Vibrational Spectroscopy at Electrified Interfaces E-Book

Andrzej Wieckowski

• Herausgeber: John Wiley & Sons
• Kategorie: Wissenschaft und neue Technologien
• Serie: The Wiley Series on Electrocatalysis and Electrochemistry
• Sprache: Englisch

Seitenzahl: 805

Veröffentlichungsjahr: 2013

Beschreibung

Reviews the latest theory, techniques, and applications Surface vibrational spectroscopy techniques probe the structure and composition of interfaces at the molecular level. Their versatility, coupled with their non-destructive nature, enables in-situ measurements of operating devices and the monitoring of interface-controlled processes under reactive conditions. Vibrational Spectroscopy at Electrified Interfaces explores new and emerging applications of Raman, infrared, and non-linear optical spectroscopy for the study of charged interfaces. The book draws from hundreds of findings reported in the literature over the past decade. It features an internationally respected team of authors and editors, all experts in the field of vibrational spectroscopy at surfaces and interfaces. Content is divided into three parts: * Part One, Nonlinear Vibrational Spectroscopy, explores properties of interfacial water, ions, and biomolecules at charged dielectric, metal oxide, and electronically conductive metal catalyst surfaces. In addition to offering plenty of practical examples, the chapters present the latest measurement and instrumental techniques. * Part Two, Raman Spectroscopy, sets forth highly sensitive approaches for the detection of biomolecules at solid-liquid interfaces as well as the use of photon depolarization strategies to elucidate molecular orientation at surfaces. * Part Three, IRRAS Spectroscopy (including PM-IRRAS), reports on wide-ranging systems--from small fuel molecules at well-defined surfaces to macromolecular complexes--that serve as the building blocks for functional interfaces in devices designed for chemical sensing and electric power generation. The Wiley Series on Electrocatalysis and Electrochemistry is dedicated to reviewing important advances in the field, exploring how these advances affect industry. The series defines what we currently know and can do with our knowledge of electrocatalysis and electrochemistry as well as forecasts where we can expect the field to be in the future.

Leseprobe

Table of Contents

Wiley Series on Electrocatalysis and Electrochemistry

Title page

Copyright page

Preface to the Wiley Series on Electrocatalysis and Electrochemistry

Foreword

Preface

Contributors

Part One: Nonlinear Vibrational Spectroscopy

Chapter 1: Water Hydrogen Bonding Dynamics at Charged Interfaces Observed with Ultrafast Nonlinear Vibrational Spectroscopy

1.1 Introduction

1.2 Experimental Methods

1.3 Linear Infrared Absorption Spectra of Water near Charged Interfaces

1.4 Population Relaxation and Orientational Relaxation

1.5 Ion–Water Hydrogen Bond Switching

1.6 Small Reverse Micelles

1.7 Spectral Diffusion Measurements of Interfacial Water Dynamics

1.8 Concluding Remarks

Acknowledgments

Chapter 2: SFG Studies of Oxide–Water Interfaces: Protonation States, Water Polar Orientations, and Comparison with Structure Results from X-Ray Scattering

2.1 Introduction: Importance of Oxide–Water Interfaces

2.2 General Features of Oxide Surface Structures and Interfacial Structures of Oxide–Water Interfaces

2.3 Macroscopic versus Microscopic Pictures of Oxide–Water Interfaces: Electrical Double Layer

2.4 Current Understanding of Oxide–Water Interfaces Excluding Information from X-ray Scattering and SFVS

2.5 X-ray Studies of Oxide–Water Interfaces

2.6 SFVS Studies of Oxide–Water Interfaces

2.7 SFVS and Surface X-ray Scattering Results on Different Oxides

2.8 Other Oxide Surfaces and Interfaces

2.9 Limitations and Difficulties

2.10 Conclusions

Acknowledgments

Chapter 3: Vibrational Sum Frequency Generation Spectroscopy of Interfacial Dynamics

3.1 Introduction

3.2 Sum Frequency Generation Spectroscopy

3.3 Time-Domain Picture and Nonresonant Suppression

3.4 Fabry–Perot Étalon

3.5 Instrumentation for SFG Spectroscopy

3.6 Interfacial Dynamics via Ultrafast Temperature Jump

3.7 Lithium Batteries

3.8 Summary and Outlook for Future Work

Acknowledgments

Chapter 4: Spectroscopy of Electrified Interfaces with Broadband Sum Frequency Generation: From Electrocatalysis to Protein Foams

4.1 Introduction

4.2 Principles of Vibrational Sum Frequency Generation and Experimental Details

4.3 Specific Adsorption of (Bi)sulfate Anions on Pt(111) Surfaces Studied with Broadband SFG

4.4 Broadband SFG Studies of Electrochemical Reactions: Electrocatalysis of Ethanol

4.5 Sum Frequency Generation of Proteins at Electrified Air–Water Interfaces

Acknowledgments

Part Two: Raman Spectroscopy

Chapter 5: Surface-Enhanced Resonance Raman Scattering (SERRS) Studies of Electron-Transfer Redox-Active Protein Attached to Thiol-Modified Metal: Case of Cytochrome c

5.1 Principles and Methodology of SERS and EC-SERRS Techniques

5.2 Attachment of Cytochrome c to Alkanethiol-Coated Electrode Prior to ET Studies

5.3 EC–SERRS Studies of Cyt c Attached to Alkenethiol-Coated Electrodes

5.4 Summary

Chapter 6: Depolarization of Surface-Enhanced Raman Scattering Photons from a Small Number of Molecules on Metal Surfaces

6.1 Introduction

6.2 Brief History of Polarized SERS Measurements

6.3 Method for SERS Polarization Measurement

6.4 Depolarization Behavior of SERS Photons

6.5 Conclusion

Acknowledgments

Part Three: IRRAS Spectroscopy (Including PM–IRRAS)

Chapter 7: DFT and In Situ Infrared Studies on Adsorption and Oxidation of Glycine, l-Alanine, and l-Serine on Gold Electrodes

7.1 Introduction

7.2 Model and Computational Details

7.3 Experimental Section

7.4 Results and Discussion

7.5 Conclusions

Acknowledgments

Chapter 8: Composition, Structure, and Reaction Dynamics at Electrode–Electrolyte Interfaces Using Infrared Spectroscopy

8.1 Introduction

8.2 Reflectance Spectroscopy

8.3 External Reflectance Infrared Spectroscopy for Structural Studies of Chemisorbed Adlayers

8.4 Time–Resolved ATR–Seiras for Study of Mechanism of Electrocatalytic Reactions

8.5 ATR–Seiras for Study of Enzymes Immobilized on Metal Electrodes

8.6 Conclusions

Acknowledgments

Chapter 9: Vibrational Stark Effect at Halide Precovered Cu(100) Electrodes

9.1 Introduction

9.2 Experimental

9.3 Results

9.4 Conclusion

Acknowledgments

Chapter 10: Vibrational Spectroscopy of the Ionomer–Catalyst Interface

10.1 Introduction

10.2 Vibrational Spectroscopy of Bulk Nafion

10.3 State of Hydration of Nafion: FTIR and DFT Analysis

10.4 Operando Spectroscopy of Ionomer–Pt Interface

10.5 Polarization-Modulated IR Reflection ABsorption Spectroscopy of Nafion–Pt Interfaces

10.6 Future Work

Chapter 11: In Situ PM–IRRAS Studies of Biomimetic Membranes Supported at Gold Electrode Surfaces

11.1 Introduction

11.2 What Is PM–IRRAS?

11.3 Experimental

11.4 Electrochemical Studies of Gold-Electrode-Supported Bilayers

11.5 IR Studies of Electrode-Supported Membranes

11.6 Preparation of Biomimetic Membranes

11.7 PM–IRRAS Studies of One-Component Biomimetic Membranes

11.8 Two-Component Bilayers

11.9 Three-Component Bilayers

11.10 Bilayers with Incorporated Peptides

11.11 PM–IRRAS Studies of Floating Bilayer Lipid Membranes

11.12 Summary and Conclusions

Acknowledgments

Index

WILEY SERIES ON ELECTROCATALYSIS AND ELECTROCHEMISTRY

Andrzej Wieckowski, Series Editor

Synthetic Diamond Films: Preparation, Electrochemistry, Characterization and Applications, Edited by Enric Brillas and Carlos Alberto Martínez-Huitle

Fuel Cell Catalysis: A Surface Science Approach, Edited by Marc T. M. Koper

Electrochemistry of Functional Supramolecular Systems, Margherita Venturi, Paola Ceroni, And Alberto Credi

Catalysis in Electrochemistry: From Fundamentals to Strategies for Fuel Cell Development, Elizabeth Santos and Wolfgang Schmickler

Fuel Cell Science: Theory, Fundamentals, and Biocatalysis, Andrzej Wieckowski and Jens Norskov

Copyright © 2013 by John Wiley & Sons, Inc. All rights reserved

Published by John Wiley & Sons, Inc., Hoboken, New Jersey

Published simultaneously in Canada

No part of this publication may be reproduced, stored in a retrieval system, or transmitted in any form or by any means, electronic, mechanical, photocopying, recording, scanning, or otherwise, except as permitted under Section 107 or 108 of the 1976 United States Copyright Act, without either the prior written permission of the Publisher, or authorization through payment of the appropriate per-copy fee to the Copyright Clearance Center, Inc., 222 Rosewood Drive, Danvers, MA 01923, [phone], fax [phone], or on the web at www.copyright.com. Requests to the Publisher for permission should be addressed to the Permissions Department, John Wiley & Sons, Inc., Ill River Street, Hoboken, NJ 07030, [phone], fax [phone], or online at http://www.wiley.com/go/permissions.

Limit of Liability/Disclaimer of Warranty: While the publisher and author have used their best efforts in preparing this book, they make no representations or warranties with respect to the accuracy or completeness of the contents of this book and specifically disclaim any implied warranties of merchantability or fitness for a particular purpose. No warranty may be created or extended by sales representatives or written sales materials. The advice and strategies contained herein may not be suitable for your situation. You should consult with a professional where appropriate. Neither the publisher nor author shall be liable for any loss of profit or any other commercial damages, including but not limited to special, incidental, consequential, or other damages.

For general information on our other products and services or for technical support, please contact our Customer Care Department within the United States at [phone], outside the United States at [phone] or fax [phone].

Wiley also publishes its books in a variety of electronic formats. Some content that appears in print may not be available in electronic formats. For more information about Wiley products, visit our web site at www.wiley.com.

Library of Congress Cataloging-in-Publication Data is available.

9781118157176

Preface to the Wiley Series on Electrocatalysis and Electrochemistry

This series covers recent advances in electrocatalysis and electrochemistry and depicts prospects for their contribution into the present and future of the industrial world. It aims to illustrate the transition of electrochemical sciences from its beginnings as a solid chapter of physical chemistry (covering mainly electron transfer reactions, concepts of electrode potentials, and structure of electrical double layer) to the field in which electrochemical reactivity is shown as a unique chapter of heterogeneous catalysis, is supported by high-level theory, connects to other areas of science, and includes focus on electrode surface structure, reaction environment, and interfacial spectroscopy.

The scope of this series ranges from electrocatalysis (practice, theory, relevance to fuel cell science and technology) to electrochemical charge transfer reactions, biocatalysis, and photoelectrochemistry. While individual volumes may appear quite diverse, the series promises updated and overall synergistic reports providing insights to help further our understanding of the properties of electrified solid–liquid systems. Readers of the series will also find strong reference to theoretical approaches for predicting electrocatalytic reactivity by such high-level theories as density functional theory. Beyond the theoretical perspective, further vehicles for growth are such significant topics such as energy storage, syntheses of catalytic materials via rational design, nanometer-scale technologies, prospects in electrosynthesis, new instrumentation, and surface modifications. In this context, the reader will notice that new methods being developed for one field may be readily adapted for application in another.

Electrochemistry and electrocatalysis have both benefited from numerous monographs and review articles due to their depth, complexity, and relevance to the practical world. The Wiley Series on Electrocatalysis and Electrochemistry is dedicated to present the current activity by focusing each volume on a specific topic that is timely and promising in terms of its potential toward useful science and technology. The chapters in these volumes will also demonstrate the connection of electrochemistry to other disciplines beyond chemistry and chemical engineering, such as physics, quantum mechanics, surface science, and biology. The integral goal is to offer a broad-based analysis of the total development of the fields. The progress of the series will provide a global definition of what electrocatalysis and electrochemistry are now, and will contain projections about how these fields will further evolve in time. The purpose is twofold, to provide a modern reference for graduate instruction and for active researchers in the two disciplines, as well as to document that electrocatalysis and electrochemistry are dynamic fields that are expanding rapidly, and are likewise rapidly changing in their scientific profiles and potential.

Creation of each volume required the editors involvement, vision, enthusiasm, and time. The Series Editor thanks each Volume Editor who graciously accepted his invitation. Special thanks go to Ms. Anita Lekhwani, the Series Acquisitions Editor, who extended the invitation to edit this series to me and has been a wonderful help in its assembling process.

Andrzej Wieckowski

Series Editor

Foreword

Despite extensive efforts, the electrochemical interface, where central processes in electrochemical reactions occur, had long been a black box until researchers started to shed light on it in the 1960s. At the beginning, the light used was mostly that in the visible region, but information obtainable by visible light was very limited, and the use of vibrational spectroscopy, which can provide detailed information on molecules, was strongly desired. In 1974, an innovation was made in spectroelectrochemistry by the application of Raman spectroscopy. It is well known that the Raman study of pyridine adsorbed on silver electrodes led to the discovery of surface-enhanced Raman scattering (SERS). The application of infrared reflection absorption spectroscopy (IRAS) to electrochemical interfaces in 1980 also was a great achievement. Surface-enhanced infrared absorption (SEIRA), an effect similar to SERS, was discovered in the same year, and the first observation of sum frequency generation (SFG) from monolayers on solid surfaces was made in 1986, although their applications to electrochemical interfaces were somewhat delayed. During the last four decades, these surface vibrational spectroscopy techniques have been advanced greatly owing to the improvement in instrumentation and the development of experimental techniques, which are still further developing year by year.

Vibrational spectroscopy is a powerful tool to identify molecules and to study their structures and reactions, as it is often mentioned that vibrational spectra are letters from molecules. It is also the case at surfaces and interfaces. It provides us information on adsorbed structures and orientations of molecules. Spectra are sensitive to changes in the environment at interfaces, from which we can obtain deeper insight into chemistry and physics at the interfaces. In situ, time-resolved monitoring of reactions taking place at surfaces and interfaces is also possible. Owing to these advantages, they have gained wide application, from fundamental electrochemistry to many other related fields of science and technology, including surface science, heterogeneous electrocatalysis, energy conversion, biochemistry, nanotechnology, and sensors. However, each technique has strong and weak points. Difficulty in interpretation of the obtained spectra is another problem. For appropriate use of the techniques and for correct interpretation of spectra, sufficient fundamental understanding of the techniques is required.

This book, which features all the aforementioned four surface vibrational spectroscopy techniques and their applications to recent research topics, will provide fundamental information for nonspecialists and an up-to-date account of recent advances in this field for specialists.

Masatoshi Osawa

Catalysis Research Center, Hokkaido University, Sapporo, Japan

Preface

Electrified interfaces play an important role in many phenomena. Electric fields that develop at junctions between different phases can align molecules and ions into configurations that greatly influence the physical and chemical nature of the interface. The molecular structure of charged interfaces impacts many practical processes, including energy conversion in batteries, solar cells, and fuel cells, corrosion at solid surfaces, chemical reactions over oxide particles in the earth and atmosphere, biochemical transformations, signal transduction in chemical sensors, and heterogeneous catalytic reactions, to name a few.

Surface vibrational spectroscopy techniques probe the structure and composition of interfaces at a molecular level. Their versatility and typically nondestructive nature often enable in situ measurements of operating devices and monitoring of interface-controlled processes under reactive conditions. This book highlights modern applications of Raman, infrared, and nonlinear optical spectroscopy in the study of charged interfaces.

Early chapters in the book provide a glimpse into the breadth of systems that can be investigated through the use of nonlinear optical techniques. Properties of interfacial water, ions, and biomolecules at charged dielectric, metal oxide, and electronically conductive metal catalyst surfaces, as probed by nonlinear optical techniques, are discussed in Part I. In addition to examples of practical experimental interest, the chapters guide readers to the latest in measurement and instrumental techniques. Part II includes coverage of Raman spectroscopy from the standpoint of sensitive approaches for detection of biomolecules at solid–liquid interfaces and the use of photon depolarization strategies to elucidate molecular orientation at surfaces. Part III reports on wide-ranging systems from small fuel molecules at well-defined surfaces to macromolecular complexes as building blocks of functional interfaces in devices that have applications in chemical sensing and electric power generation. These interfaces are amenable for infrared spectroscopy due to versatile sampling methods, that is, specular and diffuse reflectance, polarization–modulation, and total internal reflection modes.

Andrzej Wieckowski

Carol Korzeniewski

Björn Braunschweig

Contributors

Christopher M. Berg

School of Chemical Sciences

University of Illinois

Urbana, Illinois

Björn Braunschweig

Institute of Particle Technology (LFG)

Erlangen, Germany

Christa L. Brosseau

Department of Chemistry

Saint Mary's University, Halifax

Nova Scotia, Canada

Angel Cuesta

Instituto de Química Física “Rocasolano”

CSIC

Madrid, Spain

Dana D. Dlott

School of Chemical Sciences

University of Illinois

Urbana, Illinois

Jonathan Doan

Department of Chemistry and Chemical Biology

Northeastern University

Boston, Massachusetts

Kathrin Engelhardt

Institute of Particle Technology (LFG)

Erlangen, Germany

Michael D. Fayer

Department of Chemistry

Stanford University

Stanford, California

Juan M. Feliu

Instituto de Electroquímica

Universidad de Alicante

Alicante, Spain

Emily E. Fenn

Department of Chemistry

Stanford University

Stanford, California

Ian Kendrick

Department of Chemistry and Chemical Biology

Northeastern University

Boston, Massachusetts

Carol Korzeniewski

Department of Chemistry

Texas Tech University

Lubbock, Texas

Agata Królikowska

Department of Chemistry Warsaw University

Warsaw, Poland

Robert B. Kutz

Champaign, Illinois

Annia H. Kycia

Department of Chemistry

University of Guelph,

Guelph, Ontario, Canada

Jacek Lipkowski

Department of Chemistry

University of Guelph

Guelph, Ontario, Canada

Prabuddha Mukherjee

Urbana, Illinois

Kei Murakoshi

Department of Chemistry Faculty of Science Hokkaido University

Sapporo, Japan

Hideki Nabika

Department of Material and Biological Chemistry, Faculty of Science Yamagata University

Yamagata, Japan

Fumika Nagasawa

Department of Chemistry Faculty of Science Hokkaido University

Sapporo, Japan

José Manuel Orts

Instituto de Electroquímica

Universidad de Alicante

Alicante, Spain

Wolfgang Peukert

Institute of Particle Technology (LFG)

Erlangen, Germany

Duc Thanh Pham

Institute of Physical Chemistry

University of Bonn

Bonn, Germany

Antonio Rodes

Instituto de Electroquímica

Universidad de Alicante

Alicante, Spain

Melanie Röefzaad

Institute of Physical and Theoretical Chemistry

University of Bonn

Bonn, Germany

Armin Rumpel

SwissOptic AG

Heerbrugg, Switzerland

Andrea P. Sandoval

Departamento de Quimica, Facultad de Ciencias

Universidad Nacional de Colombia

Bogotá, Colombia

Y. Ron Shen

Physics Department

University of California, Berkeley

Berkeley, California

Eugene S. Smotkin

Department of Chemistry and Chemical Biology

Northeastern University

Boston, Massachusetts

ZhangFei Su

Department of Chemistry

University of Guelph

Guelph, Ontario, Canada

Mai Takase

Catalysis Research Center Hokkaido University

Sapporo, Japan

Klaus Wandelt

Institute of Physical and Theortetical Chemistry

University of Bonn

Bonn, Germany, and

Institute of Experimental Physics

University of Wroclaw

Wroclaw, Poland

Glenn A. Waychunas

Geochemistry Department Earth Sciences Division

Lawrence Berkeley National Laboratory Berkeley, California

Andrzej Wieckowski

Department of Chemistry

University of Illinois at Urbana Champaign Urbana, Illinois

Part One

Nonlinear Vibrational Spectroscopy

Chapter 1

Water Hydrogen Bonding Dynamics at Charged Interfaces Observed with Ultrafast Nonlinear Vibrational Spectroscopy

Emily E. Fenn and Michael D. Fayer

Department of Chemistry, Stanford University, Stanford, California

1.1 Introduction

The question of how charged species affect water structure and dynamics is relevant to many applications in chemistry, biology, geology, and industry. Biological systems are often crowded aqueous environments filled with proteins, membranes, vesicles, and other structures that often rely on the presence of ions for stability and proper functioning [1–6]. The ion–water interface is critical for ion exchange resins [7, 8], heterogeneous catalysis [9–11], electrochemistry [12], as well as processes involving mineral dissolution [13, 14] and ion adsorption [15, 16]. Because the behavior of water in the presence of ions impacts a wide range of technical and scientific fields, a great deal of literature over the years has been dedicated to studying the aqueous solvation of ions and the properties of water at charged interfaces. Studies that have examined ion–water interfaces have employed x-ray and neutron diffraction [17–19], Raman spectroscopy [20], ultrafast infrared spectroscopy [21–26], Fourier transform infrared (FTIR) spectroscopy [20, 27], and other spectroscopic techniques [15, 28, 29]. Theoretical models [30, 31], molecular dynamics (MD) simulations [32–35], and Monte Carlo (MC) calculations [20] have also been employed. While simulations can provide some insight into the underlying dynamics, most experimental techniques only provide steady-state data. Here we utilize ultrafast infrared spectroscopy to examine the hydrogen bonding dynamics of water at several types of charged and uncharged interfaces.

Ultrafast infrared spectroscopy has been shown to be a powerful technique for elucidating dynamics in water–ion systems [21–26], other hydrogen bonding systems [36–43], protein environments [44–52], and systems that undergo chemical exchange [25, 53–57]. Here, we apply ultrafast infrared pump–probe and two-dimensional infrared (2D IR) vibrational echo spectroscopic techniques to examine the dynamics of water when it is confined in nanoscopic environments and interacting with interfaces. The question is whether the nature of confinement or the chemical composition of the interface most significantly influences the dynamics. To explore this question, the dynamics of water at charged and neutral interfaces in reverse micelles are compared. In addition, water in ionic solutions is investigated. Some water molecules are hydrogen bonded to ions, while others are hydrogen bonded to water molecules. These are in equilibrium, with water molecules bound to ions switching and becoming bound to water molecules, and vice versa. Using 2D IR chemical exchange spectroscopy, we determine the exchange time required for a water hydroxyl initially hydrogen bonded to an anion to switch to being hydrogen bonded to another water molecule.

Reverse micelles consist of a water pool surrounded by a layer of surfactant molecules and are often used as model systems for confined environments. The surfactant molecules are terminated by a hydrophilic head group that can be either charged or neutral. These hydrophilic head groups face in toward the water pool while the alkyl (hydrophobic) tails of the surfactant are suspended in a nonpolar organic phase. A schematic of a reverse micelle utilizing the surfactant Aerosol-OT, or AOT [sodium bis(2-ethylhexyl) sulfosuccinate], is shown in Figure 1.1. The AOT surfactant (Fig. 1.2) forms spherical monodispersed reverse micelles that have been well characterized. The size of the AOT reverse micelles can be easily controlled by varying the amounts of starting materials according to the w0 parameter: w0 = [H2O]/[surfactant] [58–60]. AOT can yield sizes of w0 = 0 (essentially dry reverse micelles) all the way up to w0 = 60, which has a water pool diameter of 28 nm and contains ∼350,000 water molecules [61]. Isooctane is a common solvent used as the nonpolar phase of AOT reverse micelle systems, but other solvents such as carbon tetrachloride, cyclohexane, and benzene can also be used with minimal changes in water pool size for a given w0 [62]. A recent study has shown that the identity of the nonpolar phase has no effect on the water pool dynamics [63].

Figure 1.1 Illustration of the reverse micelle interior. The bulk water core is surrounded by a layer of interfacial water. The total water pool diameter is denoted by d. The hydrophilic AOT head groups face in toward the water pool while the alkyl tails are suspended in the organic phase. The sodium counterions are dispersed in the water pool, but they generally reside close to the head group interface.

Figure 1.2 Molecular structures for AOT and Igepal CO-520. AOT (top) is terminated by a charged sulfonate head group with a sodium counterion while Igepal (bottom) has a neutral hydroxyl head group.

As shown in Figure 1.2, AOT has a sulfonate head group with a sodium counterion. The head group region of the reverse micelle therefore creates a charged interface that surrounds the water pool. The sodium ions will generally reside in a region close to the interface. Figure 1.1 illustrates the regions of a reverse micelle. When the total water pool diameter, d, is sufficiently large (≥4.6 nm) the reverse micelle can support a core of water with bulklike properties. Below we will discuss how far perturbations from the charged sulfonate interfacial region extend into the water pool and what happens to the water dynamics as the size of the water pool changes in size. The chemical identity of the surfactant layer can be changed by using a neutral surfactant molecule called Igepal CO-520 (Fig. 1.2). Igepal is terminated with neutral hydroxyl head groups, so the interfacial water molecules will be exposed to a very different chemical surface compared to the AOT reverse micelle system. To what extent changes in surfactant identity, particularly charged versus neutral head group regions, and reverse micelle size affect water dynamics will be described.

Water dynamics are investigated through the processes of orientational relaxation, spectral diffusion, and vibrational relaxation, which can be measured with ultrafast infrared vibrational spectroscopy. These observables report on how the hydrogen bond network of water evolves and rearranges over time. The hydroxyl stretch of water is monitored during the experiments and is used as a reporter for hydrogen bond dynamics. During vibrational relaxation, vibrational energy dissipates by transferring into a combination of low-frequency modes, such as torsions and bath modes [64, 65]. Energy must be conserved during this process. Certain pathways that facilitate vibrational relaxation in one system may or not be present in a different system. Thus, vibrational relaxation is extremely sensitive to local environments. Orientational relaxation measures how quickly water molecules reorient by monitoring the direction of the transition dipole of the hydroxyl stretch. Molecular reorientation is involved in water hydrogen bond exchange, which leads to global hydrogen bond network reorganization [66, 67]. Bulk water consists of an extended network of hydrogen bonds that are continually rearranging and exchanging with one another. According to the theory of Laage and Hynes, water molecules exchange hydrogen bonds via a jump reorientation mechanism that involves concerted motions of water molecules in the first and second solvation shells [66, 67]. The mechanism proceeds when a molecule in the second solvation shell of another water molecule moves in toward the first solvation shell. In order to swap hydrogen bonds with the approaching water from the second solvation shell, a water molecule must pass through a five-coordinate transition state and then undergo a large-amplitude rotational motion (or “jump”). The jump allows it to switch one of its hydrogen bonds to the approaching water molecule. These large-amplitude jumps change the orientation of the transition dipole. Solutes and interfaces (such as the surfactant shell of the reverse micelles) can disrupt the jump reorientation mechanism, thus slowing down the process of reorientation [23, 68–72]. Both vibrational and orientational relaxation can be measured with ultrafast infrared pump–probe spectroscopy.

Ultrafast 2D IR vibrational echo spectroscopy is used to measure spectral diffusion of the water hydroxyl stretch. The linear infrared absorption spectrum of the hydroxyl stretch is very broad due to a large distribution in the lengths and strengths of hydrogen bonds. At the beginning of the 2D IR experiment, a hydroxyl will vibrate at a certain frequency, but due to dynamic structural evolution of the system, that frequency will change over time. This process of frequency evolution is known as spectral diffusion and reports on how quickly water molecules sample different structural environments.

In addition, 2D IR vibrational echo chemical exchange spectroscopy is used to examine how quickly a water hydroxyl bound to an anion will switch to being hydrogen bonded to a neighboring water hydroxyl. This process is illustrated schematically in Figure 1.3. A model system for studying water–ion exchange is a solution of sodium tetrafluoroborate (NaBF4) in water because the linear IR absorption spectrum of the solution yields two resolved peaks corresponding to waters interacting with other waters and waters interacting with the tetrafluoroborate anions. It is found that the ion–water hydrogen bond switching time is ∼7 ps [25]. This switching time has implications when treating the orientational relaxation dynamics of water molecules inside reverse micelles made of charged and neutral surfactants [71]. Together, these ultrafast infrared experiments involving water in reverse micelles and water–ion chemical exchange construct a dynamic picture of the behavior of water molecules at charged interfaces and interacting with ions.

Figure 1.3 Representation of solvation shell exchange. The time it takes for a water hydroxyl initially hydrogen bonded to an anion (left side) to switch to being hydrogen bonded to another water hydroxyl (right side) can be measured with 2D IR vibrational echo spectroscopy.

1.2 Experimental Methods

1.2.1 Sample Preparation

Carbon tetrachloride (CCl4), cyclohexane, isooctane, H2O, D2O, AOT, Igepal CO-520, and NaBF4 were used as received. Stock solutions of 0.5 M AOT were prepared in CCl4, cyclohexane, and isooctane. A variety of solvents are necessary due to certain experimental considerations that will be discussed below. A 0.3-M stock solution of Igepal CO-520 was prepared in cyclohexane. The residual water contents of the stock solutions were measured via Karl Fischer titration. The reverse micelle samples were prepared by mass by adding appropriate amounts of a solution of 5% HOD (water with one hydrogen exchanged with deuterium) in H2O to measured quantities of the AOT or Igepal stock solutions to obtain the desired w0. In the ultrafast experiments, the OD stretch of 5% HOD in H2O is probed because it not only provides an isolated stretching mode to interrogate but also prevents vibrational excitation transfer processes from artificially causing decay of the orientational correlation function and observables related to spectral diffusion [73, 74]. MD simulations demonstrate that a dilute amount of HOD does not perturb the structure and properties of H2O and that the OD stretch reports on the dynamics of water [75].

For the pump–probe experiments involving large reverse micelles presented here, the 0.5-M stock solution of AOT in isooctane was used to make samples of w0 = 10, 16.5, 25, 37, and 46 (diameters of 4.0, 5.8, 9, 17, and 20 nm, respectively). It has been found that the combination of AOT and isooctane causes distortions in vibrational echo experiments, so for vibrational echo experiments on small reverse micelles the 0.5-M AOT/CCl4 stock solution was used to make w0 = 2, 4, and 7.5 (diameters of 1.7, 2.3, and 3.3 nm, respectively) [63]. CCl4 cannot support reverse micelles larger than w0 ∼10 [61], so the 0.5-M AOT/cyclohexane stock solution was used to make w0 = 12 and 16.5 (diameters of 4.6 and 5.8 nm, respectively) for vibrational echo experiments on larger reverse micelles. Neither CCl4 nor cyclohexane cause distortions in the vibrational echo experiments. Cyclohexane can only reliably make reverse micelles of w0 < 20 [76, 77], so the vibrational echo studies are limited to the lower bound of large sizes. It will be shown that both w0 = 12 and 16.5 have well-defined bulk water cores and interfacial water regions, so analysis of these sizes should provide insight into the behaviors of water molecules in the even larger sizes.

To compare the effects of the chemical composition of the interface, Igepal reverse micelles with w0 = 12 and 20 were also made. Like AOT, Igepal also makes monodispersed spherical reverse micelles [78]. The AOT and Igepal surfactants have different aggregation numbers, thus yielding different sizes for the same w0 values. The w0 = 12 Igepal reverse micelles have the same 5.8-nm diameter as AOT w0 = 16.5 while the w0 = 20 reverse micelles have the same 9-nm diameter as AOT w0 = 25. AOT lamellar structures (sheets of AOT surfactants with water between them) were prepared by adding water to dry AOT to produce samples with various water-to-surfactant ratios, λ. These lamellar samples allow us to examine the effects of confining geometry on the water dynamics (spherical confinement versus confinement within layers).

The experimental samples are contained between two calcium fluoride windows that are separated by a Teflon spacer. The thickness of the Teflon spacer is chosen such that the optical density of the OD stretch region is ∼0.1 for the vibrational echo experiments and ∼0.5–0.7 for the pump–probe experiments.

For the 2D IR chemical exchange experiments, a 5.5-M solution of NaBF4 in water was used. Again, the water component consisted of 5% HOD in H2O for the same reasons outlined above. The 5.5-M concentration corresponds to a system with seven water molecules per NaBF4 molecule (n = 7). For linear IR absorption measurements, additional samples of 1.7 M, 3.1 M, and 4.3 M of NaBF4 in water were made, corresponding to n = 30, 15, and 10, respectively.

1.2.2 2D IR Vibrational Echo Spectroscopy

The laser system used to generate the infrared light that excites the OD hydroxyl stretch consists of a Ti–sapphire oscillator that seeds a regenerative amplifier. The output of the regenerative amplifier pumps an optical parametric amplifier that generates near-infrared wavelengths that are difference frequency mixed in a AgGaS2 crystal. The resulting mid-IR pulses are centered at ∼4 μm (2500 cm−1) but can be tuned to the peak of the absorption spectrum for a given sample (e.g., 2565 cm−1 for w0 = 2 AOT reverse micelle). The generated mid-IR beam enters a 2D IR vibrational spectrometer that can be readily converted into a pump–probe setup.

The pump–probe and 2D IR vibrational echo techniques presented here are noncollinear four-wave mixing experiments [79, 80]. In these experiments, three field–matter interactions between the sample and the incident ultrafast laser pulses create a third-order macroscopic polarization that emits a signal electric field. Depending upon the type of experiment, the signal electric field carries different types of information. As discussed in the introduction, the signal from a pump–probe experiment allows one to extract vibrational lifetimes and orientational relaxation parameters for the OD stretch of HOD in H2O. The vibrational lifetime and orientational observables are sensitive to local structural and chemical environments. 2D IR vibrational echo spectroscopy is a sophisticated technique that observes how vibrational chromophores in a system evolve in frequency over time via chemical exchange, spectral diffusion, coherence transfer, or other processes. 2D IR spectroscopy can monitor these frequency changes by manipulating the quantum pathways by which the system evolves.

The experimental pulse sequence for the 2D IR vibrational echo experiments is shown in Figure 1.4a. The pump–probe experiment (Fig. 1.4b) is similar in several ways to the 2D IR experiment, but it differs in the number of input beams and the manner by which the signal is detected. In the 2D IR experiment, three time-ordered beams interact with the sample (Fig. 1.5). The first pulse induces a coherence state between the ground (0) and first excited (1) vibrational levels. The vibrations are initially in phase, but inhomogeneous broadening of the absorption line and structural fluctuations cause the phase relationships to decay. After a period of time, τ, a second pulse impinges on the sample and creates a population state in either the 0 or 1 vibrational levels. A time period Tw elapses (the waiting time) before the third pulse reaches the sample and induces a second coherence state, partially restoring the phase relationships between the vibrational chromophores. This rephasing process causes the vibrational echo signal electric field to emit at a time t ≤ τ. The vibrational echo signal propagates in the phase-matched direction according to ksig = −k1 + k2 + k3, as denoted by the BOXCARS geometry of the input beams shown in Figure 1.5. The vibrational echo signal is temporally and spatially overlapped with a local oscillator (LO) pulse for heterodyned detection. The LO is another IR pulse identical to the excitation pulses but lower in amplitude and fixed in time. The combined vibrational echo and LO beam is frequency dispersed by a monochromator, and the heterodyned signal is detected on a 32-pixel mercury–cadmium–telluride (MCT) array detector. If we denote the vibrational echo signal as S and LO signal as L, then the quantity |S + L|2 = S2 + 2LS + L2 is measured on the detector. Assume S2 is negligible and can be ignored and L2 is a constant signal and can be subtracted. The 2LS cross term represents the heterodyned signal of interest.

Figure 1.4 Pulse sequences implemented for (a) 2D IR vibrational echo spectroscopy and (b) pump–probe spectroscopy. Both techniques are noncollinear four-wave mixing experiments.

Figure 1.5 Experimental setup for the 2D IR vibrational echo experiment. Three excitation beams in a BOXCARS geometry cross in the sample to produce the vibrational echo signal in the phase-matched direction denoted by the wave vector ksig. The vibrational echo signal combines with a local oscillator beam for heterodyned detection. The heterodyned signal is detected by a 32-pixel MCT array detector.

Ultimately, the 2D IR experiment obtains frequency correlation plots (2D spectra) for the second coherence period (known as the detection time period) and the first coherence period (known as the evolution time period). Each frequency axis of the correlation plot arises from Fourier transformation of each coherence period. The Fourier transform along the second coherence period is performed experimentally by the monochromator, yielding the vertical “ωm” axis while the Fourier transform along the first coherence period is obtained numerically during data processing, yielding the horizontal “ωτ” axis. These axes are clearly marked in Figures 1.6a, b, which show correlation spectra for bulk water (5% HOD in H2O). During the experiment, τ is scanned for a series of fixed Tw values. As τ is scanned, the echo signal field moves in time relative to the fixed LO. The echo field goes in and out of phase with the LO field producing an interferogram. Mixing the vibrational echo signal with the LO allows interferograms to be recorded, thus providing the necessary phase information for Fourier transformation.

Figure 1.6 CLS procedure: (a) bulk water 2D spectrum at Tw = 0.2 ps, showing elongation along the diagonal and (b) bulk water spectrum at Tw = 2 ps. Spectral diffusion has mostly completed, so the spectra become more circular. The white solid lines show the direction of cuts during the CLS procedure. The white dots indicate the peak positions through the slices (centerline data). The slope is found through each set of peak positions at each Tw to obtain a plot of slope versus Tw, as shown in panel (c). The CLS curve is equal to the normalized FFCF, which is shown as the black line through the CLS data.

Qualitatively, a 2D IR vibrational echo experiment works in the following manner. The first laser pulse “labels” the initial structures of the species by establishing their initial frequencies, ωτ. The second pulse ends the first time period τ and starts the reaction time period Tw during which the labeled species undergo structural evolution. For example, the local hydrogen bond network rearranges. This ends the population period of length Tw and begins a third period of length ≤τ, which ends with the emission of the vibrational echo pulse of frequency ωm, which is the signal in the experiment. The vibrational echo signal reads out information about the final structures of all labeled species by their frequencies, ωm. During the period Tw between pulses 2 and 3, the system's structural evolution occurs. The structural evolution and associated frequency changes as Tw is increased cause new off-diagonal peaks to grow in a chemical exchange experiment or the shape of the 2D spectrum to change in a spectral diffusion experiment. The growth of the off-diagonal peaks or the change in the 2D band shapes in the 2D IR spectra with increasing Tw provides the dynamical information.

The 2D IR experiments require precise timing between the excitation pulses as well as phase stability during the coherence periods. Computer-controlled precision delay lines (Aerotech ANT-50L) manipulate the delay between the excitation pulses, and a three-pulse cross-correlation measurement is used to check the timing and correct for drifts between the three excitation pulses [63]. The vibrational echo experiments are also sensitive to linear chirp in the mid-IR pulse. Linear chirp is corrected by proper insertion of materials (calcium fluoride and germanium) with opposite signs of the group velocity dispersion (GVD) [82, 83].

The 2D IR experiments presented here are used to determine the frequency–frequency correlation function (FFCF) of water molecules (OD stretch), which is a measure of spectral diffusion. The FFCF can be determined via the centerline slope (CLS) method [84, 85]. Figure 1.6 illustrates this process. In the CLS technique, the 2D IR spectrum is sliced parallel to the vertical ωm axis over a range of frequencies surrounding the center of the 2D IR spectrum. In Figures 1.6a,b, the solid white lines show the direction of slicing and the bounds over which the slices are taken. For water systems (as shown in the figure), the range is typically ±30–40 cm−1 around the measured center frequency of each 2D IR spectrum. Each slice intercepts a frequency on the horizontal ωτ axis. The slices are fit to Gaussian line shape functions to determine the peak position of each slice. Only the peak positions are required. The peak positions (white dots) are plotted versus their corresponding ωτ frequencies, and the slope of the resulting line is calculated. This process is repeated for all Tw values so that a plot of slope versus Tw is obtained (Fig. 1.6c). Generally, the 2D plots show elongated spectra at early Tw, as shown by Figure 1.6a. By 2 ps (Fig. 1.6b), the spectra show a more circular shape because most of the water hydroxyl environments in the H2O inhomogeneous line shape have been sampled by the OD vibrational chromophore, indicating that spectral diffusion is nearly complete.

It has been demonstrated theoretically that the CLS curve of slope versus Tw is equal to the Tw-dependent portion of the normalized FFCF [84, 85]. The FFCF adopts a functional form that contains both homogeneous (motionally narrowed) and inhomogeneous components,

(1.1)

where 〈δω10(t)δω10(0)〉 is the correlation function for the fluctuating 0–1 transition frequency, and δω10(t) = 〈ω10〉 − ω10(t). The summation term in Eq. (1.1) contains the processes that sample the inhomogeneous distribution of frequencies through structural evolution of the system. The Δi terms are frequency fluctuation amplitudes, and the τi are their associated time constants. The time constants describe different time scales that contribute to spectral diffusion. The magnitude of each Δi term gives the contribution to the absorption line shape from processes occurring on each time scale.

The homogeneous dephasing time T2 is given by

(1.2)

where is the pure dephasing time, T1 is the vibrational lifetime, and τor is the orientational relaxation time constant. The first term of Eq. (1.1) is known as the pure dephasing component. In the condensed phase, homogeneous broadening can arise from fast solvent motions [86]. The dynamics that give rise to pure dephasing occur on a very fast time scale such that the product Δτ < 1, meaning that this portion of the total absorption line shape comes from a motionally narrowed (Lorentzian) component. The homogeneous component is often expressed as the homogeneous line width, Γ = 1/(πT2). In water, is on the order of a couple hundred femtoseconds while T1 and τor are a few picoseconds or longer. Therefore, the homogeneous line width is virtually completely determined by pure dephasing.

Figure 1.6c shows that there is a large difference between 1 and the initial CLS data points extrapolated to Tw = 0. This large drop is caused by fast homogeneous processes, and the difference between the Tw = 0 CLS data and 1 combined with the absorption spectrum permits the homogeneous line width to be determined. Slower processes sample the inhomogeneous distribution of frequencies and cause the Tw-dependent decay of the CLS.

The first step in extracting the FFCF involves fitting the CLS to a multiexponential decay to yield a set of amplitudes and decay constants [85]. Due to the short time approximation [85, 87, 88], the amplitude associated with the fastest of the time constants can be pushed into the homogeneous contribution. As a result, the CLS data alone cannot accurately determine the full FFCF. In order to obtain the correct homogeneous component and amplitude of the fast inhomogeneous component, it is necessary to employ the absorption line shape. The linear absorption spectrum is the Fourier transform of the linear response function, R1(t),

(1.3)

where μ10 is the transition dipole moment of the 0–1 transition, 〈ω10〉 is the average 0–1 transition frequency, and g1(t) is the line shape function,

(1.4)

Equation (1.4) contains the FFCF, as given by Eq. (1.1), showing the link between the absorption spectrum and the underlying dynamic processes. The amplitude of the fast inhomogeneous decay and the homogeneous component are the only adjustable parameters as the absorption spectrum is fit simultaneously with the CLS data. The time constants and remaining amplitudes of the CLS multiexponential fit are accurate and are fixed during the procedure. This fitting routine has been shown to accurately determine the full FFCF including both homogeneous and inhomogeneous components [84, 85]. For bulk water, it is found that there is a fast ∼400-fs decay that is attributed to fluctuations of the lengths of the hydrogen bonds [89, 90], followed by a 1.7-ps decay corresponding to global hydrogen bond network randomization [23]. In addition, there is a relatively large homogeneous contribution. The full FFCF for bulk water is listed in Table 1.1 and is shown going through the CLS points in Figure 1.6c.

Table 1.1 Bulk Water FFCF Parameters

1.2.3 Polarization-Selective Pump–Probe Spectroscopy

The pump–probe experiment illustrated in Figure 1.4b also involves three field–matter interactions. In this variation, only two beams are used (instead of the three excitation beams plus the LO in the vibrational echo experiment). A strong pump pulse and weak probe pulse are crossed in the sample. The first two field–matter interactions occur during the pump pulse so that τ = 0, and the system immediately adopts a population state in either the 0 or 1 vibrational levels. The probe pulse enters at a later time to interrogate the changes in the sample due to the pump. The pump–probe signal emits in the same phase-matched direction as the probe and self-heterodynes with the probe.

The vibrational lifetime and orientational relaxation dynamics are obtained via polarization-selective pump–probe experiments. The pump is polarized at 45° relative to the probe (which is kept at horizontal polarization). Molecules whose transition dipoles are oriented at 45° will have a higher probability of being excited, while transition dipoles oriented perpendicular to the pump will have zero probability of being excited. The probe undergoes changes in absorption due to these transitions. Because the 0–1 vibrational transition is excited, there will be increased transmission of the probe as well as stimulated emission at the 0–1 transition frequency. The induced absorption of the 1–2 transition will cause a decreased transmission of the probe at the 1–2 frequency. As the molecules reorient, fewer transition dipoles will be aligned with the pump. After the sample, the probe is resolved at parallel and perpendicular polarizations relative to the pump (±45°). The resolved probe is frequency dispersed by the monochromator and detected on the 32-pixel MCT detector. The time between the pump and probe pulses is scanned to measure the time-dependent changes in absorption of the parallel and perpendicular components. The parallel and perpendicular signals are given by

(1.5)

(1.6)

where P(t) is population relaxation and C2(t) is the second Legendre polynomial orientational correlation function for a dipole transition [91]. Pure population relaxation (no contributions from orientation) may be obtained from

(1.7)

P(t) generally adopts the form of a single or biexponential decay, depending on how many distinct OD environments a system contains:

(1.8)

for one component or

(1.9)

for two components, where A1 and 1 − A1 are the fractional populations of components 1 and 2, respectively, and the terms are the vibrational lifetimes of the ODs for the ith component. The orientational correlation function, C2(t), can be obtained from calculating the anisotropy according to

(1.10)

It is important to note that Eq. (1.10) involves dividing by the pure population relaxation [P(t) of Eq. (1.7)] to obtain pure orientational relaxation dynamics. Equation (1.10) only holds for a single-component system in which the vibrational chromophores have a single vibrational lifetime and orientational correlation function. In this case, the orientational correlation function follows single exponential dynamics,

(1.11)

where τor is a time constant describing orientational relaxation. Sometimes C2(t) can also contain a contribution from a “wobbling-in-a-cone” mechanism [92, 93], which will be discussed further in Section 1.6.

If the system contains two distinct species, each with its own vibrational lifetime and orientational relaxation time constants, then P(t) does not divide out and the anisotropy adopts a more complicated form [61, 68–72]:

(1.12)

where the and terms are the vibrational lifetimes and orientational relaxation time constants for the ith component, respectively. The fractional populations of each component are given as A1 and 1 − A1.

1.3 Linear Infrared Absorption Spectra of Water near Charged Interfaces

1.3.1 Water in Salt Solutions

The hydroxyl stretch of water, in this case the OD stretch of HOD in H2O, is extremely sensitive to local environments. Figure 1.7 shows the IR absorption band of the OD stretch (5% HOD in H2O). The band is wide [∼170 cm−1 full width at half-maximum (FWHM)] because there is a large distribution in the lengths and strengths of hydrogen bonds. Hydrogen bonds influence the attractive part of the hydroxyl vibrational potential, opening up the potential and lowering the vibrational frequency compared to the gas phase. A strong hydrogen bond will lower the vibrational transition frequency more than a weak hydrogen bond. Relative to the line center of the OD stretch of HOD in H2O, strong hydrogen bonds produce a red shift and weak hydrogen bonds produce a blue shift [21, 23, 61, 68, 72]. Raman experiments and MC calculations have suggested that the magnitude of a red or blue shift in the hydroxyl stretch frequency is actually correlated to the strength and directionality of the electric field of the hydrogen bond acceptor [20]. Figure 1.8 shows linear IR absorption spectra of the OD hydroxyl stretch in water–salt solutions at a constant ratio of 12 water molecules to 1 salt ion pair. The OD hydroxyls will interact with the anions of the salts. The salts are KF, KCl, NaCl, KBr, NaBr, KI, and NaI. As the size of the anion increases, the spectra show a greater blue shift. This trend has been observed by other groups [21, 94]. Based on the Raman and MC studies mentioned above, the blue shift arises because a more diffuse and less directed electric field is projected along the OD hydroxyl as the size of the anion increases. KF, in contrast, causes the spectrum to red shift compared to the OD stretch of HOD in bulk water. F− is a very small anion, increasing the electric field felt by the hydroxyl. The Raman and MC study also concluded that the anion only caused local changes to the hydrogen bond structure of water and that only the waters in the first solvation shell of the anion felt significant perturbation [20]. The amount of blue or red shift is correlated to electric field strength of the anion and also may reflect the strength of the hydrogen bond. These conclusions refute the idea that salts are either kosmotropes (“structure makers”) or chaotropes (“structure breakers”) [95] since any changes in hydrogen bonding structure are very localized. It can also be seen from Figure 1.8 that changing the identity of the cation (K+ to Na+) does not affect the spectra when the anions are kept the same. The lack of change with cation identity indicates that it is the direct very local interaction of the hydroxyls and anions that determines the hydroxyl stretch frequency.

Figure 1.7 Linear IR absorption spectrum of the OD stretch of 5% HOD in H2O. The absorption band is very wide (∼170 cm−1) because there is a wide range in the distribution of lengths and strengths of hydrogen bonds in the water network. The red side of the line shape is dominated by strong hydrogen bonds that lengthen the OD bond. In contrast, the blue side of the line is dominated by weak hydrogen bonds that cause a shorter OD bond length.

Figure 1.8 Linear IR absorption spectra of bulk water (5% HOD in H2O) and water–salt solutions at ratios of 12 : 1 water to salt. The salts are KF, KCl, KBr, KI, NaCl, NaBr, and NaI. As the anion increases in size, the spectra shift more to the blue. Large anions exert a weaker electric field along the OD hydroxyl to cause the blue shift. There appears to be no difference when the cation is changed but the anion kept the same.

The spectra in Figure 1.8 only show one broad absorption band even though there are arguably two types of water molecules in each system: water hydrogen bonded to other water hydroxyls and water interacting with ions. The water–NaBF4 system, in contrast, shows different behavior. Linear IR absorption spectra of the 1.7-M, 3.1-M, 4.3-M, and 5.5-M solutions are shown in Figure 1.9 [25]. Bulk water shows a single absorption band at ∼2509 cm−1, but at 1.7 M NaBF4 concentration and higher concentrations, a second, narrower band grows in at ∼2650 cm−1. Because this second band increases in size as the NaBF4 concentration increases, it is concluded that this second band is due to water hydroxyls interacting with the tetrafluoroborate anions. The main band at ∼2509 cm−1 in the spectra is attributed to water–water interactions. As discussed below, in this system, 2D IR chemical exchange spectroscopy can be used to directly measure the exchange rate between the water–ion and water–water species because cross peaks will be resolved.

Figure 1.9 Linear IR absorption spectra for water–NaBF4 solutions at varying concentrations. The main lobe around 2509 cm−1 corresponds to water–water hydrogen bonding environments while the smaller peak around 2650 cm−1 corresponds to water molecules interacting with the tetrafluoroborate anions. As the NaBF4 concentration increases, the main lobe decreases in size, and the smaller lobe increases.

1.3.2 Water in AOT Reverse Micelles

Figure 1.10 displays linear IR absorption spectra for water (5% HOD in H2O) inside the full range of reverse micelle w0's examined, from w0 = 2 to w0 = 46. For clarity, a few w0's are omitted because they closely overlap with neighboring spectra. As the reverse micelle size decreases, the spectra systematically shift to the blue. While bulk water peaks around 2509 cm−1, the w0 = 2 spectrum peaks around 2565 cm−1. This blue shift trend occurs because as reverse micelle size decreases, proportionally more water molecules interact with the sulfonate head groups [61, 68, 70]. The sulfonate group is a large anion, so it exerts a weak electric field along the OD hydroxyl stretch.

Figure 1.10 Linear IR absorption spectra for water inside AOT reverse micelles from w0 = 2 through w0 = 46 along with bulk water for comparison. The spectra steadily shift to the blue as the reverse micelle size decreases.

Figure 1.1 illustrates that distinct water environments exist in the reverse micelle. If large enough, the reverse micelle can support a core of bulklike water. The bulk water core is then surrounded by a layer of interfacial water molecules associated with the head group interface. It has been shown that the spectra in Figure 1.10 can be reproduced by a linear combination of the spectrum of bulk water and the spectrum of w0 = 2 [61, 68, 70]. In the w0 = 2 sample, essentially all of the water molecules interact with the head groups, and there is no bulklike core. The w0 = 2 spectrum is taken to be the spectrum of interfacial water. This model of decomposing the reverse micelle spectra into bulk and interfacial contributions has been referred to as the core–shell model and is described by

(1.13)

where I1 and I2 are the component spectra for bulk water and w0 = 2, respectively, and a1 is the fractional population. When fitting the reverse micelle spectra, a1 is the only adjustable parameter. Figure 1.11 displays the core–shell decomposition for AOT w0 = 25. The circles are the data and the solid line through the circles is the weighted sum of the bulk water spectrum and the w0 = 2 spectrum. The agreement between the original spectrum and the linear combination is excellent. Table 1.2 [70] lists the 1 − a1 parameter (interfacial fraction) for AOT reverse micelles of w0 = 25, 37, and 46. The numbers indicate that the fraction of interfacial water decreases as water pool diameter increases. Fractions are listed for both the IR spectral decomposition analysis as well as geometric calculations. If the interfacial region is modeled as a shell of approximately one water molecule thick (about 2.8 Å), then the interfacial fractions are nearly identical to those obtained by spectral analysis. These results strongly indicate that in large reverse micelles, the interface gives rise to a very locally perturbed region of interfacial water that does not extend far beyond the first solvation shell of the head groups [70].

Figure 1.11 Two-component decomposition of the water spectrum of AOT w0 = 25, a large reverse micelle. The w0 = 25 spectrum can be reproduced by a linear combination of the bulk water spectrum and the w0 = 2 spectrum in which nearly all the water molecules interact with the head group interface.

Table 1.2 Fractional Population of Interfacial Water in Large AOT Reverse Micelles

1.3.3 Water in Igepal Reverse Micelles

The linear IR absorption spectra of water inside of Igepal reverse micelles, as shown in Figure 1.12, also show a systematic blue shift as the water pool diameter decreases [71]. Figure 1.12 compares spectra of water in Igepal reverse micelles to water in AOT reverse micelles of the same size. Igepal w0 = 20 and AOT w0 = 25 each have a water pool diameter of 9 nm while Igepal w0 = 12 and AOT w0 = 16.5 each have a water pool diameter of 5.8 nm. The Igepal spectrum of a given w0 is not as blue shifted as its AOT counterpart of the same size even though both systems contain identical amounts of water. It is important to note that identical w0's of Igepal and AOT do not yield the same water pool diameters because of different aggregation numbers of the two surfactants; that is why different w0's are used to produce reverse micelles of the same size. Igepal reverse micelles, like the AOT systems, also contain bulk and interfacial water regions. In the case of Igepal, the waters interact with hydroxyl head groups, but in AOT they interact with charged sulfonate head groups. The chemical identity of the interface clearly alters the linear absorption behavior of the water molecules.

Figure 1.12 Linear IR absorption spectra for water inside Igepal reverse micelles compared to AOT reverse micelles of the same size. Igepal w0 = 20 has the same size as AOT w0 = 25 (9 nm diameter), and Igepal w0 = 12 has the same size as AOT w0 = 16.5 (5.8 nm). Spectra for systems of the same diameter are not equivalent to each other, indicating that the hydrogen bonding interactions in the Igepal and AOT systems are different.

The Igepal spectra are not decomposed into core and shell regions like the AOT systems because Igepal reverse micelles are not well characterized below w0 ∼10 [78], so there is no low water content spectrum (akin to AOT w0 = 2) to approximate the pure interfacial water spectrum. Furthermore, the Igepal reverse micelle system contains a third environment in addition to the core and interfacial water regions. The hydroxyl head groups will exchange deuterium atoms with the 5% HOD in the water pool. As a result, the interfacial region is composed of two types of hydroxyls: waters at the interface and OD head groups. The population of OD head groups is small, and their contribution to the experimental observables has been treated in detail so that the interfacial water dynamics can still be extracted (see Section 1.4.2) [71]. Figure 1.12 shows that there are significant spectral changes upon changing the chemical identity of the interface, but it will be shown below that changing the charged sulfonate head groups at the interface to neutral hydroxyl groups does not significantly affect water's hydrogen bonding dynamics.

1.4 Population Relaxation and Orientational Relaxation

1.4.1 At the Interface of Large and Intermediate AOT Reverse Micelles

The spectral analysis in Section 1.3.2 shows that large reverse micelles can be divided into two distinct regions: bulklike core water and interfacial water. This core–shell model (Fig. 1.11) has critical implications in the treatment of large reverse micelles, as it implies that the dynamics of the core and interface regions can be separated as well, with the core exhibiting the dynamics of pure water. In pure water, the vibrational lifetime of the OD stretch is 1.8 ps [using Eq. (1.8)] [68, 70], and the orientational relaxation time is 2.6 ps [using Eq. (1.11)] [96]. Consequently, the interfacial dynamics are the only unknowns in measurements on the water pools of large reverse micelles. Based on spectral analysis, waters that interact with the interface experience a blue shift in their linear IR absorption spectra. As shown by the inset in Figure 1.13, the nearly purely interfacial w0 = 2 OD–water spectrum peaks at 2565 cm−1. The experimental pump-probe signal is frequency dispersed by a monochromator, so the dynamics at different frequencies across the absorption line shape can be monitored. For example, if the dynamics of w0 = 25 were measured at 2519 cm−1, then water molecules sampling the center of the absorption line shape would be mostly observed. Most of the spectral contribution at that frequency is due to bulklike water molecules in the core, as shown by Figure 1.11 and the inset in Figure 1.13. At higher frequencies, such as 2565 cm−1, the water molecules at the surfactant interface make a larger contribution to the signal than they do toward the center of the absorption line. To accurately extract the dynamics at the interface from the dynamics of the core (which are known), it is useful to have the greatest amount of interfacial water contributing to the signal as possible. Because of this requirement, it is best to analyze the pump–probe signals detected at frequencies greater than or equal to 2565 cm−1 where it is likely that a significant portion of the signal comes from interfacial water.

Figure 1.13 Population relaxation data for AOT w0 = 25 at three detection frequencies. The inset shows that interfacial water molecules have more population at frequencies above 2565 cm−1. The three curves have a slight frequency dependence to them because of changing fractions of core and interfacial water.

Figure 1.13 displays the population relaxation decay curves [Eq. (1.7)] for w0 = 25 at three different frequencies: 2519 cm−1 (near the absorption center of w0 = 25), 2568 cm−1 (near the peak absorption of w0 = 2 interfacial water), and 2630 cm−1 (far to the blue) [70]. The three curves are not the same, indicating that there is a frequency dependence to the population decay. When there are two separate water ensembles, the population decay is the weighted sum of the population decay of the two components, as given by Eq. (1.9). When the curves in Figure 1.13 are fit to Eq. (1.9), the vibrational lifetime of the first component (bulk water) is kept constant at its literature value such that at all frequencies. The superscript w stands for the lifetime of the OD stretch of dilute HOD in bulk water. Data at all three frequencies are fit simultaneously to Eq. (1.9), and A1 (the fractional population) and (the interfacial water lifetime) are the only adjustable parameters. It is found that this two component model for the population decay accurately describes the population relaxation dynamics for all AOT reverse micelles of w0 = 12 and larger. For all reverse micelles of w0 ≥ 12, the vibrational lifetime of interfacial water molecules is found to be 4.3 ± 0.5 ps. This value of ∼4.3 ps remains constant regardless of w0 and regardless of frequency, indicating that the dynamics of water at the interface are quite distinct from those of the bulk water core in large reverse micelles [68, 70]. Table 1.3 lists the obtained values for the large reverse micelles, showing the invariability. The only parameter that changes with wavelength (leading to the three different curves in Figure 1.13) is A1. As frequency increases, the amount of interfacial water increases, so 1 − A1 becomes larger. As w0 decreases, there is more interfacial water overall, so the 1 − A1 values collectively increase.

Table 1.3 Interfacial Vibrational Lifetime, , and Orientation Relaxation Times, , for Large AOT Reverse Micelles

Because there are two components to the population relaxation, the orientational relaxation must also be fit to a two-component model, as described by Eq. (1.12). There are quite a few variables in this equation, but population relaxation analysis allows many of the parameters in Eq. (1.12) to be fixed. A1, , , and (the known bulk water reorientation time of 2.6 ps) all can be fixed, leaving the orientational relaxation time of the interface, , as the only adjustable parameter. Figure 1.14 shows the anisotropy decays [