliumand limbal epithelium were removed by dissection. Briefly, the cornealepithelium was removed by scraping with a spatula. A circumferentialincision was made 2 mm anterior and posterior to the limbus and thelimbus was removed with scissors. The corneolimbal tissue was freshfrozen in liquid nitrogen and stored at −80 °C. HumanLESC were cultured according to the standard culture protocol 2.4. To lThefixed corneoscleral buttons were placed on a mount made from the Falcontube support acting as a mold with 2% agar in order to keep cornealcurvature. The corneobuttons were imaged with slit lamp OCT . The OCT images were obtained using a 1310 nm SLD lightsource at a scan depth of 7 mm and a speed of 200 Hz. For sectioning,corneal buttons were dehydrated in a sucrose gradient prior to freezingin optical cutting medium. An ultramicrotome (Leica EM UC7) was usedto cut corneoscleral button sections to 14 μm in thickness.The sections were permeabilized and blocked simultaneously using 0.1%Triton X-100 and 5% goat serum, respectively, for 2 h at room temperature.Sections were washed twice with PBS then stained with anti-human nuclearantigen . The sections were again washedtwice with PBS then stained against actin using phalloidin-Alexa Fluor647 (1:200) and against human nuclear antigen using goat anti-mouse-AlexaFluor 488 (1:500) simultaneously for 2 h at room temperature. Thesections were washed twice with PBS and finally stained against DNAusing DAPI . After washing twice with PBS,fluorescent images were taken using an inverted fluorescence microscope(Nikon Ti-E). 3D z-stacks were taken using a crestOptics X-Light V2spinning disk unit with a pinhole size of 40 μm.2.9k). Results were consideredstatistically significant at p < 0.05.Results are expressedas a mean ± SD (standard deviation). Statistical analysis wasperformed using GraphPad PRISM . One way andtwo way ANOVAs were used for comparison among groups. An exponentialdecay model for non-linear regression was used for the determinationof the rate constant  and surface (−NH2).Fluorescently doped mesoporous silica-coated AuNPs(AuMS) containinga thiol-functionalized core and amine-functionalized surface weresynthesized and denoted as AuMS 1. First,S1). The average sizes determined from TEM analysisof 30 NPs were 62.5 ± 6.4 nm, 155 ± 11.4 nm, 201 ±13.5 nm, and 243 ± 8.8 nm and denoted AuNP and small (S), medium(M), and large(L) AuMS, respectively (p < 0.0001). Amine externalsurface functionalization was confirmed by zeta potential analysis,where AuMS–NH2 gave highly positive values thatsignificantly increased with respect to particle size; 22.8 ±5.8 mV, 33.6 ± 5.2 mV, and 38.2 ± 5.1 mV (p < 0.0001). In contrast, control particles prepared with MS coatingwithout thiol or amine modification resulted in a negative zeta potential;−9.51 ± 3.61 mV (spr) was observed and decreasein the absorbance coefficient in comparison to bare AuNPs (p = 0.0042) (S2). RITC concentration wasfound to be 2.2 ± 0.1 μM, 4.9 ± 0.7 μM, and8.6 ± 1.2 μM in AuMSS, AuMSM, andAuMSL at 1 mg/mL, respectively. Thiol mesopore functionalizationwas characterized by the conjugation of a maleimide-modified fluorescentdye ATTO-488 followed by fluorescence spectroscopy .The AuMS were round in morphology with a uniformmesoporous structure1a and S1ectively 1b. AuMS  3.61 mV 1c. Therere AuNPs 2a. A sil 0.0042) 2b and S2troscopy 2c. This spr) after coatingwas observed. Additionally, decreased absorption was observed withincreasing AuMS silica thickness; this is likely a result of decreasingAuMS particle number when particle solutions with increasing MS thicknessare measured by weight concentration (μg/mL). Furthermore, here,we immobilized the fluorescent dye RITC in the silica matrix via pre-conjugationwith APTES.38 Loading light-responsiveprobes into the mesopores is a commonly used strategy; however, dueto reduced signal caused by dye leakage over time and subsequent possibletoxicity, it has become desirable to permanently encapsulate probesin the silica matrix with techniques such as postgrafting or co-condensation.40 By this method, fluorescent probes experience a variety of opticalenhancements such as reduced photobleaching, minimized solvatochromicshift, and increased fluorescent efficiency relative to free dye insolution.38 Choosing fluorescent probesthat avoid wavelength regions of strong cellular auto-fluorescenceis important for tracing the trajectory of SC easily with high distinguishability.41In conclusion, we showed thatwe could coat bare AuNP with mesoporoussilica. As a result of the refractive index of the silica coating,a small shift in absorbance maxima allows for site-specific post-functionalization of the particles.This approach has been mostly used for increasing loading efficiencyof cargo and attaching targeting ligands or pore closing functionalitiesto the surface.44 There are limited reportsof the utilization of the exposed functional groups in the mesoporesfor dye conjugation and, to the best of our knowledge, none that exploitthe potential of this for sensing applications.47 Additionally,due to the versatility of functionalization at the thiol group, manydifferent dyes and sensing molecules can be used toward interestingapplications such as cell barcoding.48 Inessence, silica-coated AuNP for tailored multimodal in vivo imagingusing OCT and fluorescence imaging were successfully synthesized.Here, we attempted to incorporate 0.4mol % RITC–APTES;a substantial increase from previous reports using between 0.002 and0.05 mol %.3.235 To evaluate the influence of AuMS of different sizes withoutencapsulated DCF, on cell metabolism and ROS levels, MTS, and ROSassays were conducted, respectively .h-TERTs resemble LESCs in behavior and morphology.ectively 3a,b. Fore in MTS 3a or ince in MTS 3b. AdditS, AuMSM, andAuMSL for 5 and 24 h at a dose of 100 μg/mL and NPcell uptake analyzed by a fluorescent peak shift (flow cytometry)or internalized gold content (ICP–MS). A time dependence onthe degree of AuMS internalization was observed, where at 5 h, h-TERTexposure to larger AuMS (AuMSM and AuMSL) showedhigher fluorescence , and AuMS size(p = 0.029) on cell internalization was observed.At 5 h, the uptake of AuMSL was significantly more thanAuMSS (p = 0.002), while at 24 h the uptakeof AuMSS was significantly more than that of AuMSM (p = 0.032).ICP–MS analysiswas conducted to assess the internalizedAuMS number by determining the gold content. A significant effectof incubation time . Merged microscopic images show NP aggregates locatedwithin the cytoplasm and in most cells, perinuclear accumulation wasobserved (S4).To visualize the internalizationand intracellular distributionof our particles, confocal and fluorescence microscopy were used.Confocal microscopy with orthogonal sectioning confirmed the presenceof AuMSs within the cytoplasm in the nuclear plane . Adherent cells were incubated with AuMSS, AuMSM, and AuMSL at 100 μg/mL for 24 h, then at5 h, 2 days, 4 days, 7 days, and 14 days after labeling cell populationswere analyzed by flow cytometry. After 5 h and 2 days, over 99% ofcells remained labeled .After 4 days, a peak shift for all three samples was observed, and>81% cells were labeled .This was reduced further after 7 days to about 50% of the cell populationand to 4% after 14 days .Additionally, quantitativeassessment of the longevity of celllabeling by AuMS50 Nevertheless, at particularlysmall sizes or high dosages, MSNs can be cytotoxic. Specifically,MSN sizes below 100 nm have led to increased ROS production and doses≥250 μg/mL have been reported to decrease viability butare mostly cytocompatible.52 It was unsurprising therefore that our AuMSs with diameters between150 and 250 nm applied at doses below 250 μg/mL were not cytotoxicto the h-TERT cell line.Here, we show that our AuMSs are non-toxic, cell-internalized,and highly fluorescent, also when internalized by cells. Mesoporoussilica NPs (MSNs) have been widely demonstrated as non-toxic celllabeling agents with some varieties (Cornell dots) receiving FDA approvalfor human clinical trials.55 In SC trackingapplications, the degree of the cell internalization of NPs is ofvital importance as it usually correlates with imaging longevity.MSN size has been shown to have an inverse relationship with celluptake, where larger NPs require more energy in clathrin-dependentinternalization mechanisms.54 At 5 h of incubationtime, we observed higher fluorescence of cell populations labeledwith AuMSM or AuMSL, suggesting faster uptakeof these sizes. At 24 h, where maximal uptake was seen, the fluorescenceintensities became similar and more homogeneous for the three conditionsof AuMS-labeled cells. Given that AuMS fluoresce as a function ofMS thickness and increasing thickness led to similar fluorescenceintensities in cells, it could be presumed that the intracellularAuMS number decreases with the size.Although MSN size has a limited impacton cytotoxicity, it hasbeen shown to have a dramatic effect on the degree and mechanism ofcell internalization.S were taken up by cells compared to AuMSM. However for AuMSL, the uptake increased againand no significant difference was observed compared to AuMSS, this is similar to the findings by Lu et al. (2009), where 280nm MSN cell internalization increased compared to 170 nm MSNs.24This effect was confirmedby ICP–MS analysis because allAuMS were synthesized from the same batch of AuNP, mass content ofgold in cells determined by ICP–MS was directly proportionalto particle number. This meant that at 24 h a significantly higheramount of AuMSFigure S5). The reduction in the signal is likelydue to cell proliferation and is in line with what has been reportedpreviously. For example, in our previous study, we observed that theMSN signal was halved after every cell passage, resulting in the lossof signal in flow cytometry after about 15 days.56 Similarly, Huang et al. demonstrated that FITC-labeledMSNs could be detected up to 7 days via flow cytometry. They arguedthat this was likely due to cell proliferation. Signal longevity wasmethod dependent; the NPs could still be observed by confocal microscopy21 days after single exposure to hMSCs.57 This was also observed in another study by Rosenholm et al., thepercentage of Dil-functionalized MSN-labeled MDA-MB-231 cells by flowcytometry was approximately halved every cell passage and could bedetected up to a 7-day period. However, the same labeled cells couldstill be detected in mice 32 days after implantation.42 Thus, the retention in vivo could potentially be much longerfor our NPs as well.Additionally, we demonstrated that invivo TERT labeling couldbe observed up to 7–14 days using flow cytometry . In the second strategy, DCFDA was also loaded into the mesoporesof AuMSL, but a supported lipid bilayer was added actingas a gatekeeper, slowing down DCFDA diffusion from the pores (L–DCFL(LIP)). In the third strategy, a thiol reactivederivative of DCFDA (CM–DCFDA) was conjugated to AuMSL via the exposed thiol groups in the pores of AuMSL (L–DCFc). In the fourth and last strategy, AuMSL were initially postgrafted with MPTES to create additionalthiol groups over the entire surface of AuMSL, prior toCM–DCFDA functionalization (L(PG)–DCFc). MPTES postgrafting of AuMSL was confirmed by zeta potentialanalysis and fluorescence upon functionalization with ATTO-647N . Surface functionalization of AuMS–DCFwas also characterized by zeta surface potential analysis . While the zeta potentials of AuMSL–DCFL and AuMSL–DCFc were not significantly distinct from unmodified AuMSL, AuMSL–DCFL(LIP) and AuMSL(PG)–DCFc were significantly more negativelycharged [p < 0.0001]. The DCFDA release profiles of AuMSL–DCFL and AuMSL–DCFL(LIP) were determined by analyzing a fluorescence responseof 100 μg of AuMSL–DCFL and AuMSL–DCFL(LIP) suspended in a membrane-boundcompartment in cell media supplemented with 20 μM H2O2. For AuMSL–DCFL and AuMSL–DCFL(LIP), 75% and 79% of maximum releasewas achieved at 2 days, respectively. We also observed that AuMSL–DCFL(LIP) had a slower release rate comparedto AuMSL–DCFL where k = 0.00064 and k = 0.00069, respectively .Because intracellular fluorescence intensitiesand gold contentof AuMSrategies 4a–d. In L–DCFs to sense ROS was thendetermined by measured fluorescence intensity upon addition of hydrogenperoxide (H2O2) in increasing concentrations;0–200 μM. H2O2 was always addedto homogeneous NP solutions, and thus the ROS concentration was equalunder all conditions. Under these conditions, all four AuMSL–DCF were able to detect ROS in a concentration-dependentmanner. However, the different functional approaches had significantlydifferent H2O2 detection abilities, where AuMSL(PG)–DCFc was the most sensitive and AuMSL–DCFL the least [p <0.0001] . In orderto relate the obtained fluorescent values to cellular viability, theresponse of h-TERT cells to increasing H2O2 concentrations(5–1000 μM) was evaluated in a MTS assay. At a H2O2 concentration of 50 μM, cell viabilitywas at 81.2 ± 9.3%, which decreased to 54.5 ± 10.2% at 200μM and further to 8.2 ± 7.2% at 1000 μM, indicatingcell death .The ROS sensing ability of the four synthesized AuMS–DCFparticles in h-TERT cells was evaluated after 24 h exposure to 100μg/mL to allow NP uptake. After 24 h, h-TERT cells were exposedto increasing Hn period 5a–d. All 120 min 5. Furtheobserved 5 and S9.L–DCF intensities,flow cytometry was conducted. h-TERT cells were incubated with allAuMSL types for 24 h and induced for intracellular ROSwith 100 μM H2O2 for 120 min  and then the conjugatedapproaches, where AuMSL(PG)–DCFc wasbrighter than AuMSL–DCFc. RITC fluorescencewas simultaneously monitored as the internal standard and was similarin all four constructs  demonstrated a lowerincrease compared to the other Au-MS constructs (L(PG)–DCFc were most effective in in vitro experiments(without cells), while AuMS with loaded DCF demonstrated the highestsensitivity to ROS upon internalization in TERT cells.To demonstrate that ROS signatures of AuMSme point 6a. By thnstructs 6b. In suL(PG)–DCFc was mostsensitive to increasing H2O2. The amount ofDCF that was loaded into or conjugated to AuMSL will likelyvary between the constructs because the incorporation modes are different.In addition, DCF dye availability and dispersity in solution (loadedDCF will diffuse out) may play a role in the observed differences.Although all four AuMS–DCF particles showed ROS sensingcapabilities, depending on the assay, we observed differences in thelevel of response of AuMS to ROS. Without cells in culture media,we observed that AuMSL–DCFL was the brightest. Other factors,such as the endocytic pathway, uptake efficiency, and intracellulardistribution affect the availability of DCFDA to ROS present in thecytoplasm and thus the observed fluorescence intensity. While we observedno difference in the cell uptake between the different AuMS–DCFunder flow cytometry . For AuMSL–DCFc and AuMSL(PG)–DCFc, the DCFis conjugated to the MSNs and remains stably bound. This can resultin the MSNs sensing ROS in a more local environment compared to AuMSL–DCFL and AuMSL–DCFL(LIP) and as such could explain the difference in intracellularbrightness’s between AuMS–DCF with different encapsulationmethods.However, when internalized by h-TERT cells, we observed that AuMSytometry 5f, the Aytometry 3e. The dFigure S10).61 In the case of AuMSL–DCFL with a lipid coating, the ROS response waslower than without a lipid coating .Our probes allow simultaneous use of the NP RITC signalas an internalstandard so that intracellular ROS levels can be accurately quantified5f and ca2O2. Here, the ROS response followed a similar trend as in cell culturemedia, where similar ROS sensitivity was observed for AuMSL(PG)–DCFc, AuMSL–DCFc,and AuMSL–DCFL and lower sensitivityfor AuMSL–DCFL(LIP) (L–DCFL(LIP) compared to AuMSL–DCFL . Our research demonstrates that AuMS can be used as ROS-responsiveagents by functionalization with DCFDA by four different approacheswhile using RITC as an internal standard. DCF-loaded MSNs appear toprovide a robust platform to assay intracellular ROS levels especiallywhen using flow cytometry or plate reader methods. Conjugation ofCM–DCFDA, especially to PG AuMS, enabled increased sensitivityto ROS and can be used to provide information on local ROS production.When combined with the desirable features, such as sensing longevityand intracellular localization, these probes can be a powerful approachto obtain data on subcellular ROS levels.62Fluorescence microscopy enabledreal-time single-cell trackingof AuMS–DCF-labeled cells induced with HCFL(LIP) 6. This c3.4d = 176 ± 9.8 nm) were tested againstAuMS with a smaller 18 nm AuNP core (d = 187 ±9.3 nm) with no significant difference in the overall diameter. BothAuMS (20 μL) were injected into the corneal stroma of ex vivoporcine eyes at a concentration of 25 μg/mL. Because we observedthat the AuNP size was the most determinating factor for providingOCT contrast , we here usedAuMSs with large 60 nm Au cores for further ex vivo studies. To investigatethe long-term labeling and multimodal SC tracking capability of AuMSconstructs, a model LSCT was performed using AuMS-labeled LESC. Atday 3, after seeding, human primary LESCs were exposed to 100 μg/mLof AuMSS for 24 h. At day 8, the AuMSS-labeledhuman LESC monolayer was transplanted to a decellularized rabbit corneoscleralbutton with the epithelium removed and cultured until day 10.To assess theability of our AuMS to be detected by OCT and determine whether AuNPsize has an impact on OCT contrast efficiency, AuMS with a 60 nm AuNPcore . We show thatAuMS are efficient OCT contrast agents and were internalized and retainedin human LESCs for upward of 2 weeks in a LSCT model.After fixation, the LSCT model was imaged by OCT. In comparison toa reference image of a rabbit eye with an intact corneal epithelium7a, the mFigure S11). While it is knownthat the OCT contrast primarily relies on agents with high light scatteringcross sections, both AuNP and MSNs have been shown to have high scatteringcross sections that are enhanced as a function of size.65 Therefore, it was important to understand the relative scatteringeffect of AuNP and MSNs for aiding the design of our AuMS constructand of novel AuNP and MSN tracing probes in general.In an ex vivo LSCT model, we used OCT and fluorescence microscopyto demonstrate the multimodal, long-term labeling capability of ourAuMS constructs. First, we compared our AuMS against AuMS with a smaller18 nm AuNP core, where we demonstrated that increasing the AuNP sizehas a critical impact on the OCT contrast, while MSN size had a negligibleeffect  were synthesized with multiple functionalization throughouttheir core structure. All sizes of AuMS were non-toxic to h-TERT cellsup to a concentration of 200 μg/mL and were efficiently takenup as demonstrated by flow cytometry, ICP–MS, and fluorescencemicroscopy. AuMS were successfully functionalized with DCFDA usingfour different approaches, all of which were capable of concentration-dependentintracellular ROS sensing and suitable for ROS quantification by internalstandard normalization using RITC. Postgrafted AuMS with conjugatedDCFDA exhibited the most sensitivity for ROS detection by single-celltracing using fluorescence microscopy.DCFDA-conjugated AuMSdemonstrate a new class of SC tracing probesenabling localized, highly sensitive intracellular ROS sensing, andquantification. The multimodal AuMS constructs were applied in a LSCTmodel demonstrating a high contrast efficiency in the clinically relevantimaging modalities; OCT and fluorescence microscopy. Synergistic tracingof LESCs in a LSCT model at single-cell resolution was realized. AlthoughLESC tracing for LSCT was the focus of this study, deep tissue SCtracking at meter scale penetration should also be possible due tothe CT and X-ray contrast capacity of AuMS. AuMS can also easily beadapted for SC tracing in other therapy models due to the adaptablefunctionalization possibilities where other therapeutic, sensing,or imaging agents can be incorporated. Therefore, it is proposed thatthis study describes a translatable proof-of-concept for single-cellin vivo SC monitoring using AuMS constructs.


Brassica oleracea L.) leaves. Seedlings fed with 30 mM GB via sub-irrigation showed effectively assimilated GB as evident by higher GB concentration. Exogenous GB did not retard leaf-growth  rather slightly promoted it. Temperature controlled freeze-thaw tests proved GB-fed plants were more freeze-tolerant as indicated by lower electrolyte leakage  and alleviating oxidative stress ) following a relatively moderate or severe freeze-thaw stress, i.e., −2.5 and −3.5 °C. Improved freezing tolerance induced by exogenous GB application may be associated with accumulation of compatible solute (proline) and antioxidant (glutathione). GB-fed leaves also had higher activity of antioxidant enzymes, catalase (CAT), ascorbate peroxidase (APX), and superoxide dismutase (SOD). These changes, together, may improve freezing tolerance through membrane protection from freeze-desiccation and alleviation of freeze-induced oxidative stress.Exogenous glycine betaine (GB) application has been reported to improve plant tolerance to various abiotic stresses, but its effect on freezing tolerance has not been well studied. We investigated the effect of exogenous GB on freezing tolerance of cabbage ( While plants are highly tolerant to sub-zero temperatures during the dormancy in winter, the sensitive tissues of plants, including buds, new leaves, and flowers, are typically vulnerable to frost during the active growth . AccordiPlants from temperate regions can improve their FT upon exposure to cold temperature through a process called cold acclimation ,9,10. FrBrassica oleracea L.) is an economically valuable horticultural crop in Asia, particularly in Korea, due to its high consumption  = 6.45 × (A532 − A600) − 0.56 × A450 [Leaves from NGB and GB-fed plants that were exposed to freeze-thaw stress at −2.5 and −3.5 °C and their corresponding UFCs were used to measure MDA concentration, as described in Reference . Briefly6 × A450 . The con2•−) and hydrogen peroxide (H2O2) distribution were visualized using stain by nitroblue tetrazolium (NBT) or 3,3′-diaminobenzidine (DAB) method, respectively, as detailed by Reference [Superoxide (Oeference . This exg) for 25 min at 4 °C. The resultant supernatants were then employed as the enzyme extract for CAT, APX, and SOD. The protein amount in the total enzyme extracts from 150 mg tissues was estimated as detailed by Reference [The activities of three antioxidant enzymes, i.e., catalase (CAT), ascorbate peroxidase (APX), and superoxide dismutase (SOD), were measured, as detailed by Reference . Brieflyeference .2O2. One unit of CAT activity was defined as the degradation of 1 μM H2O2 in 1 min at 240 nm. The reaction mixture for APX activity was composed of: 100 mM potassium phosphate buffer (pH 7.0), 0.5 mM ascorbic acid, 0.1 mM EDTA, 50 μL enzyme extract, and 0.1 mM H2O2. One unit of APX activity was defined as the conversion of ascorbic acid into monohydroascorbate in 1 min at 290 nm.For CAT activity, the reaction mixture consisted of: 100 mM potassium phosphate buffer (pH 7.0), 20 μL of enzyme extract, and 50 mM HThe reaction mixture for SOD activity included: 50 mM potassium phosphate buffer (pH 7.8), 15 mM nitro blue tetrazolium, 130 mM methionine, 20 μM riboflavin, 5 μM EDTA, and 50 μL enzyme extract. One unit of SOD activity was defined as the amount of enzyme required to reach 50% inhibition of formazan formation at 560 nm.g for 10 min, and resultant supernatant was further centrifuged for 5 min. The sample was then injected into a Spherisorb 5 um ODS column (250 mm × 4.6 mm) for HPLC coupled to 1200 series evaporative light scattering detector .Leaves excised from NGB and GB-fed plants were used to measure the GSH concentration using HPLC, as described by Reference . The fing at room temperature (~20 °C) for 20 min. Subsequently, the supernatant (500 μL) was diluted with 500 uL of distilled water, followed by mixing with 1 mL of ninhydrin and 1 mL of glacial acetic acid. The reaction mixtures were heated in a water bath at 95 °C for 1 h and then allowed to cool in an ice bath before being vigorously mixed with 4 mL of toluene. Samples were then set aside to allow separation of the non-polar and polar phase. The non-polar phase containing the chromophore was used to read absorbance at 520 nm using toluene as a blank. Proline concentration was determined from a standard curve , and concentration values were then calculated based on DW.Leaves excised from NGB and GB-fed plants were used to determine the proline concentration, as described by Reference . The fint-test (α = 0.05) to estimate any statistically significant differences between two treatments. The pair-wise differences among more than three treatments were compared via the Fisher’s LSD test (α = 0.05). Pearson correlation analysis between the measured physiological and biochemical indices was performed using R software .All biochemical analyses were independently repeated thrice, each with 3 technical replicates per treatment. The data were analyzed using Student’s 50  and then explored the hypothesis that exogenous GB application enhances FT of cabbage leaves. Our study offers evidence that exogenous GB application can improve FT of cabbage leaves by reducing membrane leakage and lowering MDA and ROS accumulation without causing any detrimental effect on leaf growth. Our data indicate that GB-induced FT may be linked to higher accumulation of compatible solutes  and increased activity of antioxidant enzymes , as well as non-enzymatic antioxidant . Future studies should analyze the effect of exogenous GB application on FT at whole-plant level. Furthermore, additional mechanistic elucidation of GB-induced FT of cabbage leaves is warranted, involving comparative gene expression analyses and monitoring other factors  contributing to FT.In the present study, we first determined the FT of Myeong-sung cabbage leaves as the LT


Background: Bisphenol A (BPA), a reprotoxic and endocrine-disrupting chemical, has been substituted by alternative bisphenols such as bisphenol F (BPF) and bisphenol S (BPS) in the plastic industry. Despite their detection in placenta and amniotic fluids, the effects of bisphenols on human placental cells have not been characterized. Our objective was to explore in vitro and to compare the toxicity of BPA to its substitutes BPF and BPS to highlight their potential risks for placenta and then pregnancy. Methods: Human placenta cells (JEG-Tox cells) were incubated with BPA, BPF, and BPS for 72 h. Cell viability, cell death, and degenerative P2X7 receptor and caspases activation, and chromatin condensation were assessed using microplate cytometry and fluorescence microscopy. Results: Incubation with BPA, BPF, or BPS was associated with P2X7 receptor activation and chromatin condensation. BPA and BPF induced more caspase-1, caspase-9, and caspase-3 activation than BPS. Only BPF enhanced caspase-8 activity. Conclusions: BPA, BPF, and BPS are all toxic to human placental cells, with the P2X7 receptor being a common key element. BPA substitution by BPF and BPS does not appear to be a safe alternative for human health, particularly for pregnant women and their fetuses. Bisphenol A diphenol) is a phenolic compound discovered in the late nineteenth century, used in the manufacturing of polycarbonates, epoxyresins, and other polymers. It is therefore found in a wide range of products such as food and beverage containers, compact discs, personal protective equipment, sport equipment, and medical equipment, leading to multiple sources of exposure for the entire population. In humans, BPA is detected in the blood and urine, but it is also found in the placenta and amniotic fluid ,2,3. BPA,2-diyldiThe placenta is a crucial organ for pregnancy, acting as an endocrine organ and being an interface between the mother and the fetus. Consequently, it plays a central role in the health of the fetus, and has a lifelong impact on their future wellbeing. Disordered placental development is the primary defect in the majority of pregnancy diseases, such as preeclampsia , fetal growth restriction, recurrent miscarriage, and stillbirth ,7,8,9,10Our aim was to compare bisphenol A toxicity to its substitutes, bisphenol F and bisphenol S, on their ability to induce in vitro cell death in human placental cells, at concentrations that can be found in the placenta, in order to highlight the potential risks for the placenta and then pregnancy.®, CellEventTM Caspase-3/7 Green Detection Reagent and Hoechst 33342 probes were obtained from ThermoFisher Scientific , while Caspase-Glo® 1 Inflammasome Assay, Caspase-Glo® 8 Assay, and Caspase-Glo® 9 Assay were from Promega . All chemicals were purchased from Sigma-Aldrich .Chemicals and reagents. Cell culture reagents: Minimum essential medium (MEM), fetal bovine serum (FBS), 2 mM glutamine, 100 U/mL of penicillin, 100 μg/mL of streptomycin, trypsin-EDTA 0.05%, and phosphate-buffered saline (PBS) were provided by Gibco , and cell culture material such as flasks and microplates by Corning . YO-PRO-1Bisphenol A, bisphenol F, and bisphenol S were dissolved in dimethylsulfoxyde (DMSO). Stock solutions were stored at −20 °C and work solutions were obtained after a 1/1000 dilution in culture medium. The final concentration of DMSO on cells was less than or equal to 0.1%.Human placental cell culture. JEG-3 cells were selected in our study for several reasons. Primary cultures of human placental cells, in addition to being difficult to obtain and maintain in culture, may have been exposed to BPA, BPF, and BPS or other chemicals during pregnancy, which would lead to bias in data analysis. Several human placental cell lines are available in international collections such as the American Type Culture Collection (ATCC). Previous studies have shown that the JEG-3 cell line provides an appropriate model to detect placental toxicity ,20. JEG-2 polystyrene flasks. Cell cultures were maintained in a cell culture incubator . When the JEG-3 cells reached subconfluency, they were detached using trypsin-EDTA and counted. The cellular suspension was diluted and seeded in 96-well microplates at a cellular density of 80,000 cells/mL (200 µL/well), then kept at 37 °C for 24 h. The cells were incubated for 72 h with BPA, BPF, and BPS in MEM supplemented with 2.5% FBS according to Olivier et al.’s protocol that describes the JEG-Tox model [The JEG-3 human trophoblast cell line was obtained from ATCC (HTB-36). Cells were cultured in minimum essential medium (MEM) supplemented with 10% fetal bovine serum (FBS), 1% L-glutamine, and 0.5% penicillin and streptomycin in 75 cmcells/mL 0 µL/wellexc = 540 nm and λem = 600 nm) using a Spark® microplate reader .Cell viability: Neutral Red assay. The Neutral Red solution at 0.4% (m/v in water) was diluted in cell culture medium to obtain a working concentration of 50 μg/mL. Neutral Red working solution was distributed in the plates for a 3 h incubation time at 37 °C. The cells were then rinsed with PBS and lysed with a solution of ethanol–water–acetic acid . After homogenization, the fluorescence signal was scanned (λ® microBCA Protein Assay Kit (Thermofisher Scientific). Protein sample buffer and reducing agent were added and the samples were boiled at 70 °C for 10 min. Then, 40 μg of these samples was loaded and electrophoretically separated on 4–12% polyacrylamide gels , before being transferred to nitrocellulose membrane  using iBlot™ 2 Gel transfer device (ThermoFisher Scientific). Blots were incubated first using a primary polyclonal rabbit anti-P2X7 at 1:1000  and then a secondary DyLight 800 Fluor-conjugated anti-rabbit antibody at 1:15,000 for 1 h (ThermoFisher Scientific). Actin was used as a housekeeping protein for the normalization of data using an anti-actin polyclonal antibody at 1:1000 . Blots were scanned using Odyssey® Imaging System . Quantitation of bands was performed using ImageJ software.P2X7 receptor expression: Western blot technique. Cell lysates were prepared by washing the cells twice in cold PBS, lysing them in RIPA buffer (Sigma-Aldrich) supplemented with protease inhibitor cocktail (Sigma-Aldrich) for 15 min on ice, and then, the obtained lysates were sonicated for 30 s. The total protein concentration was determined using the Pierce® assay. P2X7 cell death receptor activation was evaluated using the YO-PRO-1® assay [® probe only enters into cells after pore opening induced by P2X7 receptor activation, and binds to DNA, emitting fluorescence. A 1 mM YO-PRO-1® stock solution was diluted at 1/500 in PBS just before use and distributed in the wells of the microplate. After a 10 min incubation time at room temperature, the fluorescence signal was read (λex = 485 nm and λem = 531 nm) using a Spark® microplate reader.Cell death P2X7 receptor activation: YO-PRO-11® assay . YO-PRO-® Assays. Caspase-1, -8, and -9 activity was evaluated using the Caspase-Glo® 1, 8, and 9 Assay Kits, respectively. The assay was performed according to the manufacturer’s instructions. Luminescence was quantified using a Spark® microplate reader.Caspase-1, -8, and -9 activity: Caspase-GloTM Caspase-3/7 Green Detection Reagent. Caspase-3 activity was evaluated using the CellEventTM Caspase3/7 Green Detection Reagent. Cell EventTM Caspase-3/7 Green Detection reagent was diluted in PBS with 2.5% FBS to a final concentration of 8 μM. The cells were incubated with the reagent for 30 min and then rinsed with PBS. The cells were observed under fluorescence microscopy and pictures were taken .Caspase 3 activity: CellEventex = 350 nm and λem = 450 nm) using a Spark® microplate reader, and the cells were observed under fluorescence microscopy and pictures were taken .Quantitative and qualitative analysis of chromatin condensation: Hoechst 33342 assay. Chromatin condensation was evaluated using the Hoechst 33342 assay . Hoechst 33342 fluorescent probe enters and intercalates into DNA in living and apoptotic cells. The fluorescent signal is proportional to chromatin condensation, and the nuclei of living cells are blue, while the nuclei of apoptotic cells are bright ,23,24. AResults exploitation and statistical analysis. The results are expressed in percentage or fold change compared to control cells and presented as means of at least three independent experiments ± standard errors of the mean.p < 0.0001, *** p < 0.001, ** p < 0.01, and * p < 0.05 compared to control cells.Statistical analysis was performed using GraphPad Prism 8 software . A one-way analysis of variance for repeated measures followed by a Dunnett’s test with risk α set at 5% were performed. Thresholds of significance were **** We investigated JEG-Tox cell viability after incubation with bisphenols using the Neutral Red assay. Any concentration inducing a loss of cell viability greater than or equal to 30% was considered as cytotoxic (ISO 2009). Therefore, no statistical analysis was performed.No loss of cell viability was observed after 72 h with BPS up to 100 μM c. In con® assay. P2X7 receptor was significantly activated by the three tested bisphenols. BPA and BPF induced similar levels of P2X7 activation in a concentration-dependent manner (×1.26 at 25 μM and ×1.34 at 50 μM in P2X7 pore opening, reflecting P2X7 receptor activation, was assessed using the fluorescent YO-PRO-1P2X7 receptor expression was investigated after incubation with bisphenols using the Western blot technique. Only BPS enhanced P2X7 receptor expression compared to the control (×1.18 at 25 μM, ×1.33 at 50 μM, and ×1.31 at 100 μM; ® 1 inflammasome, Caspase-Glo® 8, and Caspase-Glo® 9 assays were used to quantify caspase-1, caspase-8, and caspase-9 activity, respectively, and the fluorescence CellEvent® Caspase-3/7 Green Detection Reagent was used to analyze caspase-3 activity by microscopy.The bioluminescent Caspase-GloOnly the highest concentration of BPA significantly activated caspase-1 (×1.57 at 50 μM; All tested concentrations of BPF significantly activated caspase-1 compared to the control (×1.60 and ×2.61 at 25 μM and 50 μM respectively; BPS had no effect on caspase-1 c, caspasThe three tested bisphenols condensed significantly the chromatin. BPA induced the most chromatin condensation, but only at 50 μM ×1.79; a,f. BPF The objective of the present study was to explore and compare the toxicity of BPA to its substitutes BPF and BPS in a human placental cell line to highlight their potential risks for pregnancy.In humans, BPA is detected in amniotic fluid, neonatal blood, and the placenta at levels of nanograms/mililiter ,3,25,26.The present study is the first to demonstrate that BPA, BPF, and BPS are all toxic to placental cells, but with different impacts. In particular, cell viability, P2X7 receptor activation, caspases activation, and chromatin condensation are differently affected by these three bisphenols, despite their structural homology. Previous studies comparing BPF and BPS were focused either on the quantification in the human placenta , on the BPA and BPF induced similar levels of P2X7 receptor activation, BPF being slightly more potent than BPA. BPS induced lower activation than both BPA and BPF, but is the only bisphenol that enhanced P2X7 receptor expression. The P2X7 receptor plays a key role in inflammation by NLRP3-inflammasome formation, leading to caspase-1 activation ,51. In oIn conclusion, the results of our study suggest that BPA, BPF, and BPS induce toxicity in human placental cells. Despite their structural homology, the induced pathways are different, but they all share P2X7 receptor activation as the key starting event, reported to trigger preeclampsia in clinics . BPF and