Molecular Assemblies: Characterization and Applications
ACS Symposium Series1355
Molecular Assemblies: Characterization and Applications
Library of Congress Cataloging-in-Publication Data
Foreword
Preface
Discovery of Monodisperse Micelles with Discrete Aggregation Numbers
Supramolecular Assembly and Mesophase Behavior of Glycopyranose-Derived Single-Chain Amphiphiles
Self-Assembly and Aggregation Studies of Simple Structural Derivatives of Stearic Acid
Förster Resonance Energy Transfer Probing of Assembly and Disassembly of Short Interfering RNA/Poly(ethylene glycol)–Poly-L-Lysine Polyion Complex Micelles
Assemblies of Hydrophobically Modified Starch Nanoparticles Probed by Surface Tension and Pyrene Fluorescence
Simple Creams, Complex Structures
Enzyme-Triggered Nanomaterials and Their Applications
Characterization of Colloidally Stabilized Latex Particles by Capillary Electrophoresis
Editor’s Biography
Indexes
Indexes
Author Index
Subject Index
Preface
1
Discovery of Monodisperse Micelles with Discrete Aggregation Numbers
Introduction
Figure 1. Schematic illustration of Platonic micelles composed of cone-shaped amphiphiles. Adapted with permission from reference 30. Copyright 2019 American Chemical Society.
Hypothesis on the Mechanism of Platonic Micelle Formation
Figure 2. Two examples of the best packing calculation using multiple spherical caps (N = 3 or 12) on a sphere’s surface. For N = 3, the coverage density D(N), defined by the ratio of the total area of the caps to the surface area of the sphere, is 75%, whereas it is 90% for N = 12.
Figure 3. The coverage density D(N) of the Tammes problem plotted against the number of caps N and the monodisperse Naggs observed to date are indicated (Nagg = 4 25, 6 (2628), 8 (2830), 12 (27283032), and 20 (2430)).
The Effect of ae on Platonic Micelle Structures: pH-Controllable Nagg Corresponding to Regular Polyhedra
Figure 4. The chemical structures of calix[4]arene-based amphiphiles bearing glutamic acids and propyl chains (ECaL3) at different pHs. The α-amines are protonated at acidic pH (pH < pKaNH3+), and the γ-carboxylic acids are deprotonated at basic pH (pH > pKaCOOH).
Figure 5. SAXS profiles of ECaL3 micelles at pH 3.0, 8.0, and 10 in 50 mM aqueous NaCl. The atomic form microscopy and TEM images at each pH are also presented.
Figure 6. (a) I(q)/c as a function of q for different ECaL3 concentrations in 50 mM aqueous NaCl at pH = 3.0 and 10. (b) Guinier plot (i.e., ln[I{q}/c] vs q2) constructed from the SAXS intensity extrapolated to zero concentration. The inset shows the concentration dependence of I(0)/c. (c) Reversible switching of the Rg of the micelle with variation of the pH. Adapted with permission from reference 27. Copyright 2017 American Chemical Society.
Figure 7. Schematic illustration of the morphological transitions between monodisperse spherical micelles with pH-controllable Nagg. The energy-minimized molecular models of ECaL3 calculated using the SCIGRESS program (Fujitsu Ltd., Tokyo, Japan) with Molecular Orbital PACkage in water are shown, where the α-amines are protonated on the right side and the γ-carboxylic acids are deprotonated on the left side.
Effect of Alkyl Chain Length on Platonic Micelle Structures
Figure 8. (a) The chemical structure of calix[4]arene-based amphiphiles bearing quaternary amines (QACaLn). R represents alkyl chains including propyl (C3), butyl (C4), heptyl (C5), hexyl (C6), heptyl (C7), octyl (C8), and nonyl (C9) tails. (b) SAXS profiles of QACaLn micelles in 50 mM aqueous NaCl. The solid curves are calculated using theoretical models of a core-shell sphere and cylinder. (c) The Guinier plots (i.e., ln[I{q}Ke−1c−1/Munimer] vs q2, where Ke is an optical constant in SAXS) derived from the spherical micelles of QACaLn. The values of Rg of the QACaLn micelles are also indicated. Adapted with permission from reference 30. Copyright 2019 American Chemical Society.
Figure 9. (a) The concentration dependence of Munimer/Mw,App (i.e., 1/Nagg at Conc. = 0) determined by AUC measurements for QACaLn micelles in 50 mM aqueous NaCl. (b) The dependence of the Nagg of the spherical micelles composed of QACaLn in 50 mM aqueous NaCl on alkyl chain length (nc: the number of carbons in each alkyl chain). Adapted with permission from reference 30. Copyright 2019 American Chemical Society.
Platonic Micelles in Other Micellar Systems
Figure 10. (a) The chemical structure of surfactin. (b) The Nagg of surfactin micelles determined by AUC measurements plotted against NaCl concentration.
Conclusions
References
2
Supramolecular Assembly and Mesophase Behavior of Glycopyranose-Derived Single-Chain Amphiphiles
Introduction
Complex Outcomes of a Single Design Plan
Chemistry of Sugar Amphiphiles
Figure 1. General structures of sugar amphiphiles.
Mesophase Behavior of Sugar Amphiphiles
Normal Micelles
Figure 2. Reconstitution of membrane proteins into bilayer-forming molecules (bottom) from micellar-forming molecules (top) using traditional methods (left) and a native chemical ligation–based method (right).
Figure 3. Structures of several amphiphiles derived from 2-amino-2-deoxy-D-glucopyranose.
Lamellar
Tubes and Fibers
Figure 4. General chemical structures of several glucose-based amphiphiles that self-assemble into tubes and fibers. Negative staining transmission electron microscopy images for the corresponding assemblies are also shown. Reproduced with permission from references (2127), and 28. Copyright 2002 and 2005 American Chemical Society.
Figure 5. Preparation of well-defined glucopyranosyl lipid nanotubes (LNTs). (a) Rapid method to generate glycolipid-based tubular structures with defined diameters. Extrusion of an aqueous dispersion of the glycolipid at 90 °C through a polycarbonate filter produces uniform vesicles with thin walls. Afterward, the extruded vesicles are filled into the nanopores of an anodic alumina membrane filter and then self-assembled into well-defined LNTs on cooling. (b) Higher magnification scanning electron microscopy and scanning transmission electron microscopy (inset) images of the obtained LNTs. (c) Low magnification scanning electron microscopy image of the resulting LNTs. Reproduced with permission from reference 29. Copyright 2006 American Chemical Society.
Figure 6. Time-lapse fluorescence microscopic images (top) and schematic representations (bottom) of NBD-containing glucopyranosylamide nanotubes upon addition of the protein QSY7-ferritin. After treatment of the NBD-nanotubes (a) with QSY7-ferritin, both open ends of the tubes start to quench (b). The quenching gradually moves toward the central part with time (c–e), being completed in 4 s (f). Reproduced with permission from reference 31. Copyright 2007 American Chemical Society.
Vesicles
Figure 7. Structures of vesicle-forming amphiphiles derived from the disaccharides maltose (a) and lactose (b).
Figure 8. Model for the self-assembly of OTG into bilayer membrane vesicles and a phase-contrast image of the resulting OTG vesicular structures. Scale bar denotes 10 μm. Reproduced with permission from reference 43. Copyright 2018 American Chemical Society.
Figure 9. CVC for OTG.
Figure 10. RCA in OTG-based vesicular microreactors. Scale bars denote 5 μm. Reproduced with permission from reference 43. Copyright 2018 American Chemical Society.
Figure 11. Chemical structures of the alkyl galactopyranosylamides GOA, GPOA, and GMOA, and negative staining transmission electron microscopy image of GOA vesicles. Scale bar denotes 100 nm. Reproduced with permission from reference 44. Copyright 2019 American Chemical Society.
Figure 12. Differential scanning calorimetry thermograms of aqueous dispersions of the acyl galactopyranosylamides GOA, GPOA, and GMOA. Reproduced with permission from reference 44. Copyright 2019 American Chemical Society.
Inverted Mesophases
QII
Figure 13. Structures of sugar-based amphiphiles forming QIIs. (a) 1-O-phytanyl-β-D-xyloside, (b) 1-O-(5,9,13,17-tetramethyloctadecyl)-β-D-xylopyranoside, (c) 1-O-(5,9,13,17-tetramethyloctadecyl)-β-D-glucopyranoside.
HII
Challenges and Future Directions
References
3
Self-Assembly and Aggregation Studies of Simple Structural Derivatives of Stearic Acid
Introduction
Figure 1. Crystal packing of stearic acid polymorphs. Adapted with permission from reference 6. Copyright 2007 Royal Society of Chemistry.
Figure 2. Structures of simple structural derivatives of stearic acid.
Lyotropic Liquid Crystalline and Gelation Properties of Stearic Acid
Figure 3. (A) POM of the stearic acid (1) and triethanolamine (2:1 molar ratio) in aqueous solution at room temperature. (B) Bright-field microscopy image. Adapted with permission from reference 26. Copyright 2007 American Chemical Society.
Figure 4. Micrograph of 2 wt % stearic acid (1) in sunflower oil. Bar: 200 nm. Adapted with permission from reference 30. Copyright 2003 John Wiley & Sons.
Figure 5. Bright-field micrographs of the gel formation of a 25 wt % stearic acid (1) in sesame oil mixture: (A) 0 s and (B) 1 min of gelation. Adapted with permission from reference 31. Copyright 2015 Elsevier.
Gelation Properties of Stearamide
Figure 6. Photographs (left) of stearamide (2) gel in isopropyl myristate (2 wt %, 1), olive oil (1 wt %, 2), and squalene oil (2 wt %, 3). Polarizing optical microscopic images of 2 wt % stearamide (2) in squalene gels (a and b). Adapted with permission from reference 33. Copyright 2014 Royal Society of Chemistry.
Self-Assembly, Gelation, and Liquid Crystalline Properties of N-(Hydroxyalkyl)stearamide
Figure 7. Optical micrographs without (A) and with (B) crossed polarizers of water penetration of N-(2-hydroxyethyl)stearamide (3, n = 2) at 90 °C. Adapted with permission from reference 34. Copyright 2010 American Chemical Society.
Figure 8. (A) Log–log frequency sweep (0.05% strain) at 25 °C for a 2 wt % N-(3-hydroxypropyl)stearamide (3, n = 3) in isostearyl alcohol gel: G′, squares; G′′, circles. (B) Plots of G′ (square) and G′′(circle) versus time for gels consisting of 2 wt % N-(3-hydroxypropyl)stearamide (3, n = 3) in isostearyl alcohol gel. Adapted with permission from reference 35. Copyright 2015 Royal Society of Chemistry.
Self-Assembly and Lyotropic Liquid Crystalline Phase Behavior of N,N-Bis(2-hydroxyethyl)stearamide
Figure 9. (A) Polarized optical micrograph of N,N-bis(2-hydroxyethyl)stearamide (4). Images were acquired at 55 °C with crossed polarizers. (B) N,N-Bis(2-hydroxyethyl)stearamide (4) water system. The diagrams consist of SAXS data (data points) superimposed on POM data (dotted lines). SAXS results show three lamellar crystalline phases: Lc1 (◻), Lc2 (X), and Lc3 (▼). They also show a fluid lamellar (Lα) phase (●) and an isotropic fluid phase (▲). Adapted with permission from reference 41. Copyright 2012 Elsevier.
Nanotubules and Microspheres from the Self-Assembly of N-(Aminoalkyl)stearamides
Figure 10. Scanning electron microscopy images of the self-assembled structures of compounds: (A, B) nanotubules formed from N-(aminoethyl)stearamide (5, n = 2) in 1,2-dichloroethane at a low concentration (0.04 wt %); (D, E) N-(aminoethyl)stearamide (5, n = 2) at a high concentration (0.16 wt %) in 1,2-dichloroethane; (C) nanobelt-like structure from N-(aminopropyl)stearamide (5, n = 3) in 1,2-dichloroethane; (F, G) microspherical structures from (F) N-(aminobutyl)stearamide (5, n = 4) in 1,2-dichloroethane; and (G) N-(aminohexyl)stearamide (5, n = 6) in 1,2-dichloroethane. Adapted with permission from reference 43. Copyright 2010 American Chemical Society.
Figure 11. Transmission electron microscopy images of the twisted structure from N-(aminoethyl)stearamide (5, n = 2) in 1,2-dichloroethane before forming the complete nanotubular morphology (a) and nanotubular structure (b, c). The inset in (c) is a scanning electron microscopy image of the nanotubule. The scale bar in all images is 100 nm. Adapted with permission from reference 43. Copyright 2010 American Chemical Society.
Figure 12. Mechanism for the formation of (a) nanotubules and (b) nanobelts and microspheres from N-(aminoalkyl)stearamide (5) in 1,2-dichloroethane. Adapted with permission from reference 43. Copyright 2010 American Chemical Society.
Gelation Studies of Stearic Acid Derivatives of N-Acyl Amino Acids and Esters as Molecular Gelators
Figure 13. IR spectra of (A) methyl stearoyl-L-serinate (7, R = -CH2OH) in hexane gel and (B) methyl stearoyl-L-serinate (7, R = -CH2OH) in chloroform. Adapted with permission from reference 44. Copyright 2012 Elsevier.
N-Phenylstearamides as Low Molecular Mass Gelators
Figure 14. Appearance of 2 wt % molecular gels of N-phenylstearamide (8, X = H) in (a) cyclohexane, (b) benzene, (c) toluene, (d) ethyl acetate, (e) acetonitrile, (f) ethanol, and (g) dimethyl sulfoxide. Adapted with permission from reference 50. Copyright 2018 Royal Society of Chemistry.
Conclusions
Acknowledgments
References
4
Förster Resonance Energy Transfer Probing of Assembly and Disassembly of Short Interfering RNA/Poly(ethylene glycol)–Poly-L-Lysine Polyion Complex Micelles
Introduction
Incorporating siRNA in Polyion Complex Micelles
Figure 1. Schematic of a polyion complex micelle formed from the complexation of siRNA with the double-hydrophilic block copolymer, PEG–PLL. PEG is neutral while PLL is cationic. Adapted with permission from reference 9. Copyright 2017 American Chemical Society.
Discerning State of Assembly versus Disassembly Using Fӧrster Resonance Energy Transfer
Figure 2. (a) siAF594 and siAF647 exist outside a micelle and are far apart. When excited at 540 nm, siAF594 emits energy (as indicated by a star) while siAF647 does not. siAF647 is also unaffected by the emission of siAF594 because they are far apart (no FRET, as indicated by the circle). (b) siAF594 and siAF647 coexist inside a micelle, where they are in close proximity for siAF594 to donate some of its emission energy to siAF647. The siAF647 then also emits energy. (FRET occurs, as indicated by the circle having changed to a star on siAF647.)
Materials and Methods
siRNA
PEG–PLL Block Copolymer
Micelle Formation
Dynamic Light Scattering for Micelle Size
FRET Measurements for State of Assembly or Disassembly
Results and Discussion
DLS Confirms Molecular Assembly
Figure 3. DLS data showing intensity-based size distributions. (a) Size distribution of micelles formed at charge neutrality (N/P = 1) shows a peak at 65 nm in diameter. (b) Size distribution of micelles formed at charge neutrality (N/P = 1) and stored for 3 days prior to the DLS measurements to examine their storage stability. The peak of the distribution occurs at 65 nm in diameter. (c) Size distribution of micelles formed at varying N/P ratios shows changes in the concentration of micelles (intensity variations) and some changes in the hydrodynamic diameter (change in location of the peak of the size distribution).
Model-Based Interpretation of Micelle Formation Behavior
FRET Confirms State of Assembly versus Disassembly
FRET for Naked siRNA Control
Figure 4. Fluorescence emission from siAF594 and siAF647 controls when present individually in HEPES buffer to demonstrate natural emission peaks. The molecules in solution were excited at 540 nm.
FRET for Neutral Micelles
Figure 5. Fluorescence emission scans for 1:1 molar mixture of the siRNA duplex with differing labels in the absence of the block copolymer and for a mixture of siRNA duplex with PEG–PLL block copolymer at N/P = 1. The two spectra compare the siRNA in a naked state and siRNA hosted within a micelle. The excitation was done at 540 nm.
FRET for Charged Micelles
Figure 6. Fluorescence emission scans for micellar solution excited at 540 nm for varying N/P ratios. The micelles are prepared with a 1:1 molar mixture of the siRNA duplex with differing labels added to the PEG–PLL solution to arrive at the specified N/P ratios.
Figure 7. Dependence of FRET on the N/P ratio. FRET values are normalized relative to those for naked siRNA in the absence of the block copolymer. The naked siRNA controls represent the condition of no energy transfer between the donor and acceptor dye labels on the siRNA duplex.
FRET Study of Micelle Disassembly and siRNA Release
Figure 8. Micelle stability and disassembly over time determined by FRET. The FRET value at time (t) was normalized with respect to that at time 0 when the solution was excited at 540 nm. Micelles were exposed to FBS, HEPES, DMEM, heparin, and trypsin to simulate different physiological conditions to determine their stability and the ability to disassemble and promote siRNA release.
Conclusions
Acknowledgments
References
5
Assemblies of Hydrophobically Modified Starch Nanoparticles Probed by Surface Tension and Pyrene Fluorescence
Introduction
Experimental Section
SNP and HM-SNP Aqueous Dispersions with Pyrene
Scheme 1. Chemical composition and structure of the HM-SNP samples used in this study. Substitution is shown in the C6 position of the anhydroglucose unit for convenience, but it can also occur at the C2 and C3 positions.
Steady-State Fluorescence
Time-Resolved Fluorescence Decays
Analysis of the Fluorescence Decays
Ultraviolet–Visible Absorption
Surface Tension
Results and Discussion
Figure 1. Fluorescence spectra (A) before and (B) after normalization at 372 nm, and (C) fluorescence decays for 0.5 μM pyrene aqueous solutions upon increasing the C6(0.15)-SNP1 concentration from 0 to 13.6 g/L. The traces in bold are for 0.5 μM pyrene in water: λex = 335 nm. The figures illustrate the influence of the C6(0.15)-SNP1 concentration on (A) the fluorescence intensity, (B) the spectral features, and (C) the lifetime of pyrene.
Figure 2. UV–vis spectra of the MAC of pyrene in (A) water and (B) a 1.9 g/L aqueous dispersion of C6(0.15)-SNP. The differences observed between the ε-values of pyrene (A) in water and (B) bound to the hydrophobic domains of C(0.15)-SNP1 in water are minor.
Figure 3. Binding of pyrene to (A) C6(0.05)-SNP1, (B) C6(0.08)-SNP1, (C) C6(0.09)-SNP1, (D) C6(0.10)-SNP1, (E) C6(0.12)-SNP1, and (F) C6(0.15)-SNP1. (○) HM-SNP1 and (●) unmodified SNP1. [Py] = 0.5 μM. Trend lines (…....) and (- - - -) are for the concentration regimes before and after the concentration (Co) at the break point, respectively. The fbound/ffree ratio shows a clear break point when plotted against the HM-SNP1 concentration.
Figure 4. Plots of (A) the binding constants (◻,■) K1 and (○,●) K2, and (B) Co as a function of the DS for (hollow) HM-SNP1 and (black) HM-SNP2. The similar values obtained for K1, K2, and Co suggest that the difference in size between HM-SNP1 and HM-SNP2 does not affect their ability to bind pyrene.
Figure 5. Schematic representation of HM-SNPs with (○) and (●) representing the hexanoyl pendants exposed to water and aggregated in the HM-SNP interior, respectively. (A) Influence of the DS on the distribution of hexanoyl pendants in the HM-SNPs, the number of hexanoyl pendants directly exposed to water remaining constant with the DS, and (B) aggregation of the HM-SNPs resulting in lower accessibility of the hexanoyl groups.
Figure 6. Binding of pyrene to (A) C6(0.08)-SNP2, (B) C6(0.10)-SNP1, and (C) C6(0.12)-SNP1. (○) HM-SNP2 and (●) unmodified SNP2. [Py] = 0.5 μM. Trend lines (…....) and (- - - -) are for the concentration regimes before and after Co, respectively. The same break point was observed in the plots of fbound/ffree-vs-[HM-SNP2] as for HM-SNP1 in Figure 3.
Figure 7. Surface tension of HM-SNP dispersions for (A) C6(0.08)-SNP1, (B) C6(0.10)-SNP1, (C) C6(0.12)-SNP1, (D) C6(0.08)-SNP2, (E) C6(0.10)-SNP2, and (F) C6(0.12)-SNP2. (○) HM-SNP and (●) unmodified SNP. Samples HM-SNP1 and HM-SNP2 exhibit a similar surface activity, despite their different sizes.
Conclusions
Acknowledgments
References
6
Simple Creams, Complex Structures
Introduction
Materials
Methods
Method of Sample Preparation
SANS
Microscopy
Rheology
Results and Discussion
Figure 1. SANS profile (obtained at 25 °C) of an aqueous cream prepared with the surfactant SDS (N1), model-fitted assuming a paracrystalline lamellar stack, with individual bilayers with the same dimensions, and a power-law description of the steep rise in scattering at low Q. Error bars on the measured data are subsumed within the plotted symbols. Cream prepared with all components protiated in 50:50 (v/v) H2O/D2O, with composition as detailed in Table 2.
Figure 2. SANS profile (obtained at 25 °C) of an aqueous cream prepared with the surfactant DTAB (N2), model-fitted assuming a paracrystalline lamellar stack, with individual bilayers with the same dimensions, and a power-law description of the steep rise in scattering at low Q. Error bars on the measured data are subsumed within the plotted symbols. Cream prepared with all components protiated in 50:50 (v/v) H2O/D2O, with composition as detailed in Table 2.
Figure 3. SANS profile (obtained at 25 °C) of an aqueous cream prepared with the surfactant hydrogenated lecithin (N3), model-fitted assuming a paracrystalline lamellar stack, with individual bilayers with the same dimensions, and a power-law description of the steep rise in scattering at low Q. Error bars on the measured data are subsumed within the plotted symbols. Cream prepared with all components protiated in 50:50 (v/v) H2O/D2O, with composition as detailed in Table 2.
Figure 4. SANS profiles measured at 25 °C for aqueous creams prepared with d70-DSPC (N4), d25-DTAB (offset at the y axis by a factor of 5 for clarity) (N6), and d25-SDS (N8) and all other (protiated) components contrast-matched to a solvent containing 97:3 (v/v) H2O/D2O (d70-DSPC and d25-DTAB) or 96:4 (v/v) H2O/D2O (d25-SDS). Arrows indicate the first three orders of Bragg reflections in the profile of the d25-DTAB cream. Cream compositions are as detailed in Table 3.
Figure 5. SANS profiles (model-fitted as detailed in Figure 1) of aqueous creams prepared in contrast-matched solvent containing (a) protiated hydrogenated lecithin or (b) deuterated d25-DTAB and a 1:1 mole ratio of hexadecanol and octadecanol as cosurfactants, with the former deuterated as d33-hexadecanol (N5 and N7, respectively). SANS measurements were made at 25 °C, and the cream compositions are as detailed in Table 3. Error bars on the measured data are subsumed within the plotted symbols.
Figure 6. SANS profiles measured at 25 °C for a fresh cream (solid line) and the corresponding dried cream (dotted line; offset at the y axis by a factor of 20 for clarity) prepared with SDS (N1), model-fitted as described in Figure 1. Error bars on the measured data are subsumed within the plotted symbols. Cream composition as detailed in Table 2.
Figure 7. SANS profiles measured at 25 °C for a fresh cream (solid line) and the corresponding dried cream (dotted line; offset at the y axis by a factor of 20 for clarity) prepared with d25-DTAB (N7), model-fitted as described in Figure 1. Error bars on the measured data are subsumed within the plotted symbols. Cream composition as detailed in Table 3.
Figure 8. SANS profiles measured at 25 °C for a fresh cream (solid line) and the corresponding dried cream (dotted line; offset at the y axis by a factor of 20 for clarity) prepared with hydrogenated lecithin (N3), model-fitted as described in Figure 1. Error bars on the measured data are subsumed within the plotted symbols. Cream composition as detailed in Table 2.
Figure 9. Photomicrographs of 1-day-old aqueous creams recorded at ~25 °C showing the dispersed oil droplets in formulations prepared with (a) SDS (N1), (b) DTAB (N2), and (c) hydrogenated lecithin (N3) under both bright-field and polarized light. Histograms of oil-droplet sizes measured from the bright-field images are fitted to a log-normal distribution. Cream compositions are as detailed in Table 2.
Figure 10. (a) Full viscosity curves recorded at 25 °C for creams based on SDS (●; N1), DTAB (■; N2), and hydrogenated lecithin (▲; N3) recorded at shear rates of 0.01–100 s–1, (b) power-law model applied to the viscosity curves over linear shear rates of 0.1–1 s–1, yielding consistency (K) indexes of 121, 79, and 30 Pa·sn and flow indexes (n) of 0.026, 0.394, and 0.312 for the SDS, DTAB, and hydrogenated lecithin creams, respectively. Cream compositions are as detailed in Table 2.
Conclusions
Figure 11. Schematic displaying the different structures formed in an aqueous cream containing DTAB (shaded) as the surfactant, a blend of the alcohols hexadecanol and octadecanol (white) as cosurfactants, liquid paraffin oil (gray dotted), and water (gray). Images are not to scale.
Figure 12. Schematic displaying the different structures formed in an aqueous cream containing SDS (shaded) as the surfactant, a blend of the alcohols hexadecanol and octadecanol (white) as cosurfactants, liquid paraffin oil (gray dotted), and water (gray). Images are not to scale.
Figure 13. Schematic displaying the different structures formed in an aqueous cream containing DSPC (shaded) as the surfactant, a blend of the alcohols hexadecanol and octadecanol (white) as cosurfactants, liquid paraffin oil (gray dotted), and water (gray). Images are not to scale.
Acknowledgments
References
7
Enzyme-Triggered Nanomaterials and Their Applications
Introduction
Figure 1. Classification of enzyme-triggered nanomaterials discussed in this chapter.
Enzyme-Triggered Self-Assembly and Morphological Transformation
Figure 2. (a) Structure of peptide used in the spatiotemporal regulation of ALP-triggered self-assembly. Images of gel at 4 °C and solution containing nanoparticles at 37 °C; transmission electron microscopy (TEM) images of the (b) nanofibers present in gel network; and (c) spherical nanoparticles. Reproduced with permission from reference 34. Copyright 2017 American Chemical Society.
Figure 3. (a) Structure of the diblock polymer and its conversion into an amphiphilic polymer when triggered by enzyme; (b) atomic force microscopy image; and (c) TEM image of the micelles after being triggered by azoreductase. Reproduced with permission from reference 35. Copyright 2014 American Chemical Society.
Figure 4. (a) Schematic of self-assembly of the chitosan-peptide conjugate and morphological transformation upon enzymatic reaction causing antibacterial action; (b) structure of the chitosan-peptide conjugate and the peptide sequences used to modify chitosan in the study. Reproduced with permission from reference 42. Copyright 2017 Wiley-VCH.
Enzyme-Triggered Disassembly or Degradation of Nanomaterials
Figure 5. (a) Structure of the block copolymer with the enzyme-responsive group at the junction of two blocks; (b) schematic showing the self-assembly and disassembly in the presence of the enzyme; and (c) images of solution of the polymer before (left) and after (right) being triggered by azoreductase. Reproduced with permission from reference 46. Copyright 2013 American Chemical Society.
Figure 6. Esterase-triggered degradation of dendritic assemblies. Reproduced with permission from reference 49. Copyright 2009 American Chemical Society.
Figure 7. Schematic of the disassembly of nanoparticles due to the surface enzymatic cleavage event of the phosphate group leading to rapid and tunable guest release behavior. Reproduced with permission from reference 50. Copyright 2018 Wiley-VCH.
Enzyme-Triggered Nonequilibrium or Dissipative Systems
Figure 8. Energy profiles of chemical systems in (a) thermodynamic equilibrium and (b) nonequilibrium conditions.
Figure 9. (a) Schematic showing the dynamic supramolecular conformation of the Zn- naphthalene diamide (NDPA) complex in the presence of ATP and adenosine diphosphate (ADP); (b) different enzymatic pathways leading to the generation and consumption of ATP and ADP. Reproduced with permission from reference 57. Copyright 2017 Wiley-VCH.
Summary and Outlook
References
8
Characterization of Colloidally Stabilized Latex Particles by Capillary Electrophoresis
Introduction
Figure 1. Schematics of three core–shell latex particles: (A) sample stabilized by both anionic and nonionic stabilizers, (B) control 1 stabilized by anionic stabilizers, and (C) control 2 stabilized by nonionic stabilizers.
Experimental Section
Emulsion Polymerization
Disk Centrifuge Photosedimentometer
CE
Sample and Buffer Preparation for Capillary Zone Electrophoresis
CE Method
AF4 with Online Multiangle Light Scattering and Differential RI Detectors
Materials and Sample Preparation
AF4 Instrument
MALS Calibration and Normalization
AF4 Method for Latex Particle Fractionation and Data Acquisition
Results and Discussion
CE Separation
Figure 2. Overlay of electropherograms of the numbered samples and AC and NC samples detected by UV at 225 ± 4 nm.
Figure 3. Electropherogram overlays of sample 3, sample 4, and sample 4 incubated with the postreaction-added nonionic stabilizers detected at (A) 225 ± 4 nm and (B) 400 ± 4 nm. (C) Expanded electropherogram of sample 3 detected at 400 ± 4 nm from 6.5 min to 7.8 min (shaded region in B).
Morphology Characterization of Bimodal Samples by CE and AF4-MALS-RI
Figure 4. CE electropherograms of sample 2 and the supernatant of sample 2 after centrifugation, detected at both 225 ± 4 nm and 400 ± 4 nm.
Figure 5. CE electropherograms of sample 3 and the supernatant of sample 3 after centrifugation, detected at both 225 ± 4 nm and 400 ± 4 nm.
Figure 6. CE electropherograms of sample 5 and the supernatant of sample 5 after centrifugation, detected at both 225 ± 4 nm and 400 ± 4 nm.
Figure 7. AF4 fractograms with right y axis representing 90 degree light scattering intensity and left y axis representing particle radius at each elution moment (x axis) from online (MALS) of sample 3, small mode of sample 3 after centrifugation, and particle size standard mixtures with diameters of 50 nm, 100 nm, 200 nm, 300 nm, 400 nm, and 700 nm and a weight ratio of 100:28:11:14:49:51.
Conclusions
References
Editor’s Biography
Ramanathan Nagarajan
Indexes
Author Index
Subject Index
C
E
G
H
M
S