Polymeric surface coatings capable of effectively integratingdesiredfunctional molecules and ligands are attractive for fabricating bio-interfacesnecessary for various applications. Herein, we report the design ofa polymeric platform amenable to such modifications in a modular fashionthrough host–guest chemistry. Copolymers containing adamantane(Ada) moieties, diethylene glycol (DEG) units, and silyloxy groupsto provide functionalization handles, anti-biofouling character, andsurface attachment, respectively, were synthesized. These copolymerswere employed to modify silicon/glass surfaces to enable their functionalizationusing beta-cyclodextrin (βCD) containing functional moleculesand bioactive ligands. Moreover, surface functionalization could bespatially controlled using a well-established technique like microcontactprinting. Efficient and robust functionalization of polymer-coatedsurfaces was demonstrated by immobilizing a βCD-conjugated fluorescentrhodamine dye through the specific noncovalent binding between Adaand βCD units. Furthermore, biotin, mannose, and cell adhesivepeptide-modified βCD were immobilized onto the Ada-containingpolymer-coated surfaces to direct noncovalent conjugation of streptavidin,concanavalin A (ConA), and fibroblast cells, respectively. It wasdemonstrated that the mannose-functionalized coating could selectivelybind to the target lectin ConA, and the interface could be regeneratedand reused several times. Moreover, the polymeric coating was adaptablefor cell attachment and proliferation upon noncovalent modificationwith cell-adhesive peptides. One can envision that the facile synthesisof the Ada-based copolymers, mild conditions for coating surfaces,and their effective transformation to various functional interfacesin a modular fashion offers an attractive approach to engineeringfunctional interfaces for several biomedical applications. Polymeric coatings on solid surfaces offer a versatileinterface for enabling the abovementioned applications.15 Generally, one of the two methods is utilized for incorporatingfunctional molecules on a polymeric interface: covalent conjugationor immobilization of noncovalent interactions. While covalent methodsfor surface immobilization are robust,17 noncovalentmethods offer flexibility such as facile reversibility and renewability/reusabilityof functional interfaces.21 Also, in addition to the aforementioned advantages,noncovalent surface functionalization strategies offer an economicalapproach to achieve diversity in a modular fashion,22 and importantly, the approach can be engineered to be nondestructiveto the substrate and the guest molecule.23Over the past decades, functional polymericcoatings on surfaceshave become prominent due to their widespread use in biomedical sciences.26 Among reported macrocyclic host molecules, crownethers,27 cyclodextrins (CDs),29 calixarenes,30 and calixpyrroles31 are among the most widely explored ones. Inaddition to these traditional hosts, cucurbiturils (CB[n]),32 blue-box,33 pillar[n]arenes,34 andresorcinarene35 have attracted attentionin recent years. Among these macrocyclic structures, CDs have beenwidely used to fabricate polymeric self-assembled materials for biomedicalapplications due to their ready availability, low toxicity, excellentbiocompatibility, and ability to make strong complexation with a varietyof guests.41 Among CD-based host–guest pairs, adamantane (Ada) is a rigidhydrophobic molecule known to form a stable complex with βCDwith a high association in an aqueous environment.42 Therefore, the assembly of βCD and Ada has been extensivelyused in various areas such as drug and gene delivery, biomoleculeimmobilization, bio-separation, and fabrication of sensors.49 Among these applications, the construction of functional polymericinterfaces to enable reversible immobilization of biomolecules hasbeen explored in recent years.56 In surface coatings, Chen and coworkers, in a seminal contribution,fabricated Ada-containing polymer brushes to construct surfaces capableof molecular recognition.50 They fabricated Ada-containingthermo-responsive polymer brushes and showed tunable and reversibleintegration of biomolecules onto their platform.18 In addition, the same group reported polymer brush-coatedsurfaces with varying thermo-sensitivity and surface wettability byadjusting the amount of the Ada moiety on the polymer brush.51 While polymer brushes offer fine-tuned interfaces,their fabrication on surfaces often involves multiple steps, suchas initiator immobilization and polymerization under controlled environments.A simple approach involving coating the surface by incubation in asolution containing the polymer will offer a practical alternative.The latter approach will also benefit from the relative ease in chemicalcharacterization of the polymeric component before coating, comparedto the characterization of surface-tethered polymers. Hence, one canenvision that a polymeric surface coating capable of undergoing amodular plug-and-play integration of functional motifs such as bioactiveligands would be an attractive approach for engineering interfacesfor various biomedical and biotechnological applications.To date, supramolecular host–guest interactions havebeenextensively explored for versatile and facile modification of variousinterfaces. Fabrication methods using host–guest supramolecularinteractions have attracted significant attention since self-assemblyprocesses drive the association of building blocks at the molecularlevel.57 functionalizationsat the interfaces are achieved in a spatially controlled manner, asdemonstrated through many of the reported functionalizations.Herein,we report the fabrication of an adamantane-containing polymericcoating to modify silicon/glass surfaces and render them functionalizablein a modular fashion using CD-based host–guest interactions1. In par2Cl2) were obtainedfrom the SciMatCo purification system, and other solvents were driedover molecular sieves. Di(ethylene glycol) methyl ether methacrylate(DEGMEMA) was obtained from Sigma-Aldrich and used after filtrationthrough the activated aluminum oxide. 2,2′-Azobis(2-methylpropionitrile)(AIBN) was purchased from Sigma Aldrich and recrystallized from methanolbefore use. 3-(Trimethoxysilyl)propyl methacrylate (TMSPMA), 4-cyano-4-(thiobenzoylthio)pentanoicacid, 1-adamantanemethanol, beta-cyclodextrin (βCD), N,N,N′, N,″N″-pentamethyldiethylenetriamine(PMDETA), copper(I) bromide (CuBr), FITC-streptavidin conjugate, and4′,6-diamidino-2-phenylindole (DAPI) were purchased from Sigma-Aldrich.Diisopropylethylamine (DIPEA) was obtained from Fluka. ConcanavalinA (FITC conjugate) was purchased from Thermo Fisher. CdSeQdot 605-streptavidinconjugate and Alexa Fluor 488 phalloidin were purchased from Invitrogen.Rhodamine-PNA conjugate was purchased from Vectorlabs. d-Mannoseand d-(+)-biotin and methacryloyl chloride were obtainedfrom Alfa-Aesar. Cyclo ) was purchasedfrom Peptides International. Lissamine rhodamine B sulfonyl chloridewas purchased from Acros Organics. AdaMA monomer,58 alkyne-bearing d-mannose,58 6-O-monotosyl-βCD (βCD-OTs),59 6-monodeoxy-6-monoazido-βCD (βCD-N3),59 6-monodeoxy-6-monoamino-βCD(βCD-NH2),59 NHS-activatedbiotin (Biotin-NHS),60 βCD-biotin,60 βCD-mannose,61 and 6-monodeoxy-6-monothio-βCD (βCD-SH)62 were synthesized according to the literature procedures.All solvents were purchased from Merck andused as obtained without further purification. Ultrapure water wasobtained from a Milli-Q Water Purification System . Anhydrous toluene, tetrahydrofuran(THF), and dichloromethane (CH1H and 13C NMR spectra were recorded on Bruker Avance Ultrashield400 (400 MHz). Molecular weights of the synthesized polymers wereestimated by size exclusion chromatography (SEC) using a PSS-SDV  gram linear column calibratedwith poly(methyl methacrylate) (PMMA) standards using a refractive-indexdetector with a mobile phase solution of 0.05 M lithium bromide inDMAc as the eluent at a flow rate of 1 mL/min at 30 °C. Attenuatedtotal reflectance Fourier transform infrared (ATR-FTIR) spectroscopywas collected on a Thermo Scientific Nicolet 380 FT-IR spectrophotometer.ATR-FTIR instrument was equipped with a Harrick Scientific GATR accessoryand a Ge crystal for surface analysis. The MALDI-TOF MS spectrum acquisitionfor the characterization of RGD-CD molecule was conducted on a ShimadzuAXIMA Performance Instrument equipped with a 337 nm N2-laser. The spectrum was acquiredin the positive ion reflection mode. 2,5-Dihydroxybenzoic acid (10mg/mL) in acetonitrile/water (1:1 v/v) with the cationization agent(sodium chloride–NaCl) was used as a matrix. Static water contactangle measurements were performed using a CAM 101 KSV goniometer.Images were obtained using an integrated digital camera observingthe deposited surface of the deionized water (5 μL). Data processand contact angle determination were carried out by using CAM2008software. Contact angle values were taken from three different locationsof each sample, and average data were reported. X-ray photoelectronspectroscopy (XPS) was carried out using a K-Alpha instrument (ThermoScientific). Fluorescence images were obtained using a Zeiss ObserverZ1 fluorescence microscope with an Axiocam MRc5 camera . The polymer film thicknesswas determined using atomic force microscopy using a CoreAFM instrument.75-AdaMA20-TMSPMA5) (P(D75-A20-T5)), DEGMEMA , TMSPMA, and AdaMA  were taken ina round-bottom flask with a magnetic bar and purged with nitrogen.Under nitrogen, anhydrous DMF (8 mL) was added, and the monomer solutionwas degassed with nitrogen. 4-Cyano-4-(thiobenzoylthio)pentanoic acid as a chain transfer reagent and AIBN  as initiator was added tothe mixture. Subsequently, the reaction flask was sealed and heatedat 70 °C for 19 h. After 19 h, the reaction mixture was cooledto room temperature, and DMF was removed under reduced pressure. Thecrude was redissolved with minimum CH2CI2, andthe polymer was precipitated in cold diethyl ether to remove unreactedmonomers. Molecular weight (Mn) and molecularweight distribution (PDI) were determined by size exclusion chromatography(SEC)  = 1.36). Since the protonresonances from the TMS-monomer overlaps with other protons, it isdifficult to estimate the amount of TMSPMA incorporation. However,since it is only 5% in the feed, as an approximation, we ignored itscontribution. The peak from Ada at 2.01 ppm was used for integration and compared to the OCH3 peak at 3.39 ppm, from the DEGMEMA monomer. Using this comparison,a ratio close to the feed ratio was observed, as given in 95-AdaMA0-TMSPMA5) (P(D95-A0-T5)). Molecular weight (Mn) and molecular weight distribution (PDI) forthe control polymer were determined using size exclusion chromatography(SEC)  = 1.40).For the synthesis of poly(DEGMEMA2) were cleaned using water (5 mL), acetone(5 mL), and ethanol (5 mL), respectively. They were sonicated in thesesolvents for 15 min and then dried under a stream of nitrogen. Finally,silicon surfaces were cleaned using the Novascan PSD series UV/Digitalozone system for 15 min.Silicon surfaces(1 × 1 cm75-A20-T5) and P(D95-A0-T5)  were spread over each silicon surface.They were left open to air for 24 h at room temperature to let thesolvent evaporate. Surfaces were placed into the vacuum oven at 60°C for 1 h. Finally, surfaces were washed and sonicated in DMFand THF for 2 min to remove the nonadhered polymer from the surface,and then, wafers were dried under a stream of nitrogen.Two polymer solutions of P and lissamine rhodamineB sulfonyl chloride  were dissolved in 0.8 mL ofanhydrous DMF. After adding N,N-diisopropylethylamine (DIPEA)  in 0.2 mL of anhydrous DMF,the mixture was stirred for 2 days at room temperature. At the endof 2 days, the mixture was added to acetone (50 mL), and the precipitatedsolid residue was centrifuged and washed with an excess amount ofacetone. The product was obtained as a purple solid. 1H-NMR 8.51 , 8.17 , 7.54 , 7.05–6.88, 6.77 , 5.05 , 3.91–3.44 , 1.34–1.24.βCD-NH2 PDMS stamp.The stamp was left to dry for 60 min, followed by further drying undera gentle stream of N2. The stamp was placed on the polymer-coatedS(D75-A20-T5) surfaces for 60 min.After printing, surfaces were washed with cold water to remove nonconjugatedmaterials. The S(D95-A0-T5) surfacewas used as a control because it did not include the adamantane groupto make a complex with the βCD ligand.A solution of βCD-grafted lissamine rhodamine B dye in water(1 mg/mL) was used to wet a 1 × 1 cm75-A20-T5) surfaces and control S(D95-A0-T5) surfaces and incubated for 24 h. After incubation, surfaceswere washed with cold water to remove unconjugated βCD grafted-biotin.For the immobilization of streptavidin, 10 μL of Qdots-streptavidinwas diluted with 90 μL of water, and 10 μL from this solutionwas spread over a 1 × 1 cm2 PDMS stamp. The stampwas left to dry for 60 min, followed by further drying under a gentlestream of N2. The stamp was placed onto the polymer-coated surfacesfor 60 min. After printing, surfaces were washed with cold water toremove nonconjugated materials.βCD-grafted-biotin solution was spread over Ada-containing S in DMF was mixed with CuSO4.5H2O and sodium ascorbate in H2O. The mixture wasstirred at 60 °C overnight. Afterward, DMF was removed, and aconcentrated mixture was added to acetone (50 mL). The formed residuewas washed with acetone (50 mL × 3 times). The solid productwas redissolved in water (5 mL), and the copper-removing resin (CupriSorb)was added. The mixture was stirred until the color mixture transformedfrom blue to colorless. Finally, the mixture was filtered to removeresin and lyophilized after freezing  1H-NMR  8.46 , 5.08 , 4.02–3.81 , 3.67–3.57 .Mono-mannose substitutedβCD was synthesized by adapting a literature procedure.75-A20-T5) andcontrol S(D95-A0-T5) surface, andsurfaces were incubated for 24 h. After incubation, surfaces werewashed with cold water to remove unconjugated βCD-Man. FITC-ConAand rhodamine-PNA conjugates were used to evaluate lectin-sugar interaction.FITC-ConA  was spreadover PDMS stamps (1 × 1 cm2). The stamps were leftto dry for 60 min, followed by further drying under a gentle streamof N2. The stamps were placed onto the S(D75-A20-T5) and S(D95-A0-T5) polymer-coated surfaces for 60 min. After printing,surfaces were washed with cold water to remove nonconjugated materials.Likewise, RhB-PNA solution (0.15 μg/mL in 20 mM HEPES buffer)was spread over a PDMS stamp (1 × 1 cm2). After drying,the stamp was placed on the S(D75-A20-T5) surface for 60 min. After printing, surfaces were washedwith cold water to remove nonconjugated materials.βCD-mannosesolution  was spread over the Ada-containingpolymer-coated S and maleimide-terminatedcyclic RGD peptide   were dissolved in anhydrous DMF (0.2 mL), and Et3N was added. The mixture was stirred at room temperature overnight.After that, the reaction mixture was concentrated and precipitatedin cold Et2O . 1H-NMR: 7.47–7.25 , 5.07 , 4.00–3.74 , 3.69–3.53 . MALDI-TOF(m/z) calcd for C76H116N10O44S [M]+ 1904.69, found[M + Na + H2O] 1945.35 g/mol.βCD-SH was synthesizedaccording to the literature.75-A20-T5) and control S(D95-A0-T5) surfaces, and surfaces wereincubated for 24 h. Finally, surfaces were washed with cold waterto remove unconjugated cRGD-βCD.The βCD-cRGDsolution  was spread over Ada-containingS(D2-containingatmosphere at 37 °C and cultured in Dulbecco’s ModifiedEagle’s Medium (DMEM) media supplemented with 10% fetal bovineserum (FBS) (PAN Biotech). Cells  were seededonto polymer-coated silicon surfaces (1 × 1 cm2) andincubated at 37 °C in a 5% CO2-containing atmospherefor 2 h. Subsequently, DMEM (1 mL) was added to each well containingthe polymer-coated silicon wafers. After 24 and 48 h incubation, cellmedia was removed, silicon surfaces were washed with 1 × PBS, and cells were fixed with 3.7% formaldehyde solutionin PBS for 10 min at room temperature. After fixation, cells werewashed with 1 × PBS . For staining filamentousactins (F-actins), cells were incubated with 0.1% Triton X-100 inPBS for 5 min at 37 °C. After washing with 1 × PBS , cells were incubated with Alexa Fluor 488 phalloidinsolution (5 units/mL) in PBS, including 1% bovine serum albumin (BSA)for 15 min at 37 °C. After washing with PBS, cell nuclei werestained with DAPI for 10 min at 37 °C. After the staining steps,the stained cells were washed with 1 × PBS. The resulting imagesof stained cells on polymer-coated silicon surfaces were taken usinga fluorescence microscope, and processing was carried out using aZeiss Zen Blue lite software .L929mouse fibroblast cells were grown in a 5% COx-Ay-Tz))where x, y, and z subscript numbers show their mol percentage in the feed) were synthesized.Two different copolymers, P(D95-A0-T5) and P(D75-A20-T5), with 0 and20% Ada groups, respectively, were synthesized (63 CopolymerP(D95-A0-T5), devoid of any Ada groups,was synthesized for control experiments to deduce the role of theAda group in surface functionalization through complexation with CD.After the polymerization, copolymers were purified by precipitatingin cold diethyl ether and were characterized by SEC and 1H NMR spectroscopy. Obtained copolymers showed high monomer conversions,and SEC analysis demonstrated reasonable control on molecular weightduring polymerization with relatively narrow molecular weight distribution.Successful incorporation of monomers in the obtained copolymers wasdeduced using 1H NMR spectroscopy (S2). Forcopolymer P(D75-A20-T5), proton resonancefrom the methoxy groups (−OCH3) of DEGMEMA was observedat 3.39 ppm, along with the proton resonances of adamantane hydrogensbetween 2.01 and 1.54 ppm. Interestingly, the proton resonances fromthe silyl-containing monomers were not visible, most likely due topeak overlaps. Notably, copolymers synthesized without any silyl-basedmonomer to the feed did not show any surface adhesion in subsequentsurface modification studies.For the synthesis of copolymers bearing Ada units, adamantane methacrylate(AdaMA) was synthesized and copolymerized with a hydrophilic di(ethyleneglycol)-based methacrylic monomer (DEGMEMA), and surface active trimethoxysilyl-basedmethacrylic monomer (TMSPMA) using RAFT polymerization 2. Copolythesized 1. The TMtroscopy 1 and S2.2 surfaces were used tofabricate polymer coating on surfaces. P(D75-A20-T5) and P(D95-A0-T5)polymer solutions were prepared in anhydrous DMF (20 mg/mL). Afterspreading polymer solutions over Si/SiO2 surfaces, incubating,and heating at 60 °C for 60 min, surfaces were rinsed with copiousamounts of organic solvents to remove unbound polymers. Polymer-coatedsurfaces obtained using P(D75-A20-T5) and P(D95-A0-T5) are referredto as S(D75-A20-T5) and S(D95-A0-T5) in subsequent sections, and “S”stands for “surface”. The coated surfaces were analyzedusing FT-IR spectroscopy. The presence of the characteristic carbonylbands around 1727 cm–1 suggested the successfulattachment of polymers onto the Si/SiO2 surfaces (75-A20-T5) and S(D95-A0-T5) also suggested the same. While the water contactangle of S(D95-A0-T5) was found as57 ± 4, that of S(D75-A20-T5) was determined as 74 ± 4, presumably due to the hydrophobicnature of the adamantane groups (75-A20-T5) was undertaken with XPS analysis.The survey scan of S(D75-A20-T5)comprises C1s and O1s signals 285.0 and 532.5 eV. The high-resolutionC1s spectrum could be deconvoluted to three Gaussians with the expectedrelative areas for three carbon atoms at 285.0 eV (C–C/C–H),286.4 eV (C–O–C/C–N/C–S), and 289.3 eV(C=O) , but this is expected since the polymer contains silyl groups.In addition, the thickness and surface roughness (Ra) of S(D75-A20-T5) surfacewere found as 71 ± 16 nm and 4.2 ± 1.2 nm using AFM analysis(Ra = 0.64 ± 0.14 nm for uncoatedsilicon surface). After characterization, the stability of polymercoatings in aqueous media was investigated. S(D75-A20-T5) and S(D95-A0-T5) surfaces were incubated in PBS solution (pH = 7.4) at 37°C for 24 h. Dried surfaces were characterized using FT-IR spectroscopy.No significant change in the carbonyl peaks after incubation withPBS suggested that polymer coatings were stable in an aqueous environment.Cleaned Si/SiOsurfaces 2b. In ade groups 2c, as weeV(C=O) 2d. ImporeV(C=O) 2e,f. The75-A20-T5) and S(D95-A0-T5) was undertakenwith βCD-lissamine rhodamine B via the μCP method usinga PDMS stamp (75-A20-T5) surface wasbrought in conformal contact with a patterned PDMS stamp inked withthe dye. After printing, the surface was washed with cold water toremove any unbound dye. For the control experiment, the S(D95-A0-T5) surface was used and treated with βCD-lissaminerhodamine B dye in the same manner. After printing, surfaces werevisualized under a fluorescence microscope. While S(D75-A20-T5) surface revealed red fluorescence(95-A0-T5) did not show any fluorescentpattern, presumably due to the lack of host–guest interactionon the surface devoid of any adamantane groups (75-A20-T5) modified with βCD-grafted lissamine rhodamine B dye was immersedin PBS buffer (pH 7.4) and incubated at 37 °C for 48 h and analyzedusing fluorescence microscopy at predetermined time intervals. Thefluorescence intensity of the βCD-grafted lissamine rhodamineB modified S(D75-A20-T5) surfaceshowed no significant change when incubated at 37 °C for 48 h. Observations suggest significantstability of thus modified polymer surface βCD/Ada-based host–guestinteractions.Initial functionalization of S.64 To this end, a βCD-biotin conjugated S(D75-A20-T5) surface was modified withFITC-streptavidin with and without a pattern (95-A0-T5), devoid of any adamantyl group, was used for controlexperiments. In addition to the S(D95-A0-T5) surface, the S(D75-A20-T5) surface, which was not modified with βCD-biotin, was alsoutilized for the control experiment. After treatment of the biotinylatedsurface with FITC-streptavidin, surfaces were washed with PBS to removeany unconjugated material. Upon visualization under a fluorescencemicroscope, green fluorescence was obtained for both patterned andnonpatterned biotinylated surfaces , further modificationwas demonstrated using the CdSe Qdot-streptavidin conjugate. Thiswas undertaken to demonstrate that such functionalizable interfacescan be also modified with comparatively large inorganic nanoparticles.In the first step, βCD-grafted biotin (βCD-biotin) wasimmobilized onto S(D75-A20-T5) byhost–guest interactions. After immobilization, the surfacewas washed with cold water to remove unbound βCD-biotin. Also,the control surface S(D95-A0–T5) was treated with βCD-biotin similarly. After conjugationof βCD-biotin, the Qdot-streptavidin conjugate was immobilizedonto a biotinylated and control surface with μCP (75-A20-T5) (95-A0-T5) revealed no fluorescence sinceno binding occurred due to the lack of adamantane groups (After the conjugation of FITC-streptavidin ontobiotin-immobilizedS(Dwith μCP 5a. After-A20-T5) 5b,c. As e groups 5c, inset75-A20-T5) was also characterized by XPS. The surveyscan of biotin-conjugatedS(D75-A20-T5) comprises C1s and O1ssignals 285.0 and 532 eV. The high-resolution C1s spectrum could bedeconvoluted to three Gaussians with the expected relative areas forthree carbon atoms at 285.0 eV (C–C/C–H), 286.2 eV (C–O–C/C–N/C–S), and 288.8eV (C=O) (75-A20-T5) surface,where the characteristic N atom peak was visible at 400.0 eV. In addition,the S2p peak coming from biotin was also observed at 164.2 eV (65 suggest successful immobilization of βCD-biotinonto the Ada-containing S(D75-A20-T5) via host–guest interactions. In addition to fluorescencemicroscopy investigations, CdSe Qdot-streptavidin conjugated biotin-S(D75-A20-T5) was also characterized byXPS. The characteristic N1s (66Biotin-conjugated surface S(DeV (C=O) 6a. Figur164.2 eV 6c. N1s astic N1s 6e and S2stic N1s 6f peaks stic N1s 6e.6675-A20-T5) and control surfaceS(D95-A0-T5) were treated with βCD-bearingmannose. Unbound βCD-bearing mannose was removed from the surfaceby rinsing the surfaces with water. βCD-Man-S(D75-A20-T5) and control S(D95-A0-T5) surfaces were subsequently evaluated for ConA binding (75-A20-T5)and control S(D95-A0-T5) surfacesbrought in conformal contact with a microstructured PDMS stamp primedwith a solution of FITC-ConA. After washing with HEPES (20 mM) solution,to remove any physically adhered FITC-ConA, the silicon surface wasvisualized under a fluorescence microscope. As expected, while theβCD-Man-S(D75-A20-T5) surfacedisplayed green fluorescence with clear patterns (95-A0-T5) surface did not show any fluorescence(75-A20-T5) and control S(D95-A0-T5) surfaces were treated with RhB-PNA using the μCP method . For this purpose, Ada-containing surfaceS surface and conjugated ConAshould decrease in the presence of a high concentration of a competitiveligand such as free sugar, and reduced interaction should lead tothe release of ConA from the surface. To this end, FITC-ConA printedβCD-Man-S(D75-A20-T5) surfaceswere incubated with d-(+)-mannose solution to enable therelease of ConA. Upon visualization under a fluorescence microscope,no green fluorescence on the surface was observed after treatmentwith a d-(+)-mannose solution (75-A20-T5) surfacewas also investigated, and it was observed that FITC-ConA could beimmobilized onto regenerated βCD-Man-S(D75-A20-T5) surfaces. As seen in 75-A20-T5) surface could immobilize FITC-ConAin subsequent cycles with almost similar efficiency . S(D75-A20-T5) surfaces were treated with cRGD-conjugatedβCD to obtain an interface displaying the cRGD peptide. Afterincubation of the surface with peptide-containing βCD solution,unbound βCD-derivatives were rinsed from the surface with water.To confirm that the peptide was immobilized onto the surface throughhost–guest interactions, the peptide-decorated surface S(D75-A20-T5) was characterized by XPS.The survey scan cRGD peptide conjugated S(D75-A20-T5) comprises C1s and O1s signals at 285.0 and 533 eV,respectively . The high-resolutionC1s spectrum could be deconvoluted to three Gaussians with the expectedrelative areas for three carbon atoms at 285.0 eV (C–C/C–H), 286.0 eV (C–O–C/C–N),and 289.1 eV (C=O) . Notably, the characteristic N1s peak from thecRGD peptide was observed at 400 eV , confirming the peptide’s successful attachment onto thesurface.Finally, we wanted to investigate if polymer-coated surfaces couldbe rendered amenable for cell attachment through noncovalent decorationwith cell-adhesive peptides. To this end, cRGD-conjugated βCDwas synthesized using maleimide-terminated cRGD and βCD-SH inthe presence of triethylamine via the thiol-maleimide Michael additionreaction (75-A20-T5)). S(D75-A20-T5) without RGD attachmentand S(D95-A0-T5) surfaces were utilizedfor the control experiment. After 24 and 48 h incubation, the actinand nuclei of attached cells were stained using an Alexa fluor-phalloidin-488kit and DAPI, respectively. After the staining processes, images ofcells on the polymer-coated surfaces were obtained using a fluorescencemicroscope. In the case of immobilization of the polymer surface withcRGD-βCD, enhanced cell attachment and proliferation were observedafter 24 and 48 h (75-A20-T5) (not treated with cRGD-βCD)and the control surface S(D95-A0-T5) remained significantly low due to the anti-biofouling characterof the PEG-based copolymer coating.After the surface immobilization of cRGD-βCD,L929 mouse fibroblast cells were seeded onto the peptide-modifiedsurface (RGD-βCD-S(Dand 48 h 10b. As aIn summary, the fabrication of a versatilepolymeric coating amenableto facile functionalization in a plug-and-play modular manner to immobilizevarious functional molecules ranging from fluorescent dyes to bioactiveligands is demonstrated. An Ada-CD-based host–guest complexationon a polymeric interface enables a variety of functionalization ina robust and reliable manner. Examples here demonstrate the modificationof polymer-coated surfaces for immobilizing fluorescent dyes, specificproteins, and protein-coated inorganic nanocrystals. Protein immobilization/sensingcan be undertaken selectively by appropriate choice of the functionalmolecules at the interface. Through catch-and-release cycles, we demonstratethat obtained interface is robust enough for regeneration and reusabilityin specific applications. Finally, we show that the inherently anti-biofoulingpolymeric interface can be tailored for the attachment of cells throughmodification with cell-adhesive peptides. One can envision that thefacile fabrication and versatile and modular nature of the interfaceengineering demonstrated here will be deemed attractive for fabricatingfunctional devices for various biomedical applications.


In acetogenic bacteria, the electron acceptor is CO2 that is reduced to enzyme bound carbon monoxide, the precursor of the carboxyl group in acetate. Here, we describe the generation of a mutant of the acetogen Acetobacterium woodii in which the last step in methyl group oxidation, formate oxidation to CO2 catalyzed by the HDCR enzyme, has been genetically deleted. The mutant grew on glycine betaine as methyl group donor, and in contrast to the wild type, formed formate alongside acetate, in a 1:2 ratio, demonstrating that methyl group oxidation stopped at the level of formate and reduced electron carriers were reoxidized by CO2 reduction to acetate. In the presence of the alternative electron acceptor caffeate, CO2 was no longer reduced to acetate, formate was the only product and all the carbon went to formate. Apparently, acetogenesis was not required to sustain formatogenic growth. This is the first demonstration of a genetic reprogramming of an acetogen into a formatogen that grows by homoformatogenesis from methyl groups. Formate production from methyl groups is not only of biotechnological interest but also for the mechanism of electron transfer in syntrophic interactions in anoxic environments.Methyl groups are abundant in anoxic environments and their utilization as carbon and energy sources by microorganisms involves oxidation of the methyl groups to CO Reoxidation of these electron carriers is coupled to the reduction of internal or external electron acceptors. In methanogens, methyl groups also serve as electron acceptors and are reduced to methane [2 is reduced to enzyme bound CO, followed by condensation with a methyl group to yield acetate [Acetobacterium woodii, 23 different MTI enzymes are encoded [Methyl groups serve as carbon and energy sources for microbial growth in anoxic environments; they are oxidized to CO methane , 2, in s methane –5 or in  acetate –8. In th acetate , 10. One acetate , 11–14:  encoded  and for  encoded , 7.A. woodii by the filamentous hydrogen-dependent carbon dioxide reductase (HDCR) [+ gradient to drive this endergonic reaction [+ in equimolar amounts with hydrogen as reductant [Methyl-THF is then oxidized via methylene- and methenyl-THF to formate –8, whiche (HDCR) –18 to cae (HDCR) , 20 is re (HDCR) , 22 but reaction  or by theductant . In sum,This metabolism yields 0.63 mol of ATP per mol of methyl group.2 is observed in pure cultures in the presence of CO2 in the gas phase. However, when methanol was fed in combination with other electron donors like H2 or CO, the fate of the methyl group changed: it was no longer oxidized but exclusively converted to acetate [Acetogenesis by disproportionation of methyl groups to acetate and CO acetate . Moreove acetate , 26. The acetate .A. woodii and other acetogens that have the HDCR enzyme, the oxidative and reductive parts of metabolism are connected by a hydrogen cycle [A. woodii [2 reduction by methanogens. This “oxidation only mode” was observed during growth on fructose but also during growth on methanol in coculture of A. woodii with a H2-consuming methanogen that removed the hydrogen thereby enabling this lifestyle energetically by removing the end product [2 + CO2 transiently, indicating the capacity to export and import formate [A. woodii. Indeed, by deleting the hdcr gene cluster we turned A. woodii into a formate producer that grew non-acetogenically on methyl groups.In syntrophic communities, there is another way for the reoxidation of reduced electron carriers. These communities are characterized by interspecies electron transfer and the electron is transferred from one cell to the other as molecular hydrogen or as formate . In A. wen cycle . Genetic. woodii . Of cour product , 30. For product  and thus formate –33. ThisA. woodii DSM1030 (wild type), ∆hydBA, ∆hdcr and ∆hydBA/hdcr were routinely cultivated under anoxic conditions at 30 °C in carbonate buffered complex medium as described previously [pyrE strain, 50 mg/l uracil was added to the medium [600).eviously . Since ae medium . As carbhydBA mutant was described before [hdcr operon , the plasmid pMTL_AW_KO_HDCR was generated in E. coli HB101  and transformed into A. woodii ΔpyrE or ∆hydBA, as described previously [hdcr and hydBA/hdcr, the entire hdcr operon was deleted, leaving only the start codon of fdhF1 and stop codon of hydA2 in the chromosome. Plasmid pMTL_AW_KO_HDCR is based on pMTL84151 [XmnI and FspI following a blunt-end ligation. It further contains the pyrE cassette from Eubacterium limosum KIST612, consisting of the pyrE gene (ELI_0961) and 66 bp of its promoter region, as described before [hdcr gene cluster, a hdcr deletion cassette was cloned into the multiple cloning site, consisting of a 1256 bp upstream flanking region ending with the start codon of fdhF1 and a 1085 bp downstream flanking region starting with the stop codon of hydA2. Both flanking regions were amplified via PCR, joined by splice-by-overlap-PCR (SOE-PCR) [EcoRI and XbaI. A detailed description of transformation into A. woodii and integration of suicide plasmids for deletion of genes as well as recombination of the plasmid at its homologous regions towards the loss of genes has been published before [hdcr operons in the A. woodii ΔpyrE strain and the ΔhydBA mutant, which lead to the loss of the 8567 bp of the hdcr operon, the deletion was verified by PCR and Sanger sequencing analysis [Generation and characterization of the ∆d before . To deleeviously . In bothMTL84151  from whid before . For tarSOE-PCR)  and clond before . After ianalysis .hdcr genes from A. woodii  downstream of a constitutive pta-ack promoter from C. ljungdahlii. Transformation of the plasmid into the Δhdcr mutant was performed as described before [2 + CO2, glycine betaine and formate.For the genetic complementation, plasmid pMTL84211_JM_Pptaack_hdcr was constructed. The plasmid is derived from pMTL84211  and contd before  and the d before . Subsequ600 of 0.3) and then harvested by centrifugation  at 8000 rpm and 4 °C for 10 min. The harvested cells were subsequently washed with 30 ml of buffer containing 50 mM imidazole (pH 7.0), 20 mM KCl, 20 mM MgSO4, 4 mM DTE and 4 µM resazurin by centrifugation at 8500 rpm and 4 °C for 10 min  and resuspended in 5 ml of imidazole buffer and kept in 16-ml Hungate tubes. All the steps were performed under strict anoxic conditions in an anoxic chamber  filled with N2/H2 . The total protein concentration in the resting cells was determined according to a previously described method [Cells were cultivated on 50 mM glycine betaine in 0.5 to 2 l carbonate buffered complex medium to late exponential growth phase  in 120-ml serum flasks to a total protein concentration of 1 mg/ml. As substrate, 50 mM glycine betaine, and if necessary, caffeate (4 mM) or H2 + CO2 was given. The suspension was incubated at 30 °C in water bath with shaking (150 rpm) and 0.5 ml samples were taken at each time point for determination of metabolites.Cells were resuspended in 10 ml imidazole buffer . Nitrogen was used as the carrier gas with a head pressure of 400 kPa and a split flow of 30 ml/s. The samples were analyzed with a thermal conductivity detector at 100 °C and at an oven temperature of 40 °C. The concentrations of acetate and formate were determined by high-performance liquid chromatography [+ ion exchange column  was applied and degassed sulfuric acid (5 mM) was used as eluent at a flow rate of 0.6 ml/min. The oven was kept at 65 °C. 10 μl sample was injected by auto-sampler and analyzed with a refractive index detector  at 55 °C. Caffeate was determined photometrically at 312 nm using an extinction coefficient of 13.72 mM−1·cm−1 [The concentrations of Heviously . The samM−1·cm−1 .hdcr operon in the thermophilic acetogen T. kivui, which does not grow on methyl groups, is essential for growth on H2 + CO2 but also for growth on glucose [T. kivui has the genetic potential to grow by the conversion of glucose to 2 acetate, 2 CO2 and 4 H2, like Thermotoga maritima [A. woodii, we deleted the hdcr gene cluster through allelic exchange mutagenesis and verified the deletion by PCR and sequence analysis   Fig. . In contly) Fig. . We then2 produces molecular hydrogen that is then oxidized by the electron-bifurcating hydrogenase with concomitant reduction of NAD+ and ferredoxin. Since only reduced ferredoxin but not NADH is required as reductant in this metabolic scenario, NADH is oxidized with reduction of ferredoxin by the Rnf complex  and growth of the mutant was inhibited by addition of molecular hydrogen.Oxidation of methyl groups to COcf. Fig. . To detease Fig. . Restingine Fig. . The ∆hyA mutant  also gre+ potential across the membrane, established by the hydrolysis of 1.2 ATP [2 are reduced to 2 CO and converted to 2 acetyl-CoA with 2 methyl groups from glycine betaine. In sum, one formate and 2 acetate are produced from 3 glycine betaine in the ∆hdcr mutant  and fhs2 (Awo_c08040) [fhs2 forms an operon with a gene encoding a formate transporter  [fhs2/fdhC genes led to a complete loss of transient formate production, indicating their involvement in formate production. However, formate is produced as a side product, most likely because the formyl-THF synthetase reaction is the kinetic bottleneck upon entry of formate into the THF pathway; formate production is not energy conserving.The hallmark of acetogenic bacteria is the production of acetate from two molecules of COetyl-CoA –50. The  species –33, it i_c08040) ; fhs2 fo2 and H2 from a methyl group  is endergonic  to H+ (E0’ = −414 mV) is endergonic and driven by simultaneous electron transfer from reduced ferredoxin (E0’ ≈ −450 mV) to H+. Whereas the latter reaction is exergonic with −6.9 kJ/mol, the former is endergonic with 18.5 kJ/mol, making the overall reaction endergonic with 11.6 kJ/mol. The reaction would be at equilibrium at a hydrogen partial pressure of 0.1 atm, lower concentrations making the reaction thermodynamically possible. A maximum of 0.4 ATP/mol of methyl group can be obtained mechanistically, one in the formyl-THF synthetase reaction and 0.6 ATP has to be invested in the ATPase/Rnf reaction to transfer 2 electrons from NADH to ferredoxin (Fig. Methanol oxidation in xin Fig. . The sam2 + CO2 or formate does not make a big difference and the demonstrated capability of A. woodii to grow on methyl groups by formatogenesis does not necessarily reflect that this will occur in nature. Indeed, this seems rather unlikely since A. woodii has the HDCR enzyme that oxidizes formate to H2 + CO2 with unprecedented rates [2/bicarbonate concentrations, or a lack of molybdenum or iron may disfavor formate oxidation but favor formate production. Reversal of the WLP also occurs in syntrophic oxidation of acetate [Thermoacetogenium phaeum [Tepidanaerobacter acetatoxydans [T. acetatoxydans does not have a formate dehydrogenase arguing for formate transfer between this acetogen and methanogens during syntrophic acetate oxidation [Blautia lack genes encoding a HDCR or formate dehydrogenase [Blautia species are active members of the gut microbiome [Energetically, the oxidation of a methyl group to Hed rates , 17. How acetate –57. Somem phaeum , 59 and toxydans , 61 can rogenase . Since Bcrobiome , formate2 are energetically equivalent and can be transformed into one another by soluble or membrane-bound enzymes. Since the equilibrium constant is almost one, the direction is controlled by the concentration of the reductants. Formate production may be favored in H2-rich ecosystems and electrons shuttled to the electron consuming partners via formate, not hydrogen. Such ecosystems are, for example, deep sea hydrothermal vents in which serpentinization leads to hydrogen production [Thermococcus paralvinellae isolated from a deep sea vent is a chemosyntroph that produces H2 from H+ in the absence of sulfur [2 is then used by the Methanococci. At high hydrogen pressures, H2 formation is inhibited, and formate is produced from H2 + CO2 by the formate hydrogen lyase in T. paralvinellae [Methanococci as carbon and electron source.Formate and Hoduction , 65. Thef sulfur ; H2 is tvinellae . Formate2 is inhibiting [2 inhibition. Whether or not natural formatogenic lifestyle exists in acetogens, remains to be elucidated.Formate production gives cells a growth advantage when Hhibiting . 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