Reviews in Fluorescence 2008

Reviews in Fluorescence 2008 Editor Chris D. Geddes University of Maryland, Baltimore, MD, USA For further volumes: h...

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Reviews in Fluorescence 2008

Editor Chris D. Geddes University of Maryland, Baltimore, MD, USA

For further volumes: http://www.springer.com/series/6946

.

Chris D. Geddes Editor

Reviews in Fluorescence 2008

123

Editor Chris D. Geddes Institute of Fluorescence University of Maryland 701 E. Pratt St. Baltimore, MD 21202 Suites 3017-21 USA [email protected]

ISSN 1573-8086 ISBN 978-1-4419-0828-5 e-ISBN 978-1-4419-1260-2 DOI 10.1007/978-1-4419-1260-2 Springer New York Dordrecht Heidelberg London Library of Congress Control Number: 2009940098 © Springer Science+Business Media, LLC 2010 All rights reserved. This work may not be translated or copied in whole or in part without the written permission of the publisher (Springer Science+Business Media, LLC, 233 Spring Street, New York, NY 10013, USA), except for brief excerpts in connection with reviews or scholarly analysis. Use in connection with any form of information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed is forbidden. The use in this publication of trade names, trademarks, service marks, and similar terms, even if they are not identified as such, is not to be taken as an expression of opinion as to whether or not they are subject to proprietary rights. Printed on acid-free paper Springer is part of Springer Science+Business Media (www.springer.com)

Preface

This is the fifth volume in the Reviews in Fluorescence series. To date, four previous volumes have been both published and well received by the scientific community. Several book reviews in the last few years have also favorably remarked on the series. In this fifth volume we continue the tradition of publishing leading edge and timely articles from authors around the world. With the recent Nobel Prize in Chemistry for 2008 being awarded for the discovery and development of the Green Fluorescent Protein (GFP) to Shimomura, Chalfie, and Tsien, we have subsequently included several timely reviews on GFP in this volume. We thank the authors for their timely and exciting contributions. We hope you find this volume as useful as past volumes, which promises to be just as diverse with regard to fluorescence-based content. Finally, in closing, I would like to thank Caroleann Aitken for helping coordinating content with authors and Michael Weston at Springer for help in publishing this current volume. Baltimore, MD, USA

Chris D. Geddes

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Contents

Fluorescence Anisotropy to Study the Preferential Orientation of Fluorophores in Ordered Bi-Dimensional Systems: Rhodamine 6G/Laponite Layered Films . . . . . . . . . . . . . . . . . . F. López Arbeloa, V. Martínez, T. Arbeloa, and I. López Arbeloa Room Temperature Tryptophan Phosphorescence of Proteins in the Composition of Biological Membranes and Solutions . . . . . . . Vladimir M. Mazhul’, Alexander V. Timoshenko, Ekaterina M. Zaitseva, Svetlana G. Loznikova, Inessa V. Halets, and Tatsiana S. Chernovets Rational Design of FRET-Based Sensor Proteins . . . . . . . . . . . . . M. Merkx Fluorescence Imaging of Calcium Loading and Mitochondrial Depolarization in Cancer Cells Exposed to Heat Stress . . . . . . . . . . Olaf Minet, Cathrin Dressler, Jürgen Beuthan, Urszula Zabaryło, Rasa Zukiene, and Vida Midaziene Energy Transfer in Silica Nanoparticles: An Essential Tool for the Amplification of the Fluorescence Signal . . . . . . . . . . . . . Sara Bonacchi, Damiano Genovese, Riccardo Juris, Ettore Marzocchi, Marco Montalti, Luca Prodi, Enrico Rampazzo, and Nelsi Zaccheroni Spectroscopic Characterization of Plasma – Chemically Functionalized and Fluorophore-Labeled Polymer Surfaces . . . . . . . Katrin Hoffmann, Renate Mix, Joerg F. Friedrich, and Ute Resch-Genger

1

37

69

89

119

139

Fluorescent Labeling and Its Effect on Hybridization of Oligodeoxyribonucleotides . . . . . . . . . . . . . . . . . . . . . . . . Ramendra K. Singh and Shipra Agarwal

161

New Method for Determining Histamine Rate in Halieutic Products . . . . . . . . . . . . . . . . . . . . . . . . . . . . Alphonse Tine and Stéphy E. Douabalé

195

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Contents

Spectroscopy of DNA–Actinomycin Complexes . . . . . . . . . . . . . . Nikolai Vekshin Fluorescence Spectroscopy in Optoelectronics, Photomedicine, and Investigation of Biomolecular Systems . . . . . . . . . . . . . . . . . . . . . . . . . . . Danuta Wróbel, Alina Dudkowiak, and Jacek Goc

219

237

Multicolor Imaging with Fluorescent Proteins in Mice . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Robert M. Hoffman

277

Genetically Encoded Fluorescent and Bioluminescent Probes for Illuminating Cellular Signaling Pathways . . . . . . . . . . . . . . . Yoshio Umezawa

303

Fluorescent Protein FRET Applications . . . . . . . . . . . . . . . . . . Annalee W. Nguyen, Xia You, Abeer M. Jabaiah, and Patrick S. Daugherty

321

Imaging Protein Interactions in Living Cells Using the Fluorescent Proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . Richard N. Day, Ammasi Periasamy, and Ignacio Demarco

337

Engineering Green Fluorescent Proteins Using an Expanded Genetic Code . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Prajna Paramita Pal and Nediljko Budisa

359

Fluorescent Proteins in Transgenic Plants . . . . . . . . . . . . . . . . . Reginald J. Millwood, Hong S. Moon, and C. Neal Stewart Jr.

387

Peptide Foldamers: From Spectroscopic Studies to Applications . . . . Lorenzo Stella, Gianfranco Bocchinfuso, Emanuela Gatto, Claudia Mazzuca, Mariano Venanzi, Fernando Formaggio, Claudio Toniolo, Antonio Palleschi, and Basilio Pispisa

405

Circularly Polarized Luminescence (CPL) of Proteins and Protein Complexes . . . . . . . . . . . . . . . . . . . . . Eugene Gussakovsky New Dual Fluorescent Dyes Based on Modified “Excited State with Extended Conjunction” Photophysical Model . . . . . . . . . . . . Michael P. Begaye, Premchendar Nandhikonda, Zhi Cao, and Michael D. Heagy Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

425

461

479

Contributors

Shipra Agarwal Department of Chemistry, Indian Institute of Technology Bombay, Powai, Mumbai-400076. India Fernando López Arbeloa Departamento de Química Física, Universidad del País Vasco UPV/EHU, Apartado 644, 48080-Bilbao, Spain, [email protected] Inigo López Arbeloa Departamento de Química Física, Universidad del País Vasco UPV/EHU, Apartado 644, 48080-Bilbao, Spain Teresa Arbeloa Departamento de Química Física, Universidad del País Vasco UPV/EHU, Apartado 644, 48080-Bilbao, Spain Michael P. Begaye Department of Chemistry, New Mexico Institute of Mining & Technology, Socorro, NM 87801, USA Jürgen Beuthan Charité – Universitätsmedizin Berlin, CC6 AG Medizinische Physik/Optische Diagnostik, Fabeckstr. 60-62, 14195 Berlin, Germany Gianfranco Bocchinfuso Dipartimento di Scienze e Tecnologie Chimiche, Università di Roma Tor Vergata, Rome, Italy Sara Bonacchi Dipartimento di Chimica “G. Ciamician,” Latemar Unit, Università degli Studi di Bologna, Via Selmi 2, 40126 Bologna, Italy Nediljko Budisa Max-Planck-Institut für Biochemie, BioFuture Research Group – Molecular Biotechnology, Am Klopferspitz 18, D-82152 Martinsried, Germany Zhi Cao Department of Chemistry, New Mexico Institute of Mining & Technology, Socorro, NM 87801, USA Tatsiana Chernovets Laboratory of Proteomics, Institute of Biophysics and Cellular Engineering, National Academy of Sciences of Belarus, 27 Academicheskaya Str., Minsk, 220072, Belarus Patrick S. Daugherty Department of Chemical Engineering, University of California, Santa Barbara, CA 93106-5080, USA

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Contributors

Richard N. Day Departments of Medicine and Cell Biology, University of Virginia Health Services, Charlottesville, VA 22908, USA Ignacio Demarco Department of Cell Biology, University of Virginia Health Services, Charlottesville, VA 22908, USA Cathrin Dressler LMTB GmbH, Fabeckstr. 60-62, 14195 Berlin, Germany Alina Dudkowiak Institute of Physics, Faculty of Technical Physics, Poznan University of Technology, Nieszawska 13A, 60-965 Poznan, Poland Stéphy Edgard Douabalé Faculté des Sciences et Techniques Université Cheikh Anta Diop, Dakar, Sénégal Fernando Formaggio Dipartimento di Scienze Chimiche, Università di Padova, Padova, Italy Joerg F. Friedrich Federal Institute for Materials Research and Testing (BAM), Richard-Willstaetter-Str. 11, D-12489 Berlin, Germany Emanuela Gatto Dipartimento di Scienze e Tecnologie Chimiche, Università di Roma Tor Vergata, Rome, Italy Damiano Genovese Dipartimento di Chimica “G. Ciamician,” Latemar Unit, Università degli Studi di Bologna, Via Selmi 2, 40126 Bologna, Italy Jacek Goc Institute of Physics, Faculty of Technical Physics, Poznan University of Technology, Nieszawska 13A, 60-965 Poznan, Poland Eugene Gussakovsky Institute for Biodiagnostics, National Research Council Canada, Winnipeg, Manitoba R3B 1Y6, Canada Inessa Halets Laboratory of Proteomics, Institute of Biophysics and Cellular Engineering, National Academy of Sciences of Belarus, 27 Academicheskaya Str., Minsk, 220072, Belarus Michael D. Heagy Department of Chemistry, New Mexico Institute of Mining & Technology, Socorro, NM 87801, USA Katrin Hoffmann Federal Institute for Materials Research and Testing (BAM), Richard-Willstaetter-Str. 11, D-12489 Berlin, Germany Robert M. Hoffman AntiCancer, Inc., San Diego, CA 92111, USA; Department of Surgery, University of California, San Diego, CA 92103, USA Riccardo Juris Dipartimento di Chimica “G. Ciamician,” Latemar Unit, Università degli Studi di Bologna, Via Selmi 2, 40126 Bologna, Italy Abeer M. Jabaiah Department of Chemical Engineering, University of California, Santa Barbara, CA 93106-5080, USA

Contributors

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Svetlana Loznikova Laboratory of Proteomics, Institute of Biophysics and Cellular Engineering, National Academy of Sciences of Belarus, 27 Academicheskaya Str., Minsk, 220072, Belarus Virginia Martínez Departamento de Química Física, Universidad del País Vasco UPV/EHU, Apartado 644, 48080-Bilbao, Spain Ettore Marzocchi Dipartimento di Chimica “G. Ciamician,” Latemar Unit, Università degli Studi di Bologna, Via Selmi 2, 40126 Bologna, Italy Vladimir Mazhul Laboratory of Proteomics, Institute of Biophysics and Cellular Engineering, National Academy of Sciences of Belarus, 27 Academicheskaya Str., Minsk, 220072, Belarus, [email protected] Claudia Mazzuca Dipartimento di Scienze e Tecnologie Chimiche, Università di Roma Tor Vergata, Rome, Italy Maarten Merkx Laboratory of Chemical Biology, Department of Biomedical Engineering, Eindhoven University of Technology, Eindhoven, Netherlands, [email protected] Vida Mildažiene Vytautas Magnus University, Vileikos 8, LT-44404 Kaunas, Lithuania Reginald J. Millwood Department of Plant Sciences, The University of Tennessee, Knoxville, TN, USA, [email protected] Olaf Minet Charité – Universitätsmedizin Berlin, CC6 AG Medizinische Physik/Optische Diagnostik, Fabeckstr. 60-62, 14195 Berlin, Germany, [email protected] Renate Mix Federal Institute for Materials Research and Testing (BAM), Richard-Willstaetter-Str. 11, D-12489 Berlin, Germany Marco Montalti Dipartimento di Chimica “G. Ciamician,” Latemar Unit, Università degli Studi di Bologna, Via Selmi 2, 40126 Bologna, Italy Hong Moon Department of Plant Sciences, The University of Tennessee, Knoxville, TN, USA Premchendar Nandhikonda Department of Chemistry, New Mexico Institute of Mining & Technology, Socorro, NM 87801, USA Annalee W. Nguyen Department of Chemical Engineering, University of California, Santa Barbara, CA 93106-5080, USA Antonio Palleschi Dipartimento di Scienze e Tecnologie Chimiche, Università di Roma Tor Vergata, Rome, Italy Prajna Paramita Pal Max-Planck-Institut für Biochemie, BioFuture Research Group – Molecular Biotechnology, Am Klopferspitz 18, D-82152 Martinsried, Germany

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Contributors

Ammasi Periasamy Keck Center for Cellular Imaging, Department of Biology, Gilmer Hall, University of Virginia, Charlottesville, VA 22904, USA Basilio Pispisa Dipartimento di Scienze e Tecnologie Chimiche, Università di Roma Tor Vergata, Rome, Italy Luca Prodi Dipartimento di Chimica “G. Ciamician,” Latemar Unit, Università degli Studi di Bologna, Via Selmi 2, 40126 Bologna, Italy Enrico Rampazzo Dipartimento di Chimica “G. Ciamician,” Latemar Unit, Università degli Studi di Bologna, Via Selmi 2, 40126 Bologna, Italy Ute Resch-Genger Federal Institute for Materials Research and Testing (BAM), Richard-Willstaetter-Str. 11, D-12489 Berlin, Germany, [email protected] Ramendra K. Singh Nucleic Acids Research Laboratory, Department of Chemistry, University of Allahabad, Allahabad-211002, India, [email protected] Lorenzo Stella Dipartimento di Scienze e Tecnologie Chimiche, Università di Roma Tor Vergata, Rome, Italy, [email protected] C. Neal Stewart Jr Department of Plant Sciences, The University of Tennessee, Knoxville, TN, USA Aleksandr Timoshenko Laboratory of Proteomics, Institute of Biophysics and Cellular Engineering, National Academy of Sciences of Belarus, 27 Academicheskaya Str., Minsk, 220072, Belarus Pr Alphonse Tine Faculté des Sciences et Techniques Université Cheikh Anta Diop, Dakar, Sénégal Claudio Toniolo Dipartimento di Scienze e Tecnologie Chimiche, Università di Roma Tor Vergata, Rome, Italy Yoshio Umezawa Department of Chemistry, School of Science, The University of Tokyo Hongo, Bunkyo-ku, Tokyo 113-0033, Japan, [email protected] Nikolai Vekshin Institute of Cell Biophysics, Pushchino, Moscow region, 142290, Russia Mariano Venanzi Dipartimento di Scienze e Tecnologie Chimiche, Università di Roma Tor Vergata, Rome, Italy Danuta Wróbel Institute of Physics, Faculty of Technical Physics, Poznan University of Technology, Nieszawska 13A, 60-965 Poznan, Poland, [email protected] Xia You Department of Chemical Engineering, University of California, Santa Barbara, CA 93106-5080, USA

Contributors

xiii

Urszula Zabaryło Charité – Universitätsmedizin Berlin, CC6 AG Medizinische Physik/Optische Diagnostik, Fabeckstr. 60-62, 14195 Berlin, Germany Nelsi Zaccheroni Dipartimento di Chimica “G. Ciamician”, Latemar Unit, Università degli Studi di Bologna, Via Selmi 2, 40126 Bologna, Italy Ekaterina Zaitseva Laboratory of Proteomics, Institute of Biophysics and Cellular Engineering, National Academy of Sciences of Belarus, 27 Academicheskaya Str., Minsk, 220072, Belarus ¯ Rasa Žukien˙ e Vytautas Magnus University, Vileikos 8, LT-44404 Kaunas, Lithuania

Fluorescence Anisotropy to Study the Preferential Orientation of Fluorophores in Ordered Bi-Dimensional Systems: Rhodamine 6G/Laponite Layered Films F. López Arbeloa, V. Martínez, T. Arbeloa, and I. López Arbeloa

Abstract Absorption and fluorescence spectroscopies with linearly polarized light are applied to study the anisotropic behavior of a fluorescence dye (rhodamine 6G, R6G) adsorbed in ordered clay (laponite, Lap) particles. Films elaborated by the spin-coating technique provide a parallel stacking of clay layers in the supported substrates. The posterior intercalation of the R6G molecules into the interlayer space of Lap films with a preferential orientation with respect to the normal to the clay layers gives rise to a macroscopic orientation of dye molecules into the 2D surfaces. Such an organization induces an anisotropic behavior with a photoresponse of dye/clay films to the plane of the polarized light. A mathematic procedure, based on the evolution of the fluorescence anisotropy with the twisting angle of the films with respect to the excitation light, is used to evaluate the preferential orientation of R6G molecules in Lap films. The fluorescence method can be extended to study the preferential orientation of fluorescent molecules adsorbed in any organized rigid 2D system. Keywords Fluorescence Anisotropy · Fluorescence Dyes · Organized 2D Systems · Rhodamine 6G · Clay Films · Absorption with Polarized Light

1 Introduction Recent advances in strategic fields such as the nanoscience and chemistry of intercalation have required collaboration between chemists, biologists, physicists, and engineers, providing a multidisciplinary character to the scientific community. In this sense, the synthesis of multifunctional materials with potential technological applications involves, in most cases, hybrid organo-inorganic materials. Thus, the combination of optical, spectroscopic, electronic, and chemical properties of organic F. López Arbeloa (B) Departamento de Química Física, Universidad del País Vasco, Apartado 644, 48080-Bilbao, Spain e-mail: [email protected]

C.D. Geddes (ed.), Reviews in Fluorescence 2008, Reviews in Fluorescence 2008, DOI 10.1007/978-1-4419-1260-2_1, C Springer Science+Business Media, LLC 2010

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F. López Arbeloa et al.

guest molecules with the structural, mechanical, and thermal characteristics of inorganic host materials is of capital importance in the development of new opto- and photo-electronic devices [1–10]. In particular, three important aspects need to be addressed: – To improve the photo-, thermal- and chemical stability of the guest molecules adsorbed into adequate frameworks. Indeed, chromophores embedded into different porous materials experience an important decrease in their photodegradation [9]. – To modify and control the electrical, magnetic, and optical properties of the guest molecules in constrained and rigid nanostructured environments in order to manufacture materials with very specific technological applications [1, 4, 6, 9]. – To use guest molecules as molecular probes in the characterization of support materials [4, 6, 8–11]. In this sense, fluorescence molecules have been largely applied as molecular probes to characterize many biological systems [12–14]. Several systems of technological interest have been successfully elaborated by encapsulating photoactive guest molecules into adequate rigid architectures with nano-scale organized frameworks. These systems have particular applications as photocatalyzers, photovoltaic cells, optical sensors, optical storage devices, and solid-state dye lasers [6–10, 15–27]. Moreover, in recent years, the design of new hybrid systems has been focused on obtaining macroscopic arrangements of organized guest molecules [28–33]. This can lead to new materials with nonlinear optical (NLO) properties, offering potential applications as dichroic filters, second harmonic generation devices, etc [1, 9, 10, 34, 35]. A great variety of organic and inorganic host materials (nanotubes, fullerenes, membranes, cyclodextrines, micelles, polymers, laminar oxides, silica gels, zeolites), offering a multitude of different mono-, bi-, and tri-dimensional architectures, can be used for this purpose [6–10, 36–39]. The elaboration of thin films of organized host systems is an interesting approach, because they can easily provide two-dimensional structured arrangements. There are several methodologies to elaborated 2D ordered layered systems, generally based on a sequential way to control the multilayer deposition on supported surfaces [40]. For instance: Langmuir–Blodgett (LB) films formed by van der Waals interaction between different amphiphilic (fatty acids, phospholipid, or surfactant) molecules [41–43]; layer-by-layer assemblies formed by electrostatic interaction between oppositely charged layers (i.e. negative and positive charged polymers) [44, 45]; or self-assembly block copolymers providing a great variety of ordered structures [46]. However, from a technological point of view, the macroscopic organization of dye molecules in a bi-dimensional surface is better performed by means of inorganic layered materials. Indeed, the inorganic frameworks provide more rigid and higher transparency to the visible radiation and better thermal and chemical stabilities. Clay minerals, layered double hydroxides, zirconium phosphate, etc. [6–10, 47–50], are

Fluorescence Anisotropy in 2D Ordered Systems

3

adequate inorganic host materials to design highly ordered dye molecules in a 2D distribution. We are particularly interested in the adsorption of rhodamine dyes into clay layers. Several reviews of dye/clay systems have been published in the last few years [9, 51–57]. The photophysics of rhodamines encapsulated in constrained nanostructured domains in general and in solid clay thin films in particular, has special interests [58–76] For instance: the improvement in the photophysical and lasing properties of the dye, the enhancement in the photo- and thermostability and the elaboration of macroscopic organized arrangements of R6G molecules with nonlinear optical properties. In the present review, a new method based on fluorescence anisotropy [77, 78] is applied to evaluate the preferential orientation of a fluorescent dye (rhodamine 6G, R6G) intercalated into the interlayer space of ordered clay (laponite, Lap) films [57]. The present methodology can be extended to study other systems with a macroscopic organized bi-dimensional arrangement.

2 Fluorescence Anisotropy in Ordered Bi-Dimensional Systems Fluorescence anisotropy is a powerful technique to evaluate the preferential orientation of fluorescent organic chromophores adsorbed in ordered assemblies [79–86]. Fluorescence anisotropy is based on the recording of the fluorescence intensity, both frequency- or time-resolved, for two orthogonally oriented polarizations, the parallel (I|| ) and the perpendicular (I⊥ ) orientations with respect to the direction of the polarized excitation light (Fig. 1). Similar to the absorption spectroscopy with linearly polarized light, the fluorescence intensity will depend on the relative orien and the direction of tation of the emission transition moment of the fluorophore (M) the emission polarizer [80, 82, 86–90], concretely proportional to the square of the in the direction of the polarizer, Eqs. 1 and 2. projection of M

Y M M’

Fig. 1 Scheme for the fluorescence polarization of a fluorophore after excitation with polarized light (a reorientation of the fluorophore during its fluorescent lifetime is also included)

Y‛

ψ X

4


I αM 2 cos2 ψ

(1)

I⊥ αM 2 sin2 ψ

(2)

Consequently, the fluorescence anisotropy should be able to provide the relative orientation of fluorescent molecules (with xyz sample coordinates) with respect to the XYZ laboratory coordinates. There are several parameters to define the fluorescence polarization of a sample, such as the anisotropy r = (I|| – I⊥ )/(I|| + 2I⊥ ) or the polarization P = (I|| – I⊥ )/(I|| + I⊥ ), easily interchangeable with each other [87]. However, the dichroic ratio (D) is the most recommended anisotropy parameter to evaluate the angle of oriented fluorophores in rigid ordered systems. The dichroic ratio is defined as the ratio between the two orthogonally polarized emission intensities (D ≡ I|| /I⊥ ), and directly provides the orientation angle by means of: D ≡ I /I⊥ = cot2 ψ

(3)

A new method, based on fluorescence anisotropy, has been developed to evaluate the preferential orientation of fluorescent molecules adsorbed in ordered bi-dimensional layered materials, such as supported clay films [77, 78]. Indeed, it has been proven that the dichroic ratio D shows a linear relationship versus the twisted angle between the normal to the film and the excitation beam (δ), from which the orientation angle of the fluorescent molecules with respect to the normal to the supported films (ψ) can be evaluated. We will now briefly describe the mathematical procedure showing this linear relationship. Figure 2 shows the experimental setup used to record linearly polarized fluorescence spectra of dye/clay films by means of a SPEX fluorometer model Fluorolog 3–22. The fluorescence intensity of supported films was recorded by means of the front-face configuration, in which the detection channel is placed at an angle θ = –22.5º with respect to the excitation beam. The mathematical procedure

δ film

Y

y Z Y •

mrror

emission polarizer

22.5°

y

.

x z film

x z

Z X

δ mirror

X emission polarizer

Y’ Z

22.5°

X’ mirror

mirror

excitation polarizer

Z’ X’

• Y’

excitation polarizer

Fig. 2 Experimental set-up to record the fluorescence polarization of solid films by the front-face configuration in a SPEX spectrofluorometer model Fluorolog 3–22


5

below is derived for this arrangement, although all mathematical equations can be adjusted for experimental setups with other configurations. The sample films were rotated around their vertical y-axis by an angle δ (counterclockwise) by means of a revolving solid sample compartment. The samples were excited in the Z -direction (light propagation) with vertical (V, Y -axis) or horizontal (H, X -axis) polarized light and the fluorescence intensity was recorded in the Z-axis direction (–22.5º with respect to the excitation beam) with the emission polarizer oriented in the V- (Y-axis) and H- (X-axis) directions. The films were rotated around their vertical y-axis by a twisting angle δ (angle between the normal to the film, z-axis, and the excitation beam, Z -axis) from –20º to 50º. The fluorescence spectra can be recorded for the different combinations of the excitation and emission polarizers, i.e. IHH , IVH , IHV , and IVV , where the first and the second subindexes refer to the orientation of the excitation and emission polarizers, respectively. The fluorescence dichroic ratio consists in the comparison of the fluorescence intensity with H- and V-orientation of the emission polarizer with a constant polarized excitation (H or V). Under these experimental conditions the recorded fluorescence signals will be proportional to: IHH = IX X αIab (X )kfl (X)

(4)

IHV = IX Y αIab (X )kfl (Y)

(5)

IVH = IY X αIab (Y )kfl (X)

(6)

IVV = IY Y αIab (Y )kfl (Y)

(7)

where Iab (X ) and Iab (Y ) are the absorption intensity for H- and V-polarized excitation light at the excitation wavelength, respectively, and kfl (X) and kfl (Y) are the radiative rate constant of the fluorescence light for the emission polarizer oriented in the H- and V-directions, respectively. The Iab and kfl parameters are given by the square of the projection of the absorption and fluorescence transition dipole moment into the excitation and emission polarizer directions, respectively. The proportional constants of Eqs. 4–7 should include the instrumental response for the excitation and emission polarized light and an orientation factor that takes into account the possible reorientation of the transition dipole moment of the chromophores during the fluorescence lifetime. This orientation factor, which can be time-dependent, is responsible for the fluorescent depolarization of the excitation polarization in many nonrigid and randomly distributed molecular systems [87], and has been largely used to determine the rotational diffusion motion of fluorescent or labeled-fluorescent proteins and other biological systems [91–93]. Many processes can affect the reorientation factor. Most important are: 1. The absorption transition moment in the excitation process is not collinear with the emission transition moment in the fluorescence process. This is not the case

6

2.

3.

4. 5.


for the S0 −S1 excitation of the fluorophore, since the same electronic states are involved in the absorption and emission processes. Reorientation of excited molecules during the fluorescence lifetime. This is an important mechanism for the fluorescent depolarization of the excitation light in fluid samples, but is neglectable in rigid (solid) fluorophore systems where the reorientation by a molecular diffusional mechanism is restricted (at least during a fluorescence lifetime in the time-scale of ns). Energy transfer (ET) between different excited states, i.e. from the excited state directly populated by the excitation process to a noncollinear fluorescent excited state. In this case, the reorientation factor could provide the relative orientation between the donor and the acceptor moieties. Energy migration of the excitation energies between two molecules of the same species but with a different relative orientation. Other trivial phenomena such as reabsorption and reemission effects or light scattering.

Since we are interested in evaluating the orientation angle of fluorescent molecules adsorbed in ordered bi-dimensional systems by fluorescence polarization, the effect of the reorientation factor on the fluorescence anisotropy has to be taken into account (and, if possible, to be minimized). For ordered rigid dye/clay solid films with low dye loadings, the following prerequisites can be considered [57, 77, 78]: – Depending on the dye loading, several dye species (monomers, dimers, and higher-order aggregates) can be adsorbed into the clay surfaces. – Dye molecules of the same species are adsorbed with a preferential orientation with respect to the normal to the clay layers. – Supported films provide a macroscopic parallel stacking of clay layers and the posterior intercalation of dye molecules with a preferential orientation with respect to the film normal induces a macroscopic organization of dye molecules in this direction. – The excitation is directly performed to the fluorescent S1 excited state (S0 −S1 absorption). – Fluorescence anisotropy is analyzed at those emission wavelengths in which the fluorescence intensity is much higher than the intensity of the scattering light. Because of the nature of the solid samples, very high scattering can be detected at those wavelengths close to the excitation wavelength. For this reason, shorter wavelengths in the S0 −S1 absorption band are recommended for the excitation process. Defining the fluorescence dichroic ratio (DHV ) as the ratio of both detected emission polarizations for a constant excitation polarization, DHV ≡ IHH /IHV = IVH /IVV , then:


DHV ≡

IHH IVH SH kfl (X) = = IHV IVV SV kfl (Y)

7

(8)

where SH and SV , which can be wavelength-dependent, represent the sensitivity of the detection channel for the emission polarizer oriented in the H- and V-directions, respectively. The kfl parameter is proportional to the square of the projection of the into the emission polarizer direction, i.e. fluorescent transition dipole moment (M) 2 2 MX or MY . So the vector M (with components Mx , My , and Mz in the xyz sample coordinates) has to be projected in the X and Y detection coordinates, leading to expressions: kfl (X)αMX 2 = [Mx cos (xX) + My cos (yX) + Mz cos (zX)]2

(9)

kfl (Y)αMY 2 = [Mx cos (xY) + MY cos (yY) + MZ cos (zY)]2

(10)

Angles xX, yX, zX, xY, yY, and zY are related with the Euler angles α, β, γ by means of [88]: cos (xX) = cos α cos β cos γ − sin α sin γ

(11)

cos(yX) = − sinα cosγ − cosα cosβ sinγ

(12)

cos(zX) = cosα sinβ

(13)

cos(xY) = cosα senγ +sinα cosβ cosγ

(14)

cos(yY) = cosα cosγ − sinα cosβ sinγ

(15)

cos (zY) = sin α sin β

(16)

The Euler angles are defined as (Fig. 3a): angle γ is the rotation around the z-axis that brings the y-axis in the XY-plane; angle β is the rotation around the y-axis, which translates the z-axis to the Z-axis; and angle α is the rotation angle around the new z-axis that makes the x- and y-axis coincident with the X- and Y-axis, respectively. For the experimental set-up used to record the fluorescence anisotropy, changing the relative orientation between the normal to the surface (z-axis) and the detection direction (Z-axis) by twisting the sample around its y-axis by a rotation angle δ (Fig. 2), the Euler angles are given by: α=0

(17)

γ= 0

(18)

β= 180 − (22.5+δ)

(19)

8


z ψ

M

y x

φ

Fig. 3 (A) Euler angle to translate the sample xyz coordinates to the laboratory XYZ axes. (B) Polar coordinates for a fluorophore adsorbed with a preferential angle with respect to the layer normal

where 22.5º is the angle between the excitation and the emission channels in the front-face configuration of the SPEX fluorometer. On the other hand Mx , My , and Mz can be expressed in their polar coordinates (Fig. 3b) by means of Mx = M sin ψ cos φ

(20)

My = M sin ψ sin φ

(21)

Mz = M cos ψ

(22)

and the z-axis and φ is the angle formed between where ψ is the angle between M in the xy-plane (Mxy = M sin ψ) and the x-axis. Actually the ψ the projection of M angle represents the preferential orientation of the transition moment of the dye and the normal to the clay layer. Applying Eqs. 11–22, Eqs. 9 and 10 are transformed in: kfl (X) = M 2 [ sin ψ cos φ cos (22.5 + δ) + cos ψ sin (22.5 + δ)]2

(23)

kfl (Y) = M 2 [ sin ψ sin φ]2

(24)

Because the plane of clay layers is an isotropic medium, all the φ angles are equally probable and the average for all possible φ values (from 0 to 2π ) has to be considered: Then 2π 2π cos φ = cos2 φ dφ = 1 2 2

0

2π sin2 φ =

0 2π

0

0

sin2 φ dφ = 1 2

(25)

(26)


9

2π cos

φ|2π 0

=

cos φ dφ = 0

(27)

sinφ dφ = 0

(28)

0

2π sinφ|2π 0 = 0

then, Eqs. 23–24 are transformed to: kfl (X) = M 2 [1 /2 sin2 ψ cos2 (22.5 + δ) + cos2 ψ sin2 (22.5 + δ)] 2

kfl (Y) = M 2 [1 /2 sin ψ]

(29) (30)

from which the dichroic ratio expressed in Eq. 8 is transformed to 2

DHV =

SH 1 /2 sin ψ cos2 (22.5 + δ) + cos2 ψ sin2 (22.5 + δ) 1 / sin2 ψ SV 2

(31)

The SH /SV ratio is related with the instrument response to the H and V emission polarizer directions, generally called the G factor [62]. This can be calculated from the ratio of the recorded intensity for H- and V-orientation of the polarizer isemission is is is IHH = IVV IVH = of an isotropic system (a nonviscous dye solution), G ≡ IHV SV SH , recorded at identical conditions to those of the sample (in front face configuration by means of a 1-mm optical pathway cuvette) and for all orientations of the sample (δ angle from −20º to 50º). Then, Eq. 31 is transformed to: Dcor HV = DHV G =

1/ 2

2

sin ψcos2 (22.5 + δ) + cos2 ψsin2 (22.5 + δ) 1/ 2

2

sin ψ

(31)

Reordering it, the final expression is: 2 2 2 Dcor HV = 2 cot ψ + (1 − 2 cot ψ) cos (22.5 + δ)

(32)

Equation 32 predicts a linear relationship between the fluorescence dichroic ratio and the rotation angle δ, and from the corresponding slope, the preferential orientation angle ψ of the fluorescent dye with respect to the film normal can be evaluated. In a similar way, a linear relationship can also be established between the absorption dichroic ratio and the rotation angle [79]. In this case, absorption spectra were recorded for horizontal (AH ) and vertical (AV ) linearly polarized incident light, and the corresponding dichroic ratio (DHV (abs) ≡ AH /AV ) was correlated with the rotation angle δ of the sample around its vertical y-axis (Fig. 4). The recorded spectra are affected by the instrument response to the H- and V-polarized light (even if a depolarizer is included between the sample and the detector), and the absorption spectra

10


y

depolarizer

δ δ

film depolarizer

x

y.

z

Y’

film

x z

Z X’

Z’ •

incident polarizer

Y’

X’ incident polarizer

Fig. 4 Experimental setup to record absorption polarization

iso of an isotropic system (Aiso V and AH ) have to be recorded to correct the instrumental responses. The corrected absorption dichroic ratio [Dcor HV (abs)] is linearly related with the twisted angle δ by means of [79]

Dcor HV (abs) ≡

2 − 3sen2 ψ AH Aiso V sen2 δ =1+ iso AV AH sen2 ψ

(33)

from which, the preferential orientation angle of the chromophore with respect to the normal to the clay layer ψ can be evaluated.

3 Dye/Clay Systems: Film Characterization Clay minerals are aluminosilicate layered materials based on the condensation of an octahedral (O) Al2 O3 or MgO planar sheet and one or two planar tetrahedral (T) SiO2 sheets, giving rise to the so-called TO or TOT layers. Clay layers can achieve a net negative charge due to isomorphic substitutions of tetrahedral Si4+ (tetrahedral substitution) and/or octahedral Al3+ o Mg2+ (octahedral substitution) by other metals with lower valence. The negative charges are compensated by inorganic exchangeable cations which are located on the clay surfaces, conferring the cation exchange capacity (CEC) to clay minerals. These exchangeable cations induce the stacking of clay layers giving rise to the tactoidal structure of clay platelets, characterized by the interlayer space (Fig. 5). The tactical structure of clays is not a rigid network, since the interlayer space can be expanded. The swelling ability of the interlaminar space depends, among other factors, on the surface charge density, the nature of the exchanged cations and their hydration degree [94, 95]. Smectite-type clays, with a CEC in the 30–100 meq/100 g range, are ideal clay systems to accommodate cationic or polar organic materials. The sodium form (Na+ as exchangeable cation) of smectite clays is specially recommended for this purpose,


11

Fig. 5 Tactoidal structure of clay layered materials

because the high hydration in this cation provides high swelling properties of the interlayer space of these clays. In this study, the synthetic clay Laponite (Lap) is chosen because of the high chemical purity and the very low particle size (≈30 nm), providing very transparent colloidal suspensions and solid films [96]. The CEC of Lap is 73.3 meq/100 g. On the other hand, rhodamine 6G (R6G, Fig. 6) is a fluorescent cationic dye which can be adsorbed on clay systems by a simple cation exchange mechanism. The photophysical properties of this dye and its tendency to self-associate in concentrated solutions are sensitive to the environmental conditions [97–99]. Probably, R6G is the most used laser dye as the active media in tunable lasers, and the study of new environmental conditions to improve the lasing properties of the dye is of special interest in photonics. Moreover, the photophysics of this dye has been largely used as a molecular probe to study a multitude of molecular systems [100–102]. Several techniques are now applied to characterize dye/clay solid films [71]. The morphology and the organization of clay particles in supported films are

13.8 Å H N

H 11.5 Å

Fig. 6 Molecular structure of rhodamine 6G dye (the transition dipole moment for the S0 –S1 absorption and emission transitions is marked by a double arrow)

N

O

C2H5

C2H5 CH3

CH3 CaOOC2H5 9.0 Å

12


checked by atomic force microscope (AFM) and X-ray diffraction (XRD) techniques. Absorption and fluorescence spectroscopy are applied to characterize the different species of the dye (monomers, dimers, and higher aggregates) adsorbed in the interlayer space of clay films [103, 104]. Finally, the anisotropy response of the dye adsorbed in the clay layers is evaluated by absorption and fluorescence spectroscopy with linearly polarized light [56, 77–80]. In order to evaluate the preferential orientation of fluorescent dyes adsorbed in clay layers by conventional polarization spectroscopy, a macroscopic organization of the clay layers in parallel planes is required first. It has been demonstrated that the spin-coating technique is an adequate method to elaborate supported films with a parallel arrangement of stacked clay layers [96]. Indeed, topographical images of clay films obtained by the AFM technique (Fig. 7a) suggest smooth and homogeneous surfaces of the clay particles in the films. The XRD profile of a pure Lap film elaborated by spin coating (Fig. 7b) shows sharp peaks assignable to 00c diffractions (001, 002, . . .), the diffraction c-axis being the normal to the clay layer. These sharp peaks indicate an internal organization of the clay layers in parallel planes in the films. The peak with the lowest diffraction angle corresponds to the 001 diffraction, from which the interlayer space of stacked clays can be evaluated. For the specific case of the Lap clay, the lowest diffraction angle appears at 2θ = 6.9º, giving rise to a free interlayer space of 3.2 Å, after subtracting the thickness of the clay layer (9.6 Å for a TOT clay, as in the present case). This interlayer space is due to the size of the hydrated inorganic cations (Na+ in the present Lap clay) responsible for the stacking and swelling properties of the clay layers in aqueous suspensions. The thickness of the supported clay films can be modified by changing the experimental conditions used for their elaboration by means of the spin-coating

d001

(B)

counts

(A)

d004 d002 0

10

20

d006 30 40 2θ

50

60

70

Fig. 7 AFM taping images (scan: 2×2 μm2 ) (A) and X-ray diffractions (B) of a Lap film elaborated by the spin-coating technique [96]


13

technique. It can be accurately evaluated by the AFM technique, by simply scanning the depth of the cantilever passing through a groove specifically performed in clay films for this purpose. Supported films of Lap clay with thicknesses in the range of 50–500 nm were elaborated by spin-coating methods [96]. Films with different thickness were obtained by changing the rheological (viscosity) properties of the clay suspension used for the spin-coating process, the spinning velocities and the number of consecutive coatings applied in the same substrates [96]. Cationic organic dyes (Rhodamine 6G, R6G, in the present case, Fig. 6) can be incorporated in clay films by the cationic interchange mechanism. The organic cation displaces the interchangeable inorganic cation by simple immersion of the clay films into adequate solutions of the dye. The dye loading in the clay films can be modified by changing the nature of the solvent in the immersing dye solution, the dye concentration, the immersion time, and the immersion temperature [96]. The final amount of the intercalated dye can be evaluated by several quantitative analytical techniques such as elemental CHN analysis and thermogravimetry. The relative dye loading on the clay system is generally expressed in % CEC, the percentage of the adsorbed dye over the total CEC of the clay. Lap films with different loadings of R6G in the 0.1–60% CEC range were prepared by immersing supported thin Lap films into dye solutions (a water/ethanol mixture with a molar fraction of 0.8 in water) with dye concentrations in the 10−5 – 10−3 M range during 5 min to 48 h immersion times [96]. The 60% CEC loading represents the saturation for the adsorption of R6G in Lap films, and further increase in the immersion conditions (time and dye concentration) did not induce an augmentation in the dye content. Indeed, it has been demonstrated that Lap surfaces are totally covered by a monolayer of highly packed dye molecules at this 60% CEC loading [79]. The intercalation of the dye molecules into the interlayer space of Lap films was confirmed by the XRD technique (Fig. 8). The 001 diffraction peak shifts to lower angles (2θ from 6.9º to 4.2º) by increasing the dye content (up to 60% CEC), suggesting an increase in the interlayer distance with the amount of the intercalated dye. The free interlayer space changed from 3.2 Å, for the most diluted dye film (0.1% CEC), to 11.4 Å for the highest dye-loaded film (60% CEC). Taking into account the molecular dimensions of R6G (13.8×11.5×9.0, in Å Fig. 6), the intercalation of R6G molecules into the interlayer space of Lap films can be performed in different ways: (1) The dye molecules could be adsorbed with the xanthene ring parallel to the clay layers forming a monolayer structure of dye molecules. (2) The dye molecules could be adsorbed with the short molecular axis of the xanthene ring perpendicular to the clay layer. (3) The dye molecules could be adsorbed with the long molecular axis of the xanthene ring with an orientation angle with respect to the clay layers. In this case, a more perpendicular orientation should be expected as the packing of the dye molecules is increased as suggested by XRD data.

14

F. López Arbeloa et al. 60%

4 × 103 45%

counts

3 × 103

2 × 103

22%

10%

1 × 103

5.7%

2% 0.1%

0 2

3

4

5

6 2θ

7

8

9

10

Fig. 8 X-ray diffraction for low scan angles of R6G/Lap films with different R6G loadings (in % CEC) [96]

Absorption and fluorescence spectroscopy with linearly polarized light (Sect. 4) will allow a correct interpretation of the present results. The sharp 001 diffraction peak and the clear 002 diffraction peak observed for the highest dye-loaded film (60% CEC, Fig. 8) suggest that the internal organization of the stacked clay layers in the films is not disrupted by the intercalation of the dye molecules into the interlaminar space of the clay. On the other hand, thermogravimetric TG and DTA curves (data not shown) indicate a higher thermostability of R6G when it is adsorbed in the Lap films [57, 105]. Indeed, pure R6G thermodegradates at around 330ºC, whereas the degradation of R6G in Lap films starts in the 390–420ºC temperature range [57, 105–107]. Consequently, the clay matrix protects dye molecules against thermal degradations, improving the operative lifetime in optoelectronic devices based on the photoresponse of R6G. The increase in the dye loading not only induces the expected increase in the red color of the R6G/Lap films (owing to an augmentation of the dye concentration) but also changes in the color tonality, observed by the naked eye. This last observation suggests new absorbing species formed in high dye contents. Indeed, the shape of the normalized absorption spectra of R6G in Lap films (normalized to the area under the curve, Fig. 9a) changes with the dye loading [103]. The absorption spectrum of R6G in Lap films for very low dye concentration (Fig. 9a, curve a) consists of the typical absorption spectrum of R6G in diluted solutions, with a main narrow absorption band centered at 528 nm and a vibronic shoulder at higher energies (at around 495 nm). Increasing the dye loading, new absorption bands appear at shorter and longer wavelengths of the main absorption band. New absorption bands in concentrated dye systems are generally assigned to the molecular aggregation [108–110]. Thus, the adsorption of R6G molecules into the clay surface facilitates the encounter of dye molecules leading to their selfassociation.


(A)

15

(B)

a

M

0.03

D

0.03

absorbance

absorbance

J-band 0.02 j

0.02 H-band 0.01

0.01

0.00 450

500 550 wavelength (nm)

0.00 450

600


600

Fig. 9 (A) Area-normalized absorption spectra of R6G in Lap films for different dye loadings (in % CEC): 0.1 (a), 1.0 (b), 2.0 (c), 3.2 (d), 5.7 (e), 10 (f), 14 (g), 21 (h), 45 (i), and 60 (j). (B) Average absorption spectrum of R6G dimers in Lap films calculated in the 2–25% CEC dye-loading range [103]

Molecular aggregation can be explained by the exciton theory [111, 112], a quantum mechanics method based in the dipole–dipole interaction of the monomeric units. This theory proposes the splitting into two excited states of the dimer: one with lower energies and the second with higher energies than the monomer excited state (Fig. 10). Depending on the geometry of the aggregates, the transition moments from the ground to the excited states can be allowed or forbidden: – Sandwich dimers and higher-order aggregates with a parallel association of the chromophoric aromatic rings (Fig. 10, case A): These aggregates are characterized by an absorption band placed at higher energies with respect to the monomer

case D1

E2

case A

ΔD

E1 ν2

case C

case D2

caseB

α M

θ = 90º >

E2

θ

α

θ = 0º

>

M

E2

ΔD

E1

E1 ν1

νM

νD

νD

νΜ

ν2

ν1

θ = 54.7º H-type dimers

J-type dimers

Fig. 10 Exciton splitting of the electronic excited states of dimers predicted by the exciton theory. The photophysical properties of these excited states for different aggregate geometries are also illustrated

16


absorption band (the so-called H-bands) and they are not fluorescent but efficient quenchers for the monomer fluorescent emission via an ET deactivation. – For collinear head-to-tale aggregates (Fig. 10, case B): These dimers are characterized by absorption and fluorescent bands placed at lower energies than the monomer spectral bands (the so-called, J-bands). – Intermediate geometries for the aggregates are possible: (1) coplanar displaced aggregates (Fig. 10, case C) presents a H- or J-absorption band depending on the displacement angle θ (H-band for short displaced dimers, θ > 54.7º, and J-band for long-displaced dimers, θ < 54.7º); (2) twisted sandwich aggregates (Fig. 10, case D1 ) with two absorption H- and J-bands, the band placed at higher energies being more intense; and (3) oblique head-to-tale aggregates (Fig. 10, case D2 ), with the two absorption bands active, the J-band which is also potentially fluorescent being more intense. The absorption spectrum of the dimer cannot be experimentally recorded because of its coexistence with monomers and higher-order aggregates. It can be recorded by subtracting the proportional contribution of the monomer absorption band from the registered spectra in moderated dye concentrations. A mathematical procedure has been developed, based on the Mass Law, to evaluate the absorption spectrum of the dimer in solid host materials with moderate dye contents [103]. The absorption spectrum of the R6G dimer in Lap films is calculated applying the Mass Law to the evolution of the absorption spectra of R6G/Lap films with a dye loading for moderate dye content (1–25% CEC), for which a clear isobestic point at around 509 nm in the area normalized spectra is observed, Fig. 9a. A detailed description of the mathematical procedure to evaluate the dimer absorption spectrum is described elsewhere [103]. The average absorption spectrum for the R6G dimer in Lap films (Fig. 9b) consists of two absorption bands centered at 545 (J-band) and 503 nm (H-band), with the J-band being around 3/2 more intense than the H-band. This result suggests the presence of either an oblique head-to-tale dimer or the presence of two coplanar displaced dimers: one, with an important contribution, corresponds to a long-displaced coplanar dimer (responsible for the absorption J-band), and a second minor dimer corresponds to a short-displaced coplanar dimer (absorption H-band). It has been observed that the intensity of the J-absorption band slightly decreases by increasing the dye loading [103], suggesting a more oblique head-to-tale dimer or a less favored long-displaced coplanar dimer in Lap films as the dye content is augmented. For high dye concentrated R6G/Lap films (>25% CEC) the loss of the isobestic point and new more energetic bands in the area normalized absorption spectra (Fig. 9a, curves i and j) should indicate the formation of trimers or higher-order aggregates. These aggregates would be characterized by more shifted H- and J-absorption bands, at ca. 475 and 550 nm, respectively. The formation of R6G aggregates is corroborated by fluorescence spectroscopy. Aggregates with J-type spectral bands are confirmed by the important bathochromic shift observed in the emission spectra of R6G/Lap films by increasing the dye content (Fig. 11) [104]: from 548 nm for the most diluted dye film, 0.1% CEC, up to


a

1000

fluorescence intensity (a.u.)

Fig. 11 Height-normalized fluorescence spectra of R6G in Lap films for different dye loadings (see caption for Fig. 9a for more information). Inset: evolution of the fluorescence efficiency, analyzed by the fluorescence area over the absorbance at the excitation wavelength, with the dye content [104]

17

Ifl/Aexc

i

% CEC

800

0

5

10

15

20

600

400

200

0 520

540

560

580

600

620

640

wavelength (nm)

596 nm for the most concentrated dye film, 60% CEC. On the other hand, the drastic reduction in the fluorescence efficiency (analyzed as the fluorescence area over the absorbance at the excitation wavelength, Ifl /Aexc ) with the dye concentration (inset Fig. 11) proves the presence of H-type aggregates, since they are not fluorescent and are potential quenchers of the fluorescent emission of monomer and J-type aggregates. This quenching effect is also corroborated by time-resolved fluorescence spectroscopy (data not shown), with an important decrease in the average fluorescence lifetime of the monomer emission by increasing the dye content [104]. On the other hand, a bathochromic shift of the time-resolved emission (TRES) band was observed for longer delay times between the excitation pulses and the detected emission photon [104], suggesting longer lifetimes for fluorescent species emitting at longer wavelengths (J-aggregates). It is difficult to establish the emission maximum for the R6G J-type dimer and higher-order aggregates because of overlapping of the fluorescent bands and because the recorded fluorescence spectra show a continuous shift rather than the observation of new fluorescent bands. It should be noted that the fluorescence spectra were recorded after excitation at 495 nm, at the vibronic shoulder of the monomer absorption band and the H-absorption band of R6G aggregates. In any case, fluorescence properties of R6G/Lap films are further analyzed in the next section using polarization spectroscopy. To summarize, R6G molecules can be intercalated into the interlayer space of Lap films by a simple ion exchange mechanism. Depending on the dye loading, different R6G species are identified,: for very low loading ( 40% CEC).

4 Dye Orientation in Ordered Clay Films. A Fluorescence Anisotropy Study In this section we demonstrate the validity of fluorescence anisotropy to evaluate the orientation of fluorescent probes adsorbed in ordered bi-dimensional surfaces. This methodology is applied to the specific case of supported R6G/Lap films. In order to apply conventional spectroscopic methods to evaluate the orientation of dye molecules adsorbed on clay layers, it is necessary to dispose of a macroscopic distribution of ordered clay particles, i.e. parallel stacking of the clay layer into a supported glass substrate. Parallel arrangements of clay layers can be obtained by the elaboration of thin films by the spin-coating technique [96]. This parallel stacking of clay layers is confirmed by absorption spectroscopy with linearly polarized light, a well-established polarization technique extensively used to evaluate the orientation of dye molecules in clay films [85, 113–115]. Figure 12 shows the absorption spectra of the most dye-diluted R6G/Lap film (0.1% CEC) recorded for horizontal (H) and vertical (V) polarized light as a function of the rotation angle δ of the film around its vertical y-axis (see experimental set-up in Fig. 4). These spectra were corrected for the instrumental response to the linearly 0.06

absorbance (a.u.)

A: V-polarized

B: H-polarized

0°

0.04 70° 0.02

0.00 450


450


600

Fig. 12 Corrected absorption spectra light of the 0.1% CEC R6G/Lap film for V-(A) and H-(B) polarized incident light as a function of the twisting angle δ around its vertical axis [79]


19

polarized light by recording the detected signal for an isotropic system (a diluted R6G solution in ethanol). The corrected V-polarized absorption spectrum of R6G monomers does not show any evolution with the twisting angle δ (Fig. 12a). This observation is a consequence of the isotropic behavior of the clay surface (there is not a preferential orientation in the R6G adsorption in the plane of the clay layer and all the φ angles in Eqs. 23–28 are equally probable). Experimental results suggest that this isotropic behavior is maintained for the macroscopic arrangements of clay layers and confirms the organized stacking of the clay layers (parallely disposed with respect to the supporting substrate) in the films. Indeed, a random distribution of the dye molecules in the clay layers, with a preferential orientation with respect to the layer normal, would induce a macroscopic average transient dipole moment oriented perpendicular to the clay layer (out-of-plane, along the z-axis of the sample, Fig. 3b), without any component in the xy-plane of the clay layer. Consequently, this perpendicular component of the macroscopic transition dipole does not change its relative orientation with respect to the plane of the vertical polarized light by twisting the films around its vertical axis (Fig. 4). This is not the case for the horizontally polarized light. In this case, twisting the sample around the vertical axis will imply a reorientation of the macroscopic transition moment in the polarization plane of the incident light. This is experimentally confirmed. The intensity of the absorption spectrum of R6G monomers in Lap films decreases with the twisting angle δ for H-polarized light (Fig. 12b), suggesting an anisotropic behavior of dye molecules with respect to the clay normal. Fluorescence spectra with linearly polarized light of the monomeric unit of R6G in Lap films (recorded for the 0.1% CEC R6G/Lap film) are shown in Fig. 13[77]. The fluorescence spectra were recorded for different rotating angles of the sample around its vertical y-axis (−20 < δ < 50º, see experimental set-up in Fig. 2) with the emission polarizer oriented in the horizontal [H, Fig. 13 (left)] and vertical [V, Fig. 13 (right)] for a constant V- [Fig. 13 (top)] or H-polarized [Fig. 13 (bottom)] excitation light. All these spectra were corrected for the instrumental response to the linearly polarized light by means of the detected fluorescence signal of an isotropic sample (a diluted solution of R6G in ethanol in a 1-mm optical pathway quartz cuvette) recorded under identical experimental conditions (including the rotation angle δ). Independently of the excitation polarization, the fluorescence intensity for the V- and H-polarized emission polarizer, IVV and IVH (or IHV and IHH ), increases and decreases by increasing the rotation angle δ of the film, respectively. This opposite evolution for both emission polarizations is a consequence of the anisotropic behavior of the dye/clay films with respect to the normal to the clay layer, and therefore, this fluorescence anisotropy can be used to evaluate the preferential orientation of fluorescent dye in ordered clay systems. Figure 14 shows the corrected fluorescence dichroic as a function of the emission wavelength for different δ angles, obtained for V-polarized excitation light (similar results were obtained for excitation with the H-polarization light [77, 78]). Except at the shortest detection wavelengths (where the important polarization of

20



6 × 105

IVV

IVH

50°

5 × 105 –20°

–20° 4 × 105 3 × 105

50°

2 × 105 2 × 105

ifluorescence intensity (a.u.)

0 2.0 × 106 IHH

–20°

IHV 50°

1.5 × 106 50° 1.0 × 106

–20°

5.0 × 105

0.0 520 540 560 580 600 620 wavelegnth (nm)

520 540 560 580 600 620 640 wavelegnth (nm)

Fig. 13 Corrected fluorescence spectra of 0.1% CEC R6G/Lap films for emission polarizer oriented in the H-(right) and the V-(left) direction after excitation at 495 nm with H-(bottom) and V-(top) polarized light as a function of the twisting angle δ [77, 78]

the scattered light by the clay particles close to the excitation wavelength (495 nm) cannot be completely corrected), the fluorescence dichroic ratio is independent of the emission wavelength, suggesting the presence of a unique fluorescent species, the R6G monomer in this very dye diluted R6G/Lap film. The graphics inserted into Fig. 14 illustrate the good linear relationship (with correlation coefficients 2 close to 1) between Dcor HV and cos (22.5+δ), according to Eq. 32. From the slope of these relationships, a preferential angle ψ = 62º is obtained for the adsorption of R6G monomer with respect to the normal to the Lap layers. This value, obtained at 548 nm (R6G monomer emission maximum), independently of the excitation polarization, is also obtained for other emission wavelengths (far away from the excitation wavelength).

Fluorescence Anisotropy in 2D Ordered Systems 1.6

(A)

1.2

λfl= 548 nm r = 0,9988 ψ = 62º

cor

0.8

1.4

DHV

0.6

cos2(22.5+α)

Dcor HV

Fig. 14 Evolution of the corrected fluorescence dichroic ratio of the 0.1% CEC R6G/Lap film with the twisting angle δ obtained after excitation at 495 nm with V-polarized light. Inset: linear relationship of the dichroic ratio with the twisting angle analyzed at 548 emission wavelength [77]

21

1.0

0.0

0.2

0.4

0.6

0.8

1.0

–20º 0.8

0.6

50º 0.4 540

560

580

600

620

640

wavelenngth (nm)

Recently, we observed that this fluorescence anisotropy methodology can also be applied for excitation with unpolarized light [116], just by recording the fluorescence intensity for two orthogonally oriented emission polarizations after conventional excitation. The use of unpolarized excitation is recommended because the method is more sensible (polarized excitation light implies a loss of the excitation intensity and hereafter of the emission intensity) and more precise (the effect of the light scattering polarization is reduced). An identical ψ value was previously obtained for R6G monomer adsorbed in Lap films by absorption spectroscopy with linearly polarized light (Fig. 12) [79]. Indeed, and according to Eq. 33, an orientation angle ψ = 62º was obtained from the slope 2 of the absorption dichroic ratio Dcor HV (abs) vs. sin δ (data not shown) [54]. The fact that both absorption and fluorescence anisotropies provide the same value for the orientation angle of R6G in Lap films confirms the validity of the new fluorescent method versus the well-established method based on the absorption with linearly polarized light. The new fluorescent method has the advantages to be a more sensible and selective technique than absorption spectroscopy. Moreover, since identical ψ values are obtained by both anisotropy methods, this means that all the required conditions needed to develop Eq. 32 are valid, at least for dye/clay films with very low dye contents as in the present case (0.1% CEC). For instance, the absence of any reorientation of the emission transition moment during the fluorescence lifetime or the absence of any excitation ET phenomena to other fluorophores with different orientation (at least along the film normal). The orientation of R6G dimers and higher-order aggregates in Lap films was studied by absorption and fluorescence anisotropies for some representative

22


R6G/Lap films (in the 2–25% range) for dimers and 40% and 60% CEC for higher aggregates. Figure 15 shows the most illustrative results for the fluorescence spectra with the H-oriented emission polarizer of two representative samples of R6G/Lap film: 5.7% (Fig. 15a) and 22% CEC (Fig. 15b) [80]. In the 5.7% CEC sample, the emission is mainly due to the monomer units (λfl ≈ 550 nm), which are the main adsorbed species for this relatively low dye-loaded sample. However, the 22% CEC sample provides a better characterization of the fluorescence J-band of R6G dimers in Lap films (Fig. 15b). In both samples an increase in the rotation angle δ induces a diminution in the fluorescent emission of the monomer and a slight augmentation in the fluorescence intensity at the lowest energetic part of the emission band. Thus, the evolution of the fluorescence dichroic ratio with the rotating angle δ [Fig. 15 (top)] presents two different behaviors depending on the emission wavelength: The Dcor HV value decreases and increases by increasing the rotating angle δ for the wavelength intervals λfl < 580 and λfl > 580 nm, respectively. The Dcor HV vs. cos2 (22.5+δ) linear relationship [inset Fig. 15 (top)] provides orientation angle ψ = 61º for the monomer fluorescence (analyzed at 550 nm) and ψ value around 49º for the emission from the J-type dimer (analyzed at 630 nm, far away from the influence of any emission of monomers) for both 5.7% and 22% CEC R6G/Lap films.

(A): 5.7% CEC

cor

DHV

–20


50 IHH

2.0 1.5

2 × 106 50 50 –20

0 520 540 560 580 600 620 640 660 wavelength (nm)

1.4

Dcor HV

λ fl = 550 nm Ψ = 61°

1.0 0.0

fl = 630 nm cos 2 (δ +22.5) Ψ = 50° 1.0 0.2 0.4 0.6 0.8

1.0 2.0 × 105

-20

(B): 22% CEC 1.6

1.2

–20

1

1 × 106

2.5

50

2

3 × 106

3.0

fluorescecne intensity (a.u.)

cor

DHV

3

1.5 × 105

IHH

50

–20

1.0 × 105 50 5.0

× 105

–20

0.0 520 540 560 580 600 620 640 660 wavelength (nm)

Fig. 15 Corrected fluorescence spectra of 5.7 (A: left) and 22 (B: right) % CEC R6G/Lap films for emission polarizer oriented in the H-direction after excitation at 495 nm with H-polarized light as a function of the twisting angle δ. The evolution of the corrected fluorescence dichroic ratio with the twisting angle δ is also included (top). Inset (top): linear relationship of the dichroic ratio vs. the twisting angle δ analyzed at two emission wavelengths: 550 (monomer) and 630 nm (J-type aggregate) [80]


23

However, these results have to be read carefully. Indeed, at the excitation wavelength used to record these emission spectra (λexc = 495 nm), both monomers and H-type dimers are promoted to their corresponding excited states (see Fig. 9b). Consequently, the fluorescent emission from the J-band is only possible after an ET process from those excited states directly populated in the excitation (monomer and H-band dimer). Therefore, the fluorescence anisotropy of the J-dimer should be affected for the fluorescence depolarization of the excitation light produced by this ET process. In order to evaluate the preferential orientation of the R6G dimers, the linear polarized light was applied to the absorption spectrum of the dimer calculated from moderate dye content (1–25% CEC) R6G/Lap films [80]. Figure 16 shows the evolution of the H-polarized absorption spectrum of the R6G dimer (after subtracting the proportional absorption spectrum of the monomer) with the rotation angle δ. The absorbance of the J-band decreases by increasing the rotating angle whereas the opposite behavior is observed for the H-absorption band. These opposite evolutions are clearly reflected in the absorption dichroic ratio [Fig. 16 (top)] showing a

3.0 λab = 495 nm

cor

1.5 DHV

DHV

cor

2.5 2.0

ψ = 48° λab = 550 nm

1.0

ψ = 60°

70° 1.5

2

0.0

0.2

sin δ 0.4 0.6

0.8

1.0

0° 1.0 0° 0.5 0.020

H-polarized 0°

0.015 absorbance

Fig. 16 Corrected absorption spectra of the R6G dimer in Lap (calculated for the 22% CEC R6G/Lap) recorded for H-polarized light as a function of the twisting angle δ. The evolution of the corresponding dichroic ratio with the twisting angle δ is also shown (top). Inset: linear relationship of the dichroic ratio vs. the twisting angle δ analyzed at two absorption wavelengths: 500 (H-band) and 550 nm (J-band) [79]

70

70°

0.010 70

0.005 0 0.000 450


600

24


dependence on the absorption wavelength. These evolutions imply positive and neg2 ative slopes in the linear representation of Dcor HV (abs) vs. sin δ [inset Fig. 16 (top)], giving rise to different orientation angles for the transition moment of the dimeric H- and J-absorption bands with respect to the clay layers of ψ = 48º and ψ = 61º, respectively. Several aspects can be drawn from these experimental data: – The orientation angle ψ = 61º obtained by absorption for the J-absorption band of the dimer is quite similar to that obtained for the R6G monomer by fluorescence anisotropy. These results suggest that the transition moment of both R6G species are oriented with similar angles with respect to the clay layers. – However, the orientation angle ψ = 49º reported for the fluorescence J-band of the dimer does not match that reported from the absorption anisotropy of the J-band but rather matches the ψ = 48º value obtained for the H-band by absorption anisotropy. These results suggest the above-mentioned excitation ET from the excited H-state to the fluorescent J-state of the dimer, as is discussed below. – The fact that the ψ angles for H- and J-bands of the dimers are not complementary to each other would indicate that the H- and J-adsorption bands do not correspond to the two absorption bands of a unique dimer (twisted sandwich or oblique headto-tail dimers).1 Present results suggest that the two H- and J-absorption bands of the aggregate are actually due to two different displaced coplanar dimers (see Fig. 10, case C). These coplanar dimers are not disposed parallel with respect to the clay layer but form a titled arrangement respect to the normal to the surface. The J-adsorption has to be assigned to a long displaced coplanar dimer (Fig. 10, case C with θ < 54.7º) which should be potentially fluorescent. Effectively, the absorption anisotropy proposes an orientation angle ψ = 61º with respect to the clay normal for this aggregate, this means a θ = 29º with respect to the clay layer. On the other hand, the H-absorption band of the aggregate, with an orientation angle ψ = 48º, should be related to a short displaced coplanar dimer (Fig. 10, case C with θ > 54.7º). As has been mentioned above, the discrepancies in the ψ value obtained for the J-type dimer by fluorescence anisotropy (ψ = 48º) and absorption anisotropy (ψ = 61º) can be assigned to the ET between different excited states. In the present case, the excitation at 495 nm directly populates the excited state of the short displaced coplanar dimer and, in order to observe the fluorescence from the J-type long-displaced coplanar dimer, an ET process has to be involved, as is illustrated in Fig. 17a. This ET effect can be minimized by increasing the excitation wavelength to a value in which the J-type dimer is the main absorbing species. In fact, it has been experimentally observed [80] that the dichroic ratio of the fluorescence of the 1 The dipole moment of the two excited states of the dimer would be the vectorial addition (algebraic addition and subtraction) of the individual dipole moment of the monomers. Such addition and subtraction will provide orthogonal dipole moments (see Fig. 10).


25 1.6

θ > 54.7° kkET

Ψ = 60°

1.4

ν(D ν(DJJ)

H-dimer -

J-dimer -

excitation

emission

cor

ν(D ν(D HH)

θ

λ exc= 545 nm Ψ = 56°

D HV

θ

λ exc = 575 nm

θ < 54.7°

1.2 1.0

λ exc = 495 nm Ψ = 49°

0.8 0.0

0.2

0.4

0.6

0.8

1.0

2

COS (δ + 22.5)

Fig. 17 (A) Schematic representation for the excitation energy transfer from the excited state of the short-displaced collinear dimer (H-absorbing band) to the fluorescent excited state of the long-displaced collinear dimer (J-fluorescent band). (B) Effect of the excitation wavelength on the linear relationship between the fluorescence dichroic ratio of the J-dimer (analyzed at the emission wavelength of 630 nm) and the twisting angle δ [80]

J-type dimer depends on the excitation wavelength (Fig. 17b). The orientation angle 2 derived from the slope of the linear relationship between Dcor HV vs. cos (22.5+δ) at different excitation wavelengths are: ψ = 49º for λex = 495 nm, ψ = 56º for λex = 545 nm, and ψ = 60º for λex = 575 nm. The excitation wavelength λex = 545 nm corresponds to the maximum absorption for the J-type dimer (Fig. 9b), but minor proportions of H-dimers and monomers are also excited, and the obtained ψ = 56º value is an average value for the orientation angle of all these species. However, for an excitation at 575 nm, only R6G J-type dimers are directly excited, and the fluorescence anisotropy reported from their emission directly provides its orientation angle. The orientation angle for the J-dimer evaluated by fluorescence polarization, ψ = 60º practically reaches the value reported by absorption anisotropy ψ = 61º. In order to study the dynamic of the ET process, fluorescence decay curves with linearly polarized light were registered. Figure 18 shows the decay curves for the emission from a J-dimer (at 620 nm) after excitation at 375 nm (the wavelength of the pulsed laser used in the excitation channel). The decay curves were recorded for H- and V-oriented emission polarizer after V-polarized excitation, and were corrected for the instrumental response to the plane of the linearly polarized light, by means of the recorded fluorescence decay curves of an isotropic system (a nonviscous R6G liquid solution) obtained under identical instrumental conditions (front-face configuration using a 1-mm optical pathway cell). The experimental setup is different to that used to record fluorescence spectra. For instance excitation and detection channels are oriented in a right-angle configuration, and the films were twisted with respect to both directions. The corresponding time-dependent dichroic ratio [Fig. 17 (top)] shows a constant value during the lifetime of the emitting species. This means that the ET process to the fluorescent J-dimer is a very rapid process, much faster than 50 ps, the time resolution of the time-correlated single photon counting instrument. In this case, the ET process is so rapid that the fluorescence anisotropy of the J-dimer is dominated by the polarization of the excitation light by the H-type aggregate (the prerequisite condition for collinear excitation and emission transition moments derived by Eq. 32 is no longer valid). Consequently,

26

D HV

cor

4 2 0 4 IVH lg(counts)

Fig. 18 Fluorescence decay curves of 22% CEC R6G/Lap films recorded at the fluorescence J-band of the dimer for H- and V-emission polarizer after excitation with V-polarized light. The time evolution of the corresponding corrected dichroic ratio is also shown (top)


3

IVV

2

1

0 0

10

20 time (ns)

30

40

the orientation angle ψ = 49º provided by the fluorescence anisotropy of the J-dimer after excitation at 495 nm is actually the orientation of the directly populated species in the excitation process, the H-type dimer in the present case. These results suggest that, depending on the experimental conditions, the fluorescence anisotropy can provide the orientation angle of the fluorescent species or the excited species. Moreover, if the dynamic of the ET is competitive with respect to the fluorescence lifetime, then fluorescence anisotropy could supply information related to the relative orientation between the donor and the acceptor moieties. Because of the efficient ET in R6G/Lap films and because of the presence of many absorbing and emitting species with high overlapping spectral bands, the application of fluorescence anisotropy to evaluate the preferential orientation of R6G molecules in Lap films is drastically reduced in moderated–high dye-loaded samples. Thus, it was not possible to adequately study the anisotropy for the 60% CEC film by fluorescence polarization, not only due to the presence of several absorbing and emitting species with several ET interconnected processes, but also because of the reminiscent emission of this sample (fluorescence efficiency is three orders of magnitude lower than that for R6G/Lap films with low dye loadings) [104]. Therefore, in this case the preferential orientation of R6G molecules was evaluated by absorption anisotropy [79]. Figure 19 illustrates the important anisotropy observed at absorption spectra of the 60% CEC R6G/Lap films at wavelengths close to 475 nm. This high anisotropy reveals a nearly perpendicular disposition of the absorption transition moment of this higher-order aggregate with respect to the clay layers. In 2 fact, the slope of the linear evolution of Dcor HV (abs) vs. sin δ [inset Fig. 19 (top)]

Fluorescence Anisotropy in 2D Ordered Systems 10 6

D HV

λ ab = 475 nm

cor

8 DHV

Fig. 19 Absorption spectra of 60% CEC R6G/Lap recorded with H-polarized light. The evolution of the corrected dichroic ratio with the twisting angle δ is also included (top) [80]

27

ψ = 28°

4

65

λ ab = 550 nm

6

2

ψ = 60° 2

4

0 0.0

0.2

sin δ 0.4 0.6

0.8

1.0

2 0

0 1.2 AH

65°

absorbance

1.0

0.8

0.6

0.4

0.2 0° 0.0 450


provides an orientation angle ψ ≈ 28º for this short-displaced coplanar higher-order aggregate. Table 1 summarizes the orientation angle evaluated from absorption and fluorescence anisotropies for several R6G species characterized in R6G/Lap films with different dye contents. From these results, it can be concluded (Fig. 20) [57]: 1. The intercalation of R6G dye in Lap films leads to a macroscopic arrangement of the dye molecules with a preferential orientation of the adsorbed dye molecules with respect to the normal of the clay layers. 2. Monomer units are adsorbed with a preferential orientation of about 62º, with the long molecular axis of the dye closely lying in the clay layers. 3. Such an orientation is maintained in R6G/Lap films with high dye contents. The packing of monomeric units with this orientation by increasing the dye loading

28


Table 1 Orientation angle ψ derived from absorption and fluorescence spectroscopy with linearly polarized light for several R6G species adsorbed in Lap films at different dye contents (in % CEC) Absorption polarization

Fluorescence polarization

% CEC

adsorbed species

ψ (º)

adsorbed species

0.1

Monomer (λab ≈ 530 nm)

62

Monomer (λfl ≈ 550 nml ) 62

5.7

H-dimer (λab < 500 nm) J-dimer (λab > 510 nm)

48 61

Monomer (λfl ≈ 550 nm) J-dimer (λfl > 600 nm)

61 49 (λexc = 495 nm)∗ 56 (λexc = 545 nm)∗ 60 (λexc = 570 nm)

22

H-dimer (λab < 500 nm) J-dimer (λab > 510 nm)

46 60

Monomer (λfl ≈ 550 nm) J-dimer (λfl > 600 nm)

≈ 60 50 (λexc = 495 nm)∗ 53 (λexc = 545 nm)∗ 62 (λexc = 570 nm)

40

H-band (λab < 500 nm) J-band (λab > 510 nm)

32 60

60

H-band (λab < 500 nm) J-band (λab > 510 nm)

28 60

J-band (λfl > 600 nm)

50 (λexc = 495 nm)∗ 52 (λexc = 545 nm)∗ 50 (λexc = 570 nm)∗

∗ Values

ψ (º)

affected by fluorescence depolarization

0.1% CEC

monomer ψ = 62º

1 – 25% CEC

H – dimer ψ = 48–50º

J – dimer ψ = 61º

> 40% CEC

H – aggregates J – aggregates ψ = 60º ψ = 32 – 28º

Fig. 20 Schematic representation for the preferential orientation of R6G molecules adsorbed as monomer, dimers, and higher-order aggregates (short- and long-displaced collinear aggregates) in ordered Lap films as a function of the dye content [57]

induces the formation of long displaced coplanar aggregates, characterized by J-type absorption and fluorescence bands placed at lower energies with respect to the monomeric spectral bands. 4. Simultaneously, other dye molecules are intercalated into the interlayer space of Lap films in a more perpendicular configuration, forming short displaced coplanar dimers and higher-order aggregates. Increasing the dye loading, these aggregates are disposed more to the perpendicular, probably for a more compact packing of the adsorbed molecules. These aggregates are characterized by different H-type absorption bands. They are not fluorescent but act as efficient deactivation for the fluorescent emission of the monomer and J-type aggregates.


29

5 Conclusions Fluorescence polarization is a good strategy to evaluate the preferential orientation of fluorescent dyes adsorbed in ordered layered materials, such as clay minerals in supported films elaborated by the spin-coating technique. In analogy with the wellestablished anisotropy by absorption spectroscopy with linearly polarized light, the fluorescence method is based on the evolution of the emission dichroic ratio with the twisting angle of the film with respect to the excitation beam. Selecting the proper experimental conditions, the fluorescence anisotropy can supply the tilted angle of different fluorescent species (monomer and J-type dimers and higher-order aggregates) adsorbed into the clay layers. Moreover, if a very efficient and rapid ET process populates the fluorescent species, then the fluorescence anisotropy can provide the tilted angle of nonfluorescent species absorbing at the excitation wavelength by means of the depolarization of the excitation light. Even more, if the ET process competes with the fluorescence lifetime, then the time-dependent fluorescence dichroic ratio could provide the dynamic for this excitation ET. The new fluorescence method can be applied to study the preferential orientation of fluorescent systems in ordered rigid 2D arrangements. Acknowledgments The University of the Basque Country is thanked for financial support (GIU06/80). V.M. thanks the Spanish Ministerio de Educación y Ciencia for a postdoctoral grant.

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Room Temperature Tryptophan Phosphorescence of Proteins in the Composition of Biological Membranes and Solutions Vladimir M. Mazhul’, Alexander V. Timoshenko, Ekaterina M. Zaitseva, Svetlana G. Loznikova, Inessa V. Halets, and Tatsiana S. Chernovets

Abstract The room temperature tryptophan phosphorescence (RTTP) technique allows studying slow internal dynamics of proteins in the millisecond and second diapasons. This chapter summarizes the key findings in the field of RTTP spectroscopy, physical nature of this phenomenon, and experimental approaches to analyze the microenvironment of tryptophan residues. Representative examples of RTTP of proteins in human erythrocyte membranes and plant lectins in solutions are discussed in details taking into account the effects of detergents on biological membranes and 3D structures of lectin molecules, respectively.

1 Introduction The regulation of almost all processes occurring in the biosphere at the molecular level is related to the functioning of proteins. The functional activity of proteins is determined in many respects by their structure – the conformation and the internal dynamics. Spectroscopic methods based on a registration of the spectral, temporal, and polarization parameters of the fluorescence of tryptophan residues [1–3] are widely used in the investigation of the structure of proteins. However, the most suitable means for the study of the internal dynamics of proteins is provided by the room temperature tryptophan phosphorescence (RTTP). As compared to the fluorescence, the RTTP monitors much slower processes, extending the observation time from the nanosecond range of fluorescence up to millisecond to second range of phosphorescence. The RTTP spectra of proteins in the condensed state were detected for the first time in 1966 [4]. The RTTP of proteins in solution was first described in 1974 [5]. The RTTP spectra and decay kinetics of two globular proteins – horse-liver alcohol dehydrogenase and Escherichia coli alkaline phosphatase – were measured in

V.M. Mazhul’ (B) Laboratory of Proteomics, Institute of Biophysics and Cellular Engineering, National Academy of Sciences of Belarus, 27 Academicheskaya Str., Minsk, 220072, Belarus e-mail: [email protected] C.D. Geddes (ed.), Reviews in Fluorescence 2008, Reviews in Fluorescence 2008, DOI 10.1007/978-1-4419-1260-2_2, C Springer Science+Business Media, LLC 2010

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oxygen-pured solution. The RTTP lifetime (τ ) of proteins was found to be sensitive to changes in pH and solvent viscosity, to the action of guanidine hydrochloride (1 M) and ligand binding. It was suggested that the efficiency of quenching of RTTP depends on the fluctuations in the structure of the protein. In 1976, the spectra and RTTP decay kinetics of the suspensions of isolated human erythrocyte membranes, chick embryo fibroblasts, and yeast microorganisms Candida utilis were described [6]. It was established that RTTP decay of isolated membranes and fibroblasts changes under the action of noradrenalin (10–4 M) and calcium cations (10–2 M). In that work the capability of horse-liver alcohol dehydrogenase in solution to exhibit RTTP was confirmed. The spectra and RTTP decay of a solution of rabbit muscle aldolase were recorded for the first time. It was shown that denaturation of rabbit muscle aldolase under the action of urea (4–6 M) or heat leads to a dramatic decrease in the lifetime and total phosphorescence intensity. In following years it was demonstrated that RTTP spectroscopy is exquisitely sensitive to slow internal dynamics of the protein structure in the vicinity of tryptophanyls. It has been shown that changes in the RTTP lifetime reflect changes in the local rigidity of protein structure and the phosphorescence lifetime can be used as an intrinsic monitor of local flexibility of protein structure in regions of localization of tryptophan residues in the millisecond and second range. The dominating quenching mechanism of tryptophan phosphorescence in the oxygen-pured medium is deactivation of triplet excited states due to collisions of the indole ring of a tryptophan residue with the surrounding structural elements of the globule. The frequency of these collisions resulting in out-of-plane vibrations of the indole ring and relaxation of triplet states defines the efficiency of dynamic quenching of the tryptophan phosphorescence. The tryptophan residues located in protein regions with high dense packing of atoms characterized by rigid structure show long-lived phosphorescence at room temperature with τ ranging from several seconds to hundreds of milliseconds. As a rule such regions of protein are disposed in rigid hydrophobic core of globule. The RTTP lifetime of residues located in more mobile environments is reduced to the values from several tens of milliseconds to several milliseconds. The triplet excited states of tryptophan residues located in the nonstructured regions at the periphery of the globule with a low dense packing of atoms (in the highly mobile environment) are deactivated mainly via a nonradiative mechanism, resulting in the effective dynamic quenching of RTTP. In this case, the RTTP lifetime decreases to 1–0.5 ms. Since the quantum yield of the RTTP of such residues is very low (lower than 10–7 ), a phosphorescence signal from it could not always be detected [7–24]. Thus, monitoring of the slow internal dynamics of proteins in the millisecond and second range with the use of RTTP spectroscopy is possible due to the high sensitivity of the lifetime of the tryptophan phosphorescence to the mobility of the chromophore microenvironment and the correspondence of the values of lifetimes of RTTP to the characteristic times of low-frequency structural fluctuations of macromolecule. Along with the dynamic quenching of the tryptophan phosphorescence, static quenching takes place when the indole ring of a tryptophan residue is in close contact with the phosphorescence quenching amino acid residue(s)

Room Temperature Tryptophan Phosphorescence of Proteins

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(intramolecular quenching). Such amino acids can be divided into three classes by their ability to quench tryptophan phosphorescence: (1) strongly quenching (kq ≈5×108 M–1 s–1 ), such as cystine and cysteine; (2) intermediate quenching (kq ≈5×105 –2×107 M–1 s–1 ), such as tyrosine, histidine, and tryptophan; (3) weakly quenching or nonquenching (kq ≤105 M–1 s–1 ), all other amino acid residues [16]. A special role is played by disulfide groups in the intramolecular quenching. Their pronounced ability to quench RTTP is due to their capability to accept electrons from tryptophan in the triplet excited state [18]. Static quenching of the RTTP by amino acids occurs when the distance between the indole ring and the quenching group does not exceed several angstroms. Besides distance, phosphorescence quenching efficiency depends substantially on the orientation of the quenching group relative to the indole ring of a tryptophan residue. Intramolecular quenching may occur not only if the indole ring is in permanent contact with the quenching amino acid residue but also if it is transiently brought into contact with the quencher by fluctuations in the structure of the protein. Since the distance between the indole ring and the quencher as well as their mutual orientation can periodically change with fluctuations in the molecular structure, the internal dynamics of the protein can influence the efficiency of the intramolecular quenching of the RTTP [16]. Therefore, the presence of the potential RTTP quenching residues in the tryptophan residue microenvironment does not necessarily result in phosphorescence quenching. Because of the difficulty in taking into account all factors causing the triplet excited state deactivation, it is not always possible to define precisely the phosphorescence parameters of individual tryptophan residues in multitryptophan proteins. The phosphorescence method based on RTTP lifetime measurements was used for the studies of changes in the flexibility of the proteins in solution induced by denaturation and renaturation [5, 6, 21, 25–31], transition of the protein conformations into partially folded states [21, 27, 32–36], upon changes in the solvent viscosity [5, 21, 37, 38], hydrostatic pressure [39], pH and ionic strength [5, 15, 21, 40–43], binding of substrates, inhibitors, or allosteric effectors [5, 11, 21, 26, 44–49], removal of metal cations from the active center of an enzyme [21, 26, 50], limited proteolysis [21, 51], and association and aggregation [11, 21, 46, 47]. The RTTP of proteins in biological membranes and cells has been investigated to a much lesser extent as compared to the proteins in solution. In one of the few studies of the RTTP of cells [52], the phosphorescence properties of suspensions of E. coli strains containing varying amounts of alkaline phosphatase (the protein with a large quantum yield and RTTP lifetime) were investigated. It was shown that only E. coli alkaline phosphatase-rich strain exhibits long-lived RTTP. RTTP lifetime of E. coli mutant, which had no alkaline phosphatase, was less than 5 ms. The ability of suspensions of E. coli strains containing varying amounts of alkaline phosphatase to exhibit RTTP was re-examined in work [53]. On the basis of the data obtained it was concluded that all E. coli strains, independent of the amount of alkaline phosphatase, exhibit long-lived RTTP. Intensity and RTTP lifetime of E. coli alkaline phosphatase-rich strain were found to be much higher than that of the bacteria containing a usual amount of alkaline phosphatase or free of it. RTTP decay

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of suspensions of E. coli alkaline phosphatase-rich strain was single exponential with a lifetime of about 2 s, i.e., it was analogous to the decay time of the RTTP of purified alkaline phosphatase in solution. Long-lived RTTP decay kinetics of suspensions of the E. coli strains, containing a usual amount of alkaline phosphatase or free of it, has been satisfactorily approximated by a sum of two components. It is most probable that the authors of [52] failed to detect the long-lived RTTP of E. coli strains with low content (usual for the E. coli cells) of alkaline phosphatase because its phosphorescence intensity was below the detection limits of their apparatus. Unlike the proteins in fluid solution, all samples of the isolated biological membranes, uncolored microorganisms, plant and animal cells showed a reliably detectable long-lived RTTP. Their RTTP spectra have maxima at 410–415, 440–449, and 465–470 nm, respectively. The decay kinetics of RTTP of isolated membranes and cells was adequately approximated by the sum of two exponentials with varying lifetimes and contributions of the components. Since the majority of proteins in solution are incapable of exhibiting long-lived RTTP [9, 12, 14, 21], the question of possible source of phosphorescence signals of cells has been raised. In order to elucidate this question the comparative analysis of the capability of the membrane proteins in situ and water-soluble proteins of animal cells (rat hepatocytes) and microorganisms (bacteria E. coli) has been carried out [9, 53, 54]. It has been shown that long-lived RTTP of cells is determined by proteins in the composition of membrane structures. The RTTP lifetimes of the most soluble proteins of animal and bacterial cells were two or three orders smaller than those of the membrane proteins in situ. By using the example of the E. coli mutant strain containing an anomalously large amount of alkaline phosphatase, it was shown that the soluble proteins can contribute significantly to a phosphorescence signal of a cell only in relatively rare cases where these proteins have a high RTTP quantum yield and are present in the cell in large quantities [53]. The existence of the dramatic differences in the capability to exhibit long-lived RTTP between the membrane proteins in situ and water-soluble proteins of a cell provides a unique possibility of a selective investigation of the internal dynamics of living cell membrane proteins in situ. Changes in the RTTP decay kinetics of cell culture of the rat pheochromocytoma PC12 under the action of the epidermal growth factor and the neuronal growth factor in physiological concentrations (10–8 –10–12 M) [54, 55], the C. utilis microorganisms experiencing intercellular contact interactions [21], transformed into the anabiosis and reactivation states [21], under the action of biologically active substances of the humic nature [56] and ethanol [21], Chinese hamster fibroblasts subjected to an oxidation stress [57], human platelets after the treatment with trypsin at low concentrations [58–60], human peripheral blood lymphocytes, induced by binding of the HLA antibodies [61], and lectin concanavalin A at mitogenic concentrations (3 μg/ml) [54] were reported. These results are the evidence of changes in the slow internal dynamics of membrane proteins in situ. Changes in RTTP lifetimes of cells at the pathology made it possible to use the RTTP method for the diagnosis of autoimmune diseases and cancer [21, 62]. The RTTP of uncolored plant cells was detected. Their RTTP spectra were recorded and the decay kinetics was monitored. Differences in the RTTP kinetic


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parameters of plant cells for diverse species and varieties have been discovered [21, 63, 64]. It has been established that the decay kinetics of RTTP of plant cells changes under the action of brassinosteroid hormones at physiological concentrations (10–8 –10–12 M) [63]. Despite a certain progress toward an understanding of the RTTP of biological membranes and cells, many problems still remain to be solved. In particular, the reasons for the dramatic differences in the capability to exhibit RTTP between the majority of proteins in solution and the proteins in the composition of biological membranes are not clearly understood. The question of differences in the capability of integral and peripheral membrane proteins to exhibit RTTP has not been investigated experimentally. Little is known about how the protein–protein and protein–lipid interactions influence the parameters of the RTTP of membrane proteins. Investigations to be carried out in the indicated directions will allow to clarify the poorly studied problem on the internal dynamics of membrane proteins. In Sect. 2, results of phosphorescence analysis of isolated membranes of human erythrocytes are presented. These data contribute to the solution to these problems. Phosphorescence measurements of deoxygenated samples (residual oxygen concentration did not exceed 1 nM) were carried out with the conventional home-made instruments, described elsewhere [54, 65, 66]. A shutter arrangement permitted the delayed emission to be detected 1 ms after the excitation cutoff. RTTP decay kinetics was analyzed in terms of a sum of exponential components by a non-linear least square fitting using Origin 8.0 software (Microcal Software, USA). All decay data reported here are averages obtained from three or more independent measurements. Spectra were not corrected for instrumental response.

2 Room Temperature Tryptophan Phosphorescence of Proteins of Isolated Human Erythrocyte Membranes Isolated membranes of human erythrocytes represent a convenient investigation object that is frequently used in the membranology. This is explained by the fact that preparations of isolated membranes can be easily obtained and they are homogeneous, i.e., free of impurities of cellular organelle membranes. Moreover, the erythrocyte membranes are organized by the principles that are common for all the biological membranes. The membrane of a human erythrocyte is heterogeneous in composition and in structural organization of the proteins embedded in it. Integral proteins, also called transmembrane proteins, span the lipid bilayer and typically contain one or more hydrophobic transmembrane helices. Integral proteins of erythrocyte membrane are represented by band 3, glycophorins, different ATPases (K+ –Na+ -ATPase, Mg2+ -ATPase, Na+ -ATPase, and Ca2+ -ATPase). A major membrane-spanning integral protein is band 3. Peripheral proteins are layered on the surface of a plasma membrane. Spectrin (bands 1 and 2), actin (band 5), ankyrins (bands 2.1–2.3), proteins of bands 4.1, 4.2, and glyceraldehyde 3-phosphate dehydrogenase (band 6) belong to this class of proteins. Spectrin and

42


actin comprise about 30% of all of the proteins contained in the plasma membrane of an erythrocyte. These proteins organize spectrin–actin network, representing the base of the cytoskeleton, on the cytoplasmic side of the membrane. The ankyrins play an important role in the stabilization of the cytoskeleton structure. Spectrin is associated with integral membrane proteins by attachment to ankyrin. The protein of band 4.1 with adducin promotes spectrin–actin interaction and forms a ternary complex with the transmembrane protein glycophorin C and the membrane-associated guanylate kinase p55. The protein of band 4.2 interacts with cytoplasmic domain of the anion exchanger (band 3), ankyrin, spectrin, and protein of band 4.1 [67–69]. The erythrocyte membranes used in our experiments were isolated from a fresh human donor blood by the method developed by Dodge et al. [70] and were suspended in a 0.15 M sodium phosphate buffer of pH 7.4. The erythrocyte membranes were depleted of the peripheral proteins of the spectrin–actin network and the proteins of bands 2.1–2.3, 4.1, 4.2, and 6 by the methods described in [71]. The extent of elution of the peripheral proteins was controlled by the method of electrophoresis in a polyacrylamide gel in the presence of sodium dodecyl sulfate (SDS) [72]. Below are the results of investigations of the RTTP of proteins of an erythrocyte membrane in situ in the native state and after depleting them from peripheral proteins – actin–spectrin network (bands 1, 2, and 5) – and the proteins of bands 2.1– 2.3, 4.1, 4.2, and 6. We also investigated the capability of the peripheral proteins in solution, extracted from the membranes, to exhibit RTTP. The RTTP spectrum of a suspension of isolated human erythrocyte membranes has maxima at 415, 441 and 465 nm (Fig. 1, left, curve 1). The decay kinetics of the RTTP of these membranes presented in Fig. 1 (right, curve 1) is clearly multiexponential and has been adequately approximated by the sum of two exponential components with different lifetimes (τ 1 and τ 2 ), amplitudes of lifetime components (α 1 and α 2 ; α 1 + α 2 = 1), and contributions of lifetime components to the total emission (S1 and S2 ; S1 + S2 = 1). For the membranes in the native state (control), the RTTP lifetimes are τ 1 = 160 ms and τ 2 = 1630 ms, α 1 = 0.9, and S1 = 0.5

Fig. 1 Spectra (left) and decay kinetics (right) of RTTP of isolated human erythrocyte membrane suspensions: 1 – intact membranes, 2 – membranes depleted of spectrin and actin, 3 – membranes depleted of bands 2.1–2.3, 4.1, 4.2, 6 (besides spectrin and actin) and extracts of peripheral erythrocyte membrane proteins: 4 – spectrin and actin, 5 – bands 2.1–2.3, 4.1, 4.2, 6. 0.15 M sodium phosphate buffer; λex = 297 nm; 20◦ C


43

Table 1 RTTP parameters of suspensions of isolated human erythrocyte membranes with and without peripheral membrane proteins Isolated human erythrocyte membranes

τ 1 (ms)∗

τ 2 (ms)∗

α1 ∗

S1 ∗

Intact Depleted of spectrin and actin Depleted of bands 2.1–2.3, 4.1, 4.2, 6 (besides spectrin and actin)

160 118 103

1630 1323 330

0.91 0.93 0.89

0.52 0.55 0.72

The data are averages of three independent experiments. ∗ The standard deviation in these parameters is less than 6%.

(Table 1). The multiexponential character of the kinetic curves defining the decay of the RTTP of the membranes being considered can be explained by the fact that the tryptophanyls of the membrane proteins are situated in different environments. These differences, first of all, influence the slow internal dynamics of the proteins. It is likely that internal dynamics of membrane proteins in the vicinity of tryptophanyls phosphorescing with the value τ 1 is more intensive as compared to that at the locations of tryptophan residues phosphorescing with the value τ 2 . Therefore, the kinetic parameters of the RTTP of isolated erythrocyte membranes reflect the internal dynamics of the protein structure in places of tryptophanyl localization. To determine the role of protein–protein interactions in the slow internal dynamics of membrane proteins, we investigated the influence of the extraction of peripheral proteins from the erythrocyte membranes on for their RTTP. It has been established that the extraction of the peripheral proteins of the spectrin–actin network from the isolated membranes of human erythrocytes results in a pronounced quenching of the phosphorescence of integral and peripheral proteins remaining in the membrane, which manifests itself by a decrease in the intensity of the RTTP (Fig. 1, left, curve 2) and by an increase in the rate of its quenching (Fig. 1, right, curve 2). Table 1 shows phosphorescence decay parameters of a typical experiment. The data clearly display decrease in lifetime components of the RTTP of erythrocyte membranes, from which spectrin and actin were extracted, as compared to the native membranes; in this case, τ 1 decreased by approximately 1.4 times and τ 2 decreased by approximately 1.2 times. The RTTP quenching effect detected evidences that the slow internal dynamics of the membrane proteins is enhanced significantly on the depletion of membranes of spectrin–actin network proteins. Consequently, the proteins of spectrin–actin network significantly limit the slow internal dynamics of membrane proteins in situ. An additional extraction of the remaining peripheral proteins of bands 2.1–2.3, 4.1, 4.2, and 6 from the membranes subjected to the depletion of the spectrin–actin network proteins has caused further decrease in the intensity of the RTTP (Fig. 1, left, curve 3) and its lifetimes (Fig. 1, right, curve 3) to further decrease. The values of τ 1 and τ 2 of the RTTP of the depleted membranes were decreased as compared to those of the native membranes: τ 1 was decreased by approximately 1.6 times and τ 2 was decreased by approximately 4.9 times (Table 1). This indicates an increased flexibility of the chromophore microenvironment of integral proteins remaining in the membrane. These results provide evidence that

44


peripheral proteins forming the cytoskeleton not only stabilize the structure of a membrane but also substantially limit the slow internal dynamics of membrane proteins in situ. The extraction of peripheral proteins from the membrane substantially enhances the slow internal dynamics of proteins remaining in it, which is proved by the strong quenching of the RTTP detected in the above-described experiments. It was shown in [73] that the elution of spectrin from an erythrocyte membrane leads to a decrease in the activity of membrane enzymes (K+ –Na+ -ATPase, Ca2+ -ATPase) and the reassociation of spectrin with the membrane leads to their reactivation. The results of these experiments show that the cytoskeletal proteins of an erythrocyte membrane play an important role in the creation of optimum conditions for the functioning of membrane enzymes. It is conceivable that an important condition for the functioning of membrane proteins in situ is the provision of an optimum level of the amplitude and frequency of fluctuations of the protein structure with the participation of the cytoskeletal proteins. As the further experiments have shown, the proteins in the composition of native erythrocyte membranes are dramatically distinct from the solutions of the peripheral proteins extracted from the membranes in their capability to exhibit RTTP. Unlike the membrane proteins in situ, the millisecond RTTP of the aqueous extracts of peripheral proteins is strongly quenched and therefore they were not capable of exhibiting long-lived RTTP. As is seen from Fig. 1 (left, curves 4 and 5), the RTTP of aqueous extracts of peripheral proteins was not detectable. These point to the fact that the slow internal dynamics of peripheral membrane proteins in solution is much more intensive than that of the membrane proteins in situ. The data obtained as a result of the investigation of the RTTP of the spectrin–actin network proteins in solution are in good agreement with the data of [13, 35, 36], according to which the RTTP of actin in solution is strongly quenched. The low level of the RTTP of proteins in the composition of a membrane as compared to that of the proteins in solution can be explained by the existence of protein associates in the membrane, the isolation of the proteins included as a compound of the lipid bilayer from the aqueous environment, and the increased content of the rigid α-helices and β-structures in them. A suitable experimental approach to the elucidation of the factors controlling the internal dynamics of membrane proteins in situ is the study of the mechanisms of action of detergent on the isolated human erythrocyte membranes by the RTTP method. Detergents are chaotropic surfactants that can enter membranes, disturb the protein–protein and protein–lipid interactions, and denature the protein by disintegration of its globule. In the case where the content of detergent molecules in a membrane is relatively low, their action results in the appearance of defects in the lipid–matrix packing. As the amount of detergents in a membrane increases, the lipid-packing defects become larger and damage the structural integrity of the membrane, which leads to its subsequent fragmentation. A further increase in the detergent concentration leads to solubilization of the membrane with the formation of lipid–detergent and protein–detergent micelles. In this case, detergent molecules interact with the hydrophobic regions of the membrane proteins and displace the


45

lipid molecules. The solubilization process manifests itself as a sharp decrease in the light scattering of the membrane suspension. Detergents, depending on their chemical nature, can solubilize membranes with denaturation of proteins or without it. In the case where the concentration of detergents exceeds the critical micelle concentration (CMC), stable micelles consisting of a large number (from several tens to several hundreds) of molecules are formed in the aqueous medium. The CMC of ionic detergents comprises several millimoles and the CMC of the nonionic detergents is smaller by at least one order of magnitude. The action of detergents on membranes is determined by the concentration of their monomeric (not incorporated into the micelles) molecules. Therefore, the ionic detergents having a relatively high CMC, as a rule, more effectively solubilize membranes and more strongly affect the protein conformation than do the nonionic ones [74]. In the present work, we used ionic (anionic) and nonionic detergents having different capabilities to influence the conformation of proteins and disturb the protein–lipid and protein–protein interactions. The ionic detergents used were SDS (Reakhim, Russia), sodium deoxycholate (Serva, Germany), and N-lauroylsarcosine Na+ salt (sarcosyl) (Serva, Germany). The nonionic detergents were digitonin (Merck, USA) and Tween 20 (Serva, Germany). The detergent SDS has a high CMC (1.2–7.1 mM depending on certain conditions) and, therefore, solubilizes membranes to the level of individual lipid and protein macromolecules, disturbs all noncovalent interactions in a protein, and causes a deep denaturation of its globule. The ionic detergent sodium deoxycholate (CMC 1.5 mM) effectively solubilizes the membrane lipids and proteins, including the integral ones; however, it does not denature proteins. The third ionic detergent used in the present work – sarcosyl (CMC 0.54 mM) – damages the structure of proteins in the process of solubilization of a membrane to a minimum extent. Digitonin (CMC 0.087 mM) belongs to nonionic detergents. It is a mildly acting detergent that practically does not change the conformation of membrane proteins. The low CMC (0.06 mM) of the nonionic detergent Tween 20 also exerts its mild (nondenaturing) action on the structure of proteins in the process of solubilization of membranes [74, 75]. In this study, suspensions of membranes were incubated with each of the detergents at varying concentrations during 10 min at 22◦ C. We controlled the process of solubilization of membranes by detergents using the light-scattering method by measuring the optical density at a wavelength of 530 nm (D530 ). It was established that an increase in the detergent concentration in a suspension leads, as a rule, to a decrease in the RTTP lifetime of isolated human erythrocyte membranes. Figure 2 (top in left column, curves 1 and 2) shows the ratios of values of τ1 d to τ1 0 (r1 ) and τ2 d to τ2 0 (r2 ) of RTTP, where τ1 0 and τ2 0 are the phosphorescence lifetimes of isolated human erythrocyte membranes before treatment with detergent (control) and τ1 d , τ2 d are the phosphorescence lifetimes of isolated human erythrocyte membranes after treatment with detergent. It is seen that when the content of SDS increases to 5 mM, the value of r1 of the RTTP of the membranes decreases by 1.6 times as compared to the control value. The value of r2 of the RTTP decreases in the 0–3 mM narrower range of SDS concentrations. When the SDS concentration reaches 3 mM, the value of r2 of the RTTP of the membranes decreases by

46

V.M. Mazhul’ et al. 1,2

1,2

1,2

1,0

1,0

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2 0,8

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[Sodium dodecyl sulfate] / mM

1 2

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0,8

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0,2 0,0

0,6 0,4

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0,0

[Digitonin] / mM

1,2 1,0

0,4

0

2

4

6 [N-Lauroylsarcosine, Na+ salt] / mM

0,0

Fig. 2 The dependence of ratios of τ 1 d /τ 1 0 and τ 2 d /τ 2 0 on concentration of ionic (left column) and nonionic (right column) detergents in isolated human erythrocyte membrane suspensions. τ 1 0 and τ 2 0 are the phosphorescence lifetimes of membranes before treatment with detergent (control) and τ 1 d and τ 2 d are the phosphorescence lifetimes of membranes after treatment with detergent. Protein concentration is 5 mg/ml; 0.15 M sodium phosphate buffer; λex = 297 nm; 22◦ C

1.8 times and retains at this minimum level in the 3–6 mM range of detergent concentrations. Thus phosphorescence lifetimes of isolated human erythrocyte membranes substantially decreases under the action of SDS. The results of investigation of the solubilization activity of SDS by the method of turbidimetry are presented in Fig. 3 (left, curve 1). It is seen that when the SDS concentration reaches 2 mM, a suspension of erythrocyte membranes becomes transparent and the value of D530 approaches zero pointing out that the membranes are totally solubilized. Sodium deoxycholate influenced differently, depending on its concentration, on kinetics parameters of the RTTP of the erythrocyte membranes (Fig. 2, middle in the left column). When the sodium deoxycholate concentration in a suspension reached 2 mM, the values of r1 and r2 of the RTTP decreased approximately by 1.5 times and a further increase in the detergent concentration caused a rise in the lifetimes of

Room Temperature Tryptophan Phosphorescence of Proteins 0,6

0,3

Optical Density

0,6

Optical Density

47

3 2

2

0,3

1

1

0,0 0

2 4 Concentration/mM

6

0,0

0

2 4 Concentration/mM

6

Fig. 3 The dependence of the optical density at 530 nm on the concentration of ionic (left) and nonionic (right) detergents in isolated human erythrocyte membrane suspensions. Left: 1 – SDS, 2 – sodium deoxycholate, 3 – N-lauroylsarcosine Na+ salt (sarcosyl). Right: 1 – digitonin, 2 – Tween 20. Protein concentration is 0.3 mg/ml; 0.15 M sodium phosphate buffer; 22◦ C

the RTTP. As is seen from Fig. 3 (left, curve 2), the action of sodium deoxycholate of concentration 0–3 mM on a suspension of erythrocyte membranes leads to its pronounced transparency, decreasing the value of D530 to 0.13. Consequently, sodium deoxycholate is able to solubilize the membranes to the lesser extent in comparison to SDS. Sarcosyl (0–6 mM) induced a dose-dependent decrease of the value of r1 of the erythrocyte membranes by approximately 1.6 times (Fig. 2, bottom in the left column, curve 1). In this case, the value of r2 decreased by approximately 3 times with an increase in the sarcosyl concentration from 0 to 3 mM and remained unchanged when the concentration of the detergent increased further (Fig. 2, bottom in the left column, curve 2). This curve corresponds well to the curve of the dependence of the optical density of a suspension of erythrocyte membranes on the concentration of sarcosyl (Fig. 3, left, curve 3). The degree of transparency of a suspension of erythrocyte membranes treated with sarcosyl was relatively small (as small as D530 = 0.23) as compared to the degree of transparency of a suspension of erythrocyte membranes treated with SDS and sodium deoxycholate. The action of digitonin on a suspension of membranes also resulted in quenching of its RTTP (Fig. 2, top in the right column). This effect was most pronounced when the digitonin concentration varied within the range 0–1 mM. When the detergent concentration reached 1 mM, the value of r1 of the membranes decreased by 1.8 times (Fig. 2, top in the right column, curve 1), and the value of r2 decreased by 1.12 times (Fig. 2, top in the right column, curve 2). The results of the estimation of the degree of solubilization of the membranes being investigated with the use of digitonin by the decrease in the optical density of their suspension correspond well to the data of phosphorescence investigations. It is seen (Fig. 3, right, curve 1) that the change of transparency of a suspension of erythrocyte membranes (to D530 = 0.18) is most pronounced in the 0–1 mM range of digitonin concentrations. As a result of the action of Tween 20 in the concentration range of 0–6 mM on the membranes, the values of r1 and r2 were decreased by 1.24 and 1.2 times,

48


respectively (Fig. 2, bottom in the right column, curves 1 and 2). Phosphorescence lifetimes of membranes decreased most strongly when the Tween 20 content in a suspension increased from 0 to 0.5 mM. In this range of Tween 20 concentrations, the suspension of erythrocyte membranes underwent the major change in transparency (Fig. 3, right, curve 2). The experimentally detected changes in the values of r1 and r2 indicated the quenching of RTTP of erythrocyte membranes under the action of ionic and nonionic detergents (Fig. 2) caused by enhancing of slow internal dynamics of membrane proteins in regions of tryptophan residue localization. The initial portions of the dependences of r1 and r2 on the detergent concentration point out the fragmentation of the membranes and their further solubilization. The differences in the behavior of concentration dependences of r1 and r2 reflect an individual capability of detergents to solubilize the membrane. The good correspondence between the ranges of detergent concentration, in which the internal dynamics of the membrane proteins was enhanced (the values of r1 and r2 were decreased) and the membranes were solubilized (the optical density D530 of the suspension decreased), points to the contingency of processes of the enhancement of the internal dynamics of membrane proteins and the solubilization of membranes. The following effects can cause an enhancement of the internal dynamics of proteins in the composition of erythrocyte membranes in the process of their solubilization by detergents: 1. Denaturation of the structure of proteins. 2. An increase in the mobility of the molecular surroundings of the proteins due to the dissociation of the protein complexes in the membrane. 3. Transition of proteins from the viscose lipid bilayer of a membrane into the mobile molecular environment. It is likely that SDS, capable of deeply denaturating proteins and solubilizing membranes to the level of individual molecules of lipids and proteins, enhances the internal dynamics of the membrane proteins through the above-described three mechanisms. According to the light-scattering data, the capability of SDS to clarify a suspension of membranes to a maximum level is due to its better solubilization activity as compared to that of other detergents. Detergents such as sodium deoxycholate, sarcosyl, digitonin, and Tween 20 solubilize membranes without denaturation of the proteins (see above) through the disturbance of the protein–lipid interactions and the destruction of protein associates. Consequently, the restriction of the internal dynamics of the membrane proteins in situ, as compared to that of the soluble proteins, can be due to the isolation of the macromolecules from the mobile aqueous environment entering into the composition of the viscous lipid bilayer and the existence of protein associates in a membrane. These associates prevent the structure of the proteins from being damaged under the action of a solvent. The assumption that the viscose lipid bilayer of a membrane damps the slow fluctuations of the structure of the membrane proteins is supported by the results of experiments, myelin basic protein did not phosphoresce at room temperature in solution but it showed phosphorescence after being included


49

into the phosphatidylserine vesicles [10]. In addition to the above-listed factors that limit the internal dynamics of the integral membrane proteins in situ, this dynamics is evidently decreased by the rigid α-helices and β-strands present in large amounts in the proteins. The transmembrane regions of the integral proteins contain, as a rule, a large amount of ordered structures and are packed more closely than are the soluble proteins [76, 77]. Thus, the low internal dynamics of the membrane proteins in situ, as compared to that of the majority of soluble proteins, can be explained (1) by the presence of protein associates in a membrane, (2) by the isolation of the protein domains from the aqueous environment in the composition of the lipid bilayer, and (3) by the large amount of α-helices and β-strands in the macromolecules. An increase in the rigidity of the structure of the membrane proteins in situ, caused by the abovedescribed factors, substantially decreases the efficiency of the dynamic quenching of the tryptophan phosphorescence and is responsible for the pronounced capability of the membrane proteins to exhibit a millisecond RTTP. It has been noted above that the capability of tryptophanyls to exhibit phosphorescence is determined by the phosphorescence quenching amino acid residues located in close proximity to the tryptophanyls (intramolecular quenching). The efficiency of this quenching also depends on the slow internal dynamics of a protein in regions of tryptophan residue localization. Unfortunately, it is not possible to identify the concrete quenchers of phosphorescence of tryptophanyls of membrane proteins because of the absence of necessary X-ray structural analysis data of human erythrocyte membrane proteins. In this respect, soluble proteins with known 3D structures have advantages over the erythrocyte membrane proteins. In the next section, we present results of investigations of the RTTP of lectins, soluble plant proteins.

3 Room Temperature Tryptophan Phosphorescence of Plant Lectins in Solution Lectins are objects of intensive investigations in different fields of biology, medicine, and pharmacology. Representatives of the plant lectin class (concanavalin A, phytohemagglutinin) are widely used in the laboratory practice. These plant proteins are used as pharmaceutical raw materials and are a part of food. Some plant lectins show resistance against high temperatures and proteolytic enzymes and, therefore, can retain their biological activity in food, which is not always healthy [78]. Biological properties of lectins are determined in many respects by their structure. The 3D structure of many lectins is described; however, their slow internal dynamics has not been investigated. As it was noted above, information on the slow internal dynamics of proteins can be obtained from the analysis of its RTTP decay kinetics. In the present section, the results of investigations of plant lectins in solution by the RTTP method are presented. We investigated the following eight lectins: concanavalin A (Con A) from Canavalia ensiformis (jack bean),

50


phytohemagglutinin-L (PHA-L) from Phaseolus vulgaris (red kidney bean seeds), wheat germ agglutinin (WGA) from Triticum spp. (Wheat germ), peanut agglutinin (PNA) from Arachis hypogaea (peanut seed), Pisum sativum agglutinin (PSA) from P. sativum (pea seed), Sambucus nigra agglutinin (SNA-I) from S. nigra (elder bark), Laburnum anagyroides lectin (LAL) from L. anagyroides (bean tree seed), and Solanum tuberosum agglutinin (STA) from S. tuberosum (potato tuber). All lectins were purchased from Lektinotest (Ukraine) except for Con A, which was from Sigma-Aldrich (USA). Lectins used in our experiments were solvents in 0.15 M sodium phosphate buffer of pH 7.4 and phosphorescence measurements were carried out at 20◦ C. The data on the 3D lectin structures have been taken from the Protein Data Bank (PDB) [79] and included the following files: Con A (5CNA), PHA-L (1FAT), WGA (1WGT), PNA (2PEL), and PSA (2LTN). The program RasMol [80] was used to visualize and to analyze the structures. In the present work, we investigated the packing density, i.e., number of atoms (N) in the volume of a protein globule at distances of 2–10 Å from the surface of individual tryptophan residues, and the presence of intramolecular quenchers of phosphorescence in their vicinity. We also attempted to identify the tryptophanyls responsible for the room temperature phosphorescence of plant lectins.

3.1 Concanavalin A (Con A) Lectin Con A from the C. ensiformis (jack bean) is one of the most known plant lectins. Con A macromolecule consists of four identical subunits, each having a molecular mass of 25.6 kDa. The amino acid sequence of Con A subunit includes four tryptophanyls: Trp-40A, Trp-88A, Trp-109A, and Trp-182A. In Fig. 4a, the structure of the Con A tetramer with the highlighted tryptophanyls, based on the X-ray structural analysis data (file 5CNA in PDB) [81], is presented. According to these data, Trp-88A and Trp-109A are located in the β-strands of the Con A globule and are embedded into it; Trp-40A and Trp-182A are located in the nonstructured regions and are exposed to the solvent. Con A in solution shows a reliably detectable RTTP. The RTTP spectrum of Con A has maxima at 418, 444, and 461 nm (Fig. 5a). The decay kinetics of the RTTP of Con A is multiexponential and is adequately approximated by the sum of two exponentials with τ 1 = 17 ms and τ 2 = 520 ms (Fig. 6a). The values of τ 1 , τ 2 , α 1 , and S1 of Con A and other lectins are summarized in Table 2. The biexponential character of the decay kinetics of the RTTP can be explained by the differences in local flexibility of protein structure in regions of tryptophan residue localization and the presence of quenching amino acids in the nearest vicinity of tryptophanyls. Internal dynamics of the protein structure in the vicinity of tryptophanyls is determined by the packing density of the globule, i.e., number of atoms (N) in the volume of a protein globule. The analysis of the X-ray structural data using the RasMol program shows that the packing densities at various distances from the surface of individual


51

Fig. 4 Molecular structures of plant lectins: concanavalin A (a), phytohemagglutinin-L (b), wheat germ agglutinin (c), peanut agglutinin (d), and P. sativum agglutinin (e)

tryptophan residues Trp-40A, Trp-88A, Trp-109A, and Trp-182A are different (Fig.7a). The packing density of Trp-109A microenvironment is the highest. The packing densities of Trp-40A and Trp-88A surroundings are practically equal; however, they are lower than the packing densities of the Trp-109A microenvironment. The lowest packing density was detected in the vicinity of Trp-182A. Table 3 lists the amino acid residues found at different distances from the surface of individual tryptophan residues of Con A. The table was constructed on the

52


Fig. 5 Spectra of RTTP of lectins: concanavalin A (a), leucoagglutinin (b), peanut agglutinin (c), P. sativum agglutinin (d), S. nigra agglutinin (e), L. anagyroides lectin (f), S. tuberosum agglutinin (STA) (g)


53

Fig. 6 Decay kinetics of RTTP of lectins: concanavalin A (a), leucoagglutinin (b), peanut agglutinin (c), P. sativum agglutinin (d), S. nigra agglutinin (e), L. anagyroides lectin (f), S. tuberosum agglutinin (STA) (g)

54

V.M. Mazhul’ et al. Table 2 RTTP lifetimes and maxima of RTTP spectrum of plant lectins

Lectin

Quantity of components τ (ms)∗

τ1 (ms)∗

τ2 (ms)∗

α1 ∗

S1 ∗

Con A PHA-L WGA PNA PSA SNA-I LAL STA

2 2 – 2 2 2 2 1

17 190 – 73 39 18 111 –

520 456 – 150 252 340 446

0.84 0.52 – 0.38 0.92 0.95 0.94

0.25 0.3

– – – – – – – 1070

0.22 0.64 0.50 0.30

λ1 (nm)

λ2 (nm)

λ3 (nm)

418 420 – 418 415 416 416 417

444 445 – 444 441 443 442 444

461 465 – 466 463 466 467 465

The data are averages of three independent experiments. ∗ The standard deviation in these parameters is less than 6%.

basis of the X-ray structural analysis data (file 2CNA of PDB) obtained using the RasMol program [81]. As follows from the table, the quenchers of phosphorescence are absent in the region of location of Trp-109A. His-24A, Tyr-22A, and Tyr-64A are found in the microenvironment of Trp-40A. His-180A is located near Trp-182A. These amino acid residues are moderate quenchers (see Sect. 1) of RTTP. The features of the location of the tryptophanyls in the Con A globule allow us to conclude that, at room temperature, the Trp-109A residue incorporated into the rigid β-strain shows the long duration phosphorescence corresponding to the lifetime τ 2 . The packing density in the region of location of this tryptophanyl is the highest as compared to that of the other Con A tryptophanyls, and amino acid residues quenching phosphorescence are absent in their microenvironment. Trp-40A and Trp-88A are located in the globule regions with similar atomic packing density. However, the packing density of the surrounding of Trp-40A and Trp-88A is lower as compared to that of Trp-109A. Near Trp-40A, His-24A, Tyr-22A, and Tyr-64A are found, and His-180A is located in the vicinity of Trp-88A. Therefore, it is possible that both residues Trp-40A and Trp-88A can phosphoresce with a lifetime close to τ 1 at room temperature. It is likely that, at room temperature, Trp-182A does not phosphoresce in the millisecond range. Such a strong quenching of Trp-182A should be due to its location in the highly mobile nonstructured region of the globule, having a low packing density, and the existence of His-180A residues near Trp-182A.

3.2 Phytohemagglutinin-L (PHA-L) The structure of lectin PHA-L from P. vulgaris (red kidney bean seeds) represents a tetramer consisting of four identical subunits, each having a molecular mass of 27 kDa. The amino acid sequence of PHA-L subunit includes five tryptophanyls: Trp-58A, Trp-131A, Trp-153A, Trp-202A, and Trp-226A. Figure 4b presents the structure of the PHA-L tetramer with the indicated tryptophanyls, constructed using


55

Fig. 7 The number of atoms (N) in the volume of a protein globule at different distances from the surface of individual tryptophan residues of lectins: concanavalin A (a), leucoagglutinin (b), wheat germ agglutinin (c), peanut agglutinin (d), P. sativum agglutinin (e)

the RasMol program on the basis of the X-ray structural analysis data (file 1FAT in PDB) [82]. The RTTP spectrum of the PHA-L in solution has maxima at 420, 445, and 465 nm (Fig. 5b). The decay kinetics of the RTTP of PHA-L in solution is multiexponential and is adequately approximated by the sum of two exponentials with τ 1 = 190 ms and τ 2 = 456 ms (Fig. 6b). As seen from Fig. 7b, the number of atoms in the surrounding of Trp-58A, Trp-131A, Trp-153A, Trp-202A, and Trp-226A tryptophanyls is different. The structure of PHA-L in the regions of

Trp-40A

Asn-41A, Lys-39A Met-42A, Pro-23A Ala-38A, Asn-41A, His-24A, Ile-25A, Ley-9A, Thr-11A

Ala-73A, Asp-10A, Tyr-22A, Tyr-64A, Val-65A

Å

2 3 4

5

Asp-136B, Asp-139B, His-180A, Lys-138B, Ile-217A, Pro-86A, Pro-178A, Val-179A Asn-216A, Gln-137B, Ile-181, Ser-134B, Ser-215A

Glu-87, Val-89

Trp-88A Ser-108A, Ser-110A Phe-130A Gly-92A, Leu-93A, Leu-140A, Met-129A, Phe-128A, Phe-175A, Phe-195A, Ser-108A, Val-91A Ala-173A, Ala-193, Leu-154A, Leu-174A, Leu-193A, Phe-111A, Phe-133A, Phe-212A, Thr-194A

Trp-109A

Ala-189A, Gly-87A, Leu-81A, Leu-85A, Leu-115A, Pro-86A

Glu-183A, Ile-181A His-180A, Ley-85A Asp-82A, Ser-184A

Trp-182A

Table 3 Amino acid residues located at different distances from the concanavalin A tryptophanyls. Potential quenchers of phosphorescence are indicated in bold

56 V.M. Mazhul’ et al.


57

location of Trp-58A and Trp-131A is the least densely packed. According to the X-ray structural analysis data, these tryptophanyls are located in the nonstructured regions of the PHA-L globule and are exposed to the aqueous surroundings. The protein structure in these regions is usually subject to large thermal fluctuations. Therefore, it may be suggested that Trp-58A and Trp-131A are incapable of exhibiting RTTP with lifetime of more than several milliseconds. The packing density of PHA-L in the regions of location of Trp-153A, Trp-202A, and Trp-226A is much higher, which permits to conclude that these tryptophanyls are potentially capable of exhibiting RTTP. In the region where Trp-226A is located, the packing density of the PHA-L globule is the highest. This tryptophanyl is a part of the β-strand and the intramolecular quencher of phosphorescence Tyr-51A is located in its microenvironment. Trp-153A and Trp-202A are located in the nonstructured regions of the PHA-L globule. In the nearest vicinity of Trp-153A, two amino acid residues classified as moderate quenchers of phosphorescence (His-137A and Tyr-180A) are found (Table 4). In the microenvironment of Trp-202A, intramolecular quenchers of phosphorescence are absent. Since it is difficult to take into account all factors influencing the phosphorescence of the PHA-L tryptophanyls, the estimation of their actual capability to exhibit RTTP is a challenging task. Table 4 Amino acid residues located at different distances from the leucoagglutinin tryptophanyls. Potential quenchers of phosphorescence are highlighted in bold Å

Trp-153A

Trp-202A

Trp-226A

2 3 4

Asp-154A, Arg-152A Arg-136A Asp-124A, His-137A, Leu-178A, Phe-123A, Phe-155A, Phe-187A, Thr-125A, Thr-151A, Tyr-180A

Ser-227A, Ser-225A Phe-8A, Thr-70A Asn-7A, Glu-206A, Phe-6A, Phe-11A, Phe-207A, Ser-71A, Val-205A

5

Ile-138A, Phe-74A

Glu-201A, Val-203A Thr-14B Asn-12B, Asn-15B, Gln-56A, Glu-201A, Ile-55A, Pro-54A, Pro-94A, Pro-200A, Ser-204A, Val-95A, Val-203A Ile-57A, Thr-202A, Val-93A

Ley-16A, Ley-224A, Phe-72A, Phe-90A, Phe-228A, Tyr-51A, Val-223A

3.3 Wheat Germ Agglutinin (WGA) Lectin WGA from the Triticum spp. (wheat germ) consists of two identical subunits, each having a molecular mass of 18.6 kDa. The amino acid sequence of these subunits includes three tryptophanyls generally capable of exhibiting RTTP: Trp-41B, Trp-107A, and Trp-150A. However, we failed to detect the RTTP of WGA in the millisecond range. The actual reasons for the strong quenching of the RTTP can be the high mobility of the WGA tryptophanyl micro surrounding and the existence of

58


static quenchers in the nearest vicinity of them. The structure of the WGA dimer, constructed using the RasMol program based on the X-ray structural analysis data (file 1WGT in PDB) [83], is presented in Fig. 4c. According to the X-ray structural analysis data, Trp-41B, Trp-107A, and Trp150A are located in the nonstructured region at the periphery of the WGA globule. The packing density of the protein globule in the regions where the three WGA tryptophanyls are located is shown in Fig. 7c. It follows from the figure that the packing density of the surrounding of Trp-107A and Trp-150A of WGA is not high. Approximately the same packing density is characteristic of Trp-182A of Con A as well as Trp-58A and Trp-131A of PHA-L, all of which, as it was shown above, are incapable of exhibiting the millisecond phosphorescence at room temperature. Since quenchers of phosphorescence are absent in the regions of location of Trp-107A and Trp-150A of WGA (Table 5), the most probable reason for the pronounced quenching of their phosphorescence is the strong fluctuation of the protein structure with a millisecond characteristic time. Even though the packing density of atoms at the location of Trp-41B is higher than that at the location of Trp-107A and Trp-150A, the strong quenchers of phosphorescence Cys-31B, Cys-40B, and Cys-126A, located in the nearest vicinity of Trp-41B, apparently disable this WGA tryptophanyl to exhibit RTTP. Table 5 Amino acid residues located at different distances from the wheat germ agglutinin tryptophanyls. Potential quenchers of phosphorescence are highlighted in bold Å

Trp-41B

Trp-107A

Trp-150A

2 3 4

Thr-42B, Cys-40B

Gln-106A, Gly-108A

Ala-39A, Ala-125B, Cys-31B, Cys-126A, Glu-27A, Glu- 28B, Gly-28B, Gly-32B, Ser-43B, Ser-127B Asp-12BA

Gln-92A, Tyr-109A

Gly-151A Lys-149A Asp-29B, Gly-158A, Ser-148A, Ser-152A, Tyr-159A

Gln-122A

Asp-135A, Gln-165A

5

3.4 Peanut Agglutinin (PNA) Lectin PNA from A. hypogaea (peanut seed) represents a tetramer consisting of four identical subunits, each having a molecular mass of 25 kDa. The amino acid sequence of a PNA subunit includes three tryptophanyls: Trp-55A, Trp-153A, and Trp-223A. In Fig. 4d, the structure of the PNA tetramer with the highlighted tryptophanyls, constructed using the RasMol program based on X-ray structural analysis data (file 2PEL in PDB) [84], is presented. According to the X-ray structural analysis data, Trp-223A is incorporated into the β-strand and is embedded into the interior of the PNA globule. Trp-153A is also incorporated into the β-strand. The Trp-55A is located in the nonstructured region of the PNA globule, closer to its periphery.


59

It has been established that PNA in solution exhibits a reliably detectable RTTP. The RTTP spectrum of PNA in solution has maxima at 418, 444, and 466 nm (Fig. 5c). The decay kinetics of the RTTP of PNA in solution is multiexponential and is adequately approximated by the sum of two exponentials (Fig. 6c) and is characterized by the following lifetimes: τ 1 = 73 ms, τ 2 = 150 ms (Table 2). As is seen from Fig. 7d, the packing densities of the PNA globule at the localization of the PNA Trp-55A, Trp-153A, and Trp-223A are different. The packing density is relatively high for Trp-223A and Trp-153A and is low for Trp-55A. Table 6 presents the amino acid residues found at different distances from the surface of the PNA tryptophanyls. As follows from the table, static quenchers of phosphorescence are absent in the microenvironment of Trp-55A. His-137A is located in the nearest vicinity of Trp-153A, while Tyr-48A is located near Trp223A. Despite the presence of the static quencher near Trp-223A, this tryptophanyl residue may phosphoresce at room temperature with a lifetime close to τ2 because it is located in the most densely packed region of the PNA globule and incorporated into the β-strand. The packing density in the vicinity of Trp-153A is lower than that of Trp-223A (Fig. 7d). This gives reasons to expect that Trp-153A, also incorporated into the β-strand, is capable of exhibiting phosphorescence at room temperature with a lifetime close to τ 1 . It seems likely that Trp-55A located in the nonstructured region at the periphery of the PNA globule with a sparse packing of atoms does not phosphoresce at room temperature in the millisecond range. Table 6 Amino acid residues located at different distances from the peanut agglutinin tryptophanyls. Potential quenchers of phosphorescence are highlighted in bold type Å

Trp-55A

Trp-153A

Trp-223A

2 3 4

Ile-54A, Ser-56A Leu-198A Asn-61A, Arg-53A, Glu-200A, Leu-198A, Lys-195A, Pro-199A, Ser-57A

Asn-154A, Pro-152A

Ser-222A, Ser-224A Phe-8A, Thr-67A Asn-7A, Phe-6A, Phe-69A, Phe-204A, Phe-206A, Tyr-48A

5

Ala-63A, Leu-194A, Leu-229A, Val-62A

Asp-136A, Asn-180A, His-137A, Ser-155A, Thr-185A, Thr-186A, Phe-122A, Val-138A, Val-151A, Val-178A Asp-123A, Met-173A, Thr-124A, Thr-135A, Val-161A

Arg-221A, Gly-205A, Ile-220A, Leu-47A, Phe-11A, Phe-87A, Phe-225A, Ser-68A

3.5 Pisum sativum Agglutinin (PSA) Lectin PSA lectin from P. sativum (pea seed) consists of two identical subunits, each having a molecular mass of 25.2 kDa. The amino acid sequence of a PSA subunit

60


includes five tryptophanyls: Trp-19B, Trp-40B, Trp-53A, Trp-128A, and Trp-152A. In Fig. 4e, the structure of the PSA dimer with the highlighted tryptophanyls, constructed using the RasMol program based on the X-ray structural analysis data (file 2LTN in PDB) [85], is presented. Table 7 presents the amino acid residues found in the vicinity of the PSA tryptophanyls. PSA in solution is capable of exhibiting millisecond RTTP. The RTTP spectrum of PSA has maxima at 415, 441 and 463 nm (Fig. 5d). The decay kinetics of RTTP of PSA is multiexponential. It is adequately approximated by the sum of two exponentials (Fig. 6d) with τ 1 = 39 ms and τ 2 = 252 ms (Table 2). Tryptophanyls of PSA are different in packing density in the regions of its location. Unfortunately, identification of PSA tryptophanyls capable of exhibiting phosphorescence at room temperature is very difficult because of the presence of amino acid residues quenching phosphorescence in their microenvironment. Out of the five tryptophanyls, only Trp-53A is included in the β-strand. However, distal position of Trp-53A and relatively low packing density in the region of its location (Fig. 7e) did not allow establishing its capability to exhibit phosphorescence at room temperature.

Table 7 Amino acid residues located at different distances from the P. sativum agglutinin tryptophanyls. Potential quenchers of phosphorescence are highlighted in bold Å

Trp-19B

Trp-40B

2

Gly-18B

3

Val-20B,

Ser-39B, Ser-41B Ile-8A, Thr-65A

4

Asn-17C, His-51A, Gln-15C, Gln-16C, Ile-50A, Pro-17B, Pro-49A,

5

Ala-88A, Gly-23B, Asp-14C, Leu-18A, Asn-17C, Glu Phe-11A, 2A, Pro-89A, Phe-42B, Ser-12C, Phe-67A, Val-90A Phe-85A, Tyr-46A, Val-37B

Trp-53A

Asp-54A, Ile-52A Glu-18B, Val-16B Ile-22B, Leu-7A, Arg-55A, Phe-6A, Asn-59A, Phe-24B, His-51A, Ser-66A Lys-13B, Pro-17B, Val-60A

Ala-61A, Leu-12B, Leu-46B

Trp-128A

Trp-152A

Ala-127A, Asp-129A

Lys-153A, Ser-151A Arg-135A

Asn-125A, Glu-99A, Ile-144A, Lys-145A, Tyr-100A

Asp-121A, His-136A, Ile-71A, Ile-137A, Leu-154A, Leu-177A, Lys-150A, Phe-120A, Thr-122A, Tyr-4B, Tyr-174A Phe-69A

Ala-107A


61

3.6 Sambucus nigra Agglutinin (SNA-I) Lectin SNA-I lectin from S. nigra (elder bark) is a tetramer. The molecular mass of the SNA-I tetramer is equal to 140 kDa. The subunits of this lectin are connected by disulfide bonds. Figure 5e shows the RTTP spectrum of SNA-I lectin in aqueous solution. As in the case of the above-described lectins, the RTTP spectrum of SNA-I has the maxima at 416, 443, and 466 nm. The decay kinetics of the RTTP of SNAI in solution is multiexponential and adequately approximated by the sum of two exponentials (Fig. 6e) with τ 1 = 18ms and τ 2 = 340 ms (Table 2). Unfortunately, the 3D structure of SNA-I is not available to date and therefore, we could not analyze the amino acid sequence, the packing density near the tryptophan residues, the presence of quenchers, and the tryptophanyls responsible for the RTTP of this lectin.

3.7 Laburnum anagyroides Lectin (LAL) It was established that lectin LAL from L. anagyroides (bean tree seed) in solution is capable of exhibiting RTTP. As is seen, its RTTP spectrum has maxima at 416, 442, and 467 nm (Fig. 5f). The decay kinetics of the RTTP of LAL lectin is multiexponential (Fig. 6f) and adequately approximated by the sum of two exponentials with τ 1 = 111 ms and τ 2 = 446 ms (Table 2), which points to the fact that the properties of the surroundings of the tryptophanyls capable of exhibiting RTTP are heterogeneous. These differences, first of all, can be due to differences in packing density between the regions of location of the tryptophanyls in the LAL globule and the presence of the amino acid residues, representing potential quenchers of phosphorescence, in their nearest environment. However, as for SNA-I, at present, the 3D structure of LAL has not been yet determined, hence the analysis of the features of the localization of the tryptophanyls in the LAL globule and the identification of the tryptophan residues phosphorescing at room temperature with different lifetimes are impossible.

3.8 Solanum tuberosum Agglutinin (STA) Figure 5g represents the RTTP spectrum of lectin STA extracted from S. tuberosum (potato tuber) in solution. Figure 6g presents the monoexponential decay kinetics of the RTTP of STA in solution with τ = 1070 ms. The large (for the protein in solution) lifetime of the RTTP of STA (more than 1 s) points to an extremely high packing density of atoms at the location of the tryptophanyl responsible for the phosphorescence and, consequently, to a high rigidity of the structure of the hydrophobic core of the STA globule in the region of location of this tryptophanyl. Unfortunately, the absence of X-ray structural analysis data on the STA structure gives no way to discuss the features of the location of the tryptophanyls in the STA globule and their capability to exhibit RTTP in detail.

62


Thus, as a result of the above-described works, the RTTP of seven of the eight plant lectins have been detected for the first time. They are as follows: Con A from C. ensiformis (jack bean), PHA-L from P. vulgaris (red kidney bean seeds), PNA from A. hypogaea (peanut seed), PSA from P. sativum (pea seed), SNA-I from S. nigra (elder bark), LAL from L. anagyroides (bean tree seed), and STA from S. tuberosum (potato tuber). It has been established that RTTP decay kinetics of lectins, depending on the characteristic features of their tryptophan residues localization, can be adequately approximated by the sum of two exponentials (Con A, PHA-L, PNA, PSA, SNA-I, LAL) or one exponential (STA). On the basis of 3D protein structures taken from the PDB, the features of the location of the tryptophan residues of plant lectins have been analyzed and their capability for the phosphorescence at room temperature has been discussed taking into account the packing density of atoms in the lectin globule and the existence amino-acid residues possessing the properties of static quenchers of phosphorescence. The relatively large values of the lifetimes of the RTTP of the plant lectins in solution point to the notion that the slow internal dynamics of these lectins is smaller than that of the majority of proteins in solution which are incapable of exhibiting long-lived RTTP. It can be explained by the high content of β-strands in the plant lectins. The high conformational rigidity of these lectins is probably a reason for their resistance to the action of temperature and to other physico-chemical factors. In conclusion, it is necessary to note that the “RTTP” abbreviation is not always used in the literature. In works devoted to the tryptophan phosphorescence at room temperature, the abbreviation “RTP” (room temperature phosphorescence) is frequently used. In our opinion it is preferable to use the abbreviation “RTTP” for the following reasons: 1. Except for tryptophanyls, tyrosine residues of proteins may also phosphoresce at room temperature [86]. 2. In biological membranes and cells, apart from proteins, the lipid peroxidation (LPO) products are capable of exhibiting room temperature phosphorescence. Excitation spectra of room temperature phosphorescence of LPO products are localized in the spectral range of 240–420 nm. LPO products phosphoresce at room temperature in the range of 420–600 nm [6, 87–91]. Taking into account that in biological objects not only tryptophanyls but also other chromophores are capable of exhibiting room temperature phosphorescence, the use of the abbreviation “RTTP” better defines the nature of the phosphorescence. Acknowledgment This work is supported by International Science and Technology Center, Grant B-1332.


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Rational Design of FRET-Based Sensor Proteins M. Merkx

Abstract Real-time imaging of molecular events inside living cells is important for understanding the basis of physiological processes and diseases. Genetically encoded sensors that use fluorescence resonance energy transfer (FRET) between two fluorescent proteins are attractive in this respect because they do not require cell-invasive procedures, can be targeted to different locations in the cell and are easily adapted through mutagenesis and directed evolution approaches. Most FRET sensors developed so far show a relatively small difference in emission ratio upon activation, which severely limits their application in high throughput cell-based screening applications. In our work, we try to develop strategies that allow design of FRET-based sensors with intrinsically large ratiometric changes. This rational design approach requires a better understanding and quantitative description of the conformational changes in these fusion proteins. In this chapter, I first discuss some of the key factors and strategies that determine the ratiometric response of FRET sensors, followed by an overview of our recent work in this area. Important concepts that will be discussed are (1) the conformational behavior of flexible peptide linkers to quantitatively describe the dependence of energy transfer on linker length and (2) the control of intramolecular domain interactions using the concept of effective molecular concentration.

1 Introduction Real-time imaging of molecular events inside living cells is important for understanding the molecular basis of physiological processes and diseases [1]. Fluorescence is ideally suited for this purpose because it combines high sensitivity with subcellular resolution. A wide variety of synthetic fluorescent probes have been M. Merkx (B) Laboratory of Chemical Biology, Department of Biomedical Engineering, Eindhoven University of Technology e-mail: [email protected] C.D. Geddes (ed.), Reviews in Fluorescence 2008, Reviews in Fluorescence 2008, DOI 10.1007/978-1-4419-1260-2_3, C Springer Science+Business Media, LLC 2010

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developed that allow specific detection of pH, redox status, metal ions, enzymatic activity, membrane potential, etc. Although some of them have become important molecular tools to probe intracellular events such as calcium signaling, these probes need to be introduced across cellular membranes and little control is possible over their cellular localization and concentration. Protein-based sensors based on native fluorescent proteins such as GFP and its color derivatives have become attractive alternatives for these synthetic probes [2–5]. Because these sensors are genetically encoded, their concentration and subcellular localization can be easily controlled. In addition, native protein domains can be used as receptor building blocks to ensure physiologically relevant binding affinities and specificities. Finally, molecular biology approaches such as directed evolution and site-directed mutagenesis can be used to optimize various sensor properties. Several fluorescent sensor proteins consisting of a single GFP domain have been developed in which the chromophore environment was reengineered to become sensitive to ions such as H+ , Cl− , Ca2+ [2, 4, 5]. This approach is useful for small molecules that have relatively high physiological concentrations and therefore do not require specific and high-affinity binding sites. A second, more general and therefore more interesting approach takes advantage of fluorescence resonance energy transfer (FRET) between a donor and an acceptor fluorescent domain. FRET is a photophysical effect whereby the energy that is absorbed by one fluorophore (donor) is transferred non-radiatively to a second fluorophore (acceptor) [6, 7]. The distance and orientation dependence of FRET make it extremely useful to detect conformational changes on the scale of individual proteins. Any biochemical event (e.g., ligand binding) that leads to a significant conformational change can be detected, irrespective of the protein and the nature of the protein–ligand interaction. Another important characteristic of FRET for sensor design is the possibility of ratiometric detection. An increase in FRET leads to a decrease in donor emission and an increase of acceptor emission. The ratio of donor and acceptor emission therefore provides a measure of the binding state that is independent of the absolute concentration of the sensor, which is important to obtain reliable, quantitative results in biological samples. Alternatively, FRET can be monitored by determining the lifetime of either the donor or the acceptor emission, both of which are sensitive to energy transfer. The most common approach in the development of FRET-based sensors has been one of trial and error in which many constructs (which differ in linker lengths, fluorescent and receptor domains, etc.) are screened for optimal intracellular performance. While this strategy may be suitable to obtain a FRET sensor for a single, specific target with sufficient dynamic range to allow fluorescence imaging, little insight is gained into the factors that determine sensor performance. With some notable exceptions (see below) most FRET sensors show a relatively small difference in emission ratio upon activation, which severely limits their application in high throughput cell-based screening applications. In our work, we try to develop strategies that allow design of FRET-based sensors with intrinsically large ratiometric changes. This rational design approach requires a better understanding and quantitative description of the conformational changes in these fusion proteins. In this

Rational Design of FRET-Based Sensor Proteins

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review, first some of the key factors and strategies will be discussed that determine the ratiometric response of FRET sensors, followed by an overview of our recent work in this area. Important concepts that will be discussed are (1) the conformational behavior of flexible peptide linkers to quantitatively describe the dependence of energy transfer on linker length and (2) the control of intramolecular domain interactions using the concept of effective molecular concentration.

2 Factors That Affect the Ratiometric Change in FRET-Based Sensor Proteins One of the strengths of FRET-based sensor proteins is their modular design in which a conformational change in a receptor domain is translated into a change in energy transfer between donor and acceptor domains. Alternatively, two receptor domains can be employed that interact in one state but not in the other. Examples of the latter approach are more common and include the calcium sensors developed by Tsien and others [8] and a variety of sensors for protein phosphorylation [2]. Despite the strong distance dependence of the energy transfer efficiency (Eq. (1)), it has proven surprisingly difficult to translate conformational changes into ratiometric changes that are larger than 1.5-fold. To understand why this is, we need to take a look at the factors that determine energy transfer efficiency in these fusion proteins. The relation between the energy transfer efficiency E and the distance between the donor and fluorescent domains is described by the Förster equation: E=

R60 R60 + r6

(1)

whereby the Förster distance, R0 , is given by 1/6 R0 = 0.21 κ 2 QD n−4 J(λ)

(2)

Thus, the energy transfer efficiency not only depends on the distance between the chromophores but is also dependent on the quantum yield of the donor (QD ), the spectral overlap between donor emission and acceptor absorption (J(λ)), the refractive index of the medium (n), and an orientational factor κ 2 , which is related to the relative orientation of the donor emission and acceptor absorption dipole moments. Of the parameters that determine R0 , κ 2 is the most likely to change upon sensor activation. Equation (3) describes how κ 2 depends on the angles (θ D , θ A , θ T , and φ) between the dipole moments of the donor and acceptor, as depicted in Fig. 1[6]: κ 2 = ( cos θT − 3 cos θD cos θA )2 = ( sin θD sin θA cos φ − 2 cos θD cos θA )2

(3)

If the orientation of donor and acceptor domains undergoes complete randomization while the donor is in the excited state, κ 2 averages out to a value of 2/3. If

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Fig. 1 Definition of angles between the donor emission dipole moment (D) and the acceptor absorption dipole moment (A), separated by distance r. Values for κ 2 are given for perpendicular, parallel, and collinear orientations of D and A

the orientation is fixed, κ 2 can vary between 0 (perpendicular), 1 (parallel), and 4 (collinear). Having shown how the energy transfer efficiency depends on the distance and orientation of the donor and acceptor domains, we can identify several reasons for the low dynamic range of many genetically encoded FRET-based sensors: (1) Changes in the distance between donor and acceptor are detected with the greatest sensitivity around the Förster distance, which is 48 Å for the most common pair of cyan and yellow fluorescent domains [9, 10]. However, because the fluorophores are buried inside the fluorescent protein’s β-barrel and the intervening receptor domain is often also large, the energy transfer is relatively inefficient in most sensors, resulting in small changes in FRET. (2) Flexible linkers of at least a few amino acids are required between two domains to allow proper folding of the individual domains. As a result, effective translation of the conformational change in the receptor domain into a significant conformational change for the entire sensor protein is hard to achieve. (3) Because κ 2 can vary between 0 and 4, relatively small differences in orientation can in principle have a large effect on energy transfer efficiency. However, because of the limited possibilities to predict conformational preferences in fusion proteins, taking advantage of the orientational factor using rational design approaches is very difficult. In addition, transition dipole moments have been determined experimentally only for green fluorescent proteins [11, 12]. While the transition dipole moment of the yellow fluorescent domains are likely to be similar to that of EGFP, the transition dipole moments of frequently used donor domains such as ECFP and Cerulean are not known. Despite the lack of true rational design approaches, two more empirical approaches are worth mentioning. A strategy that has been successfully applied in the optimization of several FRET sensors is the use of circularly permuted fluorescent protein domains [13–15]. In a circular permuted protein, the original C- and N-termini are connected by a short linker and new termini are generated at a solvent accessible loop [16]. Employment of a circular permuted form of Venus (a YFP variant) was shown to increase the ratiometric response of the cameleon calcium


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sensors by 560%, probably due to a change in the orientational factor. However, no clear correlation between the type of circular permutation and the effect on the ratiometric change was observed, making this approach still (to some extent) a process of trial and error. A second strategy that was reported by Daugherty and coworkers is the use of directed evolution combined with FACS screening to select FRET sensors with an increased ratiometric change [17]. Variants of ECFP and EYFP, CyPet and YPet, were identified that, when linked via a flexible caspase recognition sequence, yielded a FRET sensor with a 20-fold ratiometric change upon cleavage by caspase-3. Although YPet and CyPet have been advertised as the donor and fluorescent domains of choice for FRET sensors [18, 19], we and others have recently shown that their improved FRET properties are caused by an increased tendency to form an intramolecular complex when linked via a flexible peptide linker, limiting their potential as general applicable fluorescent domains for FRET sensors [20, 21].

3 Quantitative Understanding of Energy Transfer by Modeling the Conformational Behavior of Flexible Linkers Despite their importance in determining the functional properties of FRET sensors, little attention has been given to understand the effects of linker length and constitution on the energy transfer properties. Many studies reported a decrease in energy transfer upon increasing the linker length between donor and acceptor domains, but no attempt was made to quantitatively understand the efficiency of energy transfer as a function of linker length [22, 23]. Repeats of flexible (Gly) and hydrophilic (Ser) residues are popular linker peptides, because they are assumed to form a random coil structure in solution and to not interact with the protein domains [24]. We constructed a series of ECFP-linker-EYFP proteins in which the linker length was systematically varied between 1 and 9 GGSGGS repeats (CLY1–9) and carefully determined the amount of energy transfer for each linker length (Fig. 2) [10]. The energy transfer efficiency decreased from 71 to 43% when going from CLY1 (23 residue linker) to CLY9 (71 residue linker). Direct application of the Förster equation (assuming R0 = 48 Å), would yield interchromophore distances ranging from 41 Å for CLY1 to 50 Å for CLY9. In other words, an increase in linker length of 48 amino acids results in an apparent increase in distance of only 9 Å! This analysis assumes that the energy transfer is due to a single fixed distance, however, which is clearly not the case when the fluorescent domains are linked via a flexible linker and a distribution of interchromophore distances is present. We therefore analyzed the energy transfer by considering the distribution of conformations available to the two fluorescent domains, which in turn is based on the conformational properties of the linker peptide. Two models frequently used to describe the conformational behavior of random coil structures are the Gaussian chain model developed by Flory [25] and the WormLike Chain (WLC) model [26, 27]. For long flexible chains (as is the case here) both models are actually equivalent and only differ in the definition of the parameters

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Fig. 2 (A) ECFP-linker-EYFP proteins containing a linker of 1–9 copies of the GGSGGS repeat (CLY1–9) that were used to understand the influence of linker length on energy transfer efficiency. Shown are the residues that are assumed to be part of the linker, as well as the spherical distribution of orientations that are accessible to the vectors that connect the ends of the peptide linkers and the chromophores (vc and vy ). The end-to-end distance of the linker (re ) is translated into a chromophore-to-chromophore distance (rc ) by averaging over all possible orientations of vc and vy . (B) Emission spectra of CLY1-9, ECFP, and EYFP were obtained using 420 nm excitation. Measurements were done using 0.2 μM protein in 50 mM Tris-HCl, 100 mM NaCl, 20 μM EDTA, 10% (v/v) glycerol, pH 8.0. Figures reprinted with permission from reference [10]. Copyright 2006 American Chemical Society

used to describe chain stiffness. The Gaussian chain model uses the characteristic ratio (C∞ ), whereas the WLC model uses the persistence length (lp ). Here, we will analyze our data using the WLC model, because it has the advantage that it can be applied more easily to short and stiff polymers. In the WLC model the probability (P) of finding a certain end-to-end distance (re ) for a chain with a total contour length lc is described by Eq. (4). P(re ) = 4πre2 (3/4πlp lc )3/2 exp ( − 3re2 /4lp lc )(1 − 5lp /4lc − 2re2 /lc2 + 33re4 /80lp lc3 + 79lp2 /160lc2 + 329re2 lp /120lc3

(4)

− 6799re4 /1600lc4 + 3441re6 /2800lp lc5 − 1089re8 /12800lp2 lc6 ) The total contour length is equal to b0 n, with n being the number of peptide bonds and b0 = 3.8 Å (the average distance between adjacent Cα atoms). Using a persistence length lp of 4.5 Å, the probability of finding a certain end-to-end distance can be calculated for each of the peptide linkers present in CLY1-9 (Fig. 3). To calculate the amount of energy transfer at a certain re , it should be realized that ECFP and EYFP can assume many different orientations, which differ in the interchromophore distance rc and can therefore all contribute differently to the energy transfer efficiency. Consequently, the average energy transfer efficiency, <E>, was determined by averaging the value of E calculated for each combination of ECFP and EYFP orientations, assuming distances of 20 and 24 Å between the ends of the peptide linker and the ECFP and EYFP chromophores, respectively (Fig. 3b). In doing so, we can account for the fact that orientations with a short interchromophore distance contribute more to the energy transfer efficiency than orientations in which ECFP and EYFP are relatively far apart. Finally, the overall energy transfer


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Fig. 3 Modeling the energy transfer efficiency for CLYx using the WLC model to describe the peptide linker. (A) Distribution functions showing the probability P(re ) for each end-to-end distance of the peptide linker for CLY1–9 calculated using the WLC model assuming a persistence length of 4.5 Å. (B) Plot showing the average energy transfer ( E, solid line) and the average interchromophore distance ( rc , dashed line) as a function of re . (C) The contribution of each re to the overall energy transfer

Eensemble , calculated by multiplying the probability of each re with the corresponding E. Figure reprinted with permission from reference [10]. Copyright 2006 American Chemical Society

efficiency was calculated by multiplying the function describing <E> as a function of re by the probability of finding a certain re . Integration of this curve yields the predicted overall energy transfer efficiency (<E>ensemble ) for each of the CLY proteins. Figure 4 shows that there is excellent agreement between the predicted value for <E>ensemble using the random coil model and the experimentally determined energy transfer efficiency over the entire range of linker lengths. To show the strong dependence of the predicted energy transfer on the persistence length, Fig. 4 also shows the predicted curves for lp values of 4.0 and 3.0 Å. The value of 4.5 Å corresponds to a C∞ value of 2.3 in the Gaussian chain model, which is close to the C∞ value of 2.16 previously reported for poly(Gly) [28].

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Fig. 4 Comparison of the experimentally determined energy transfer efficiency (triangles) with the energy transfer efficiency predicted by the WLC model using persistence lengths of 3.0, 4.0, and 4.5 Å. Figure reprinted with permission from reference [10]. Copyright 2006 American Chemical Society

The fact that the energy transfer efficiency between ECFP and EYFP can be quantitatively understood by describing the flexible linker as a random coil has several important implications for FRET sensor design. First, this study shows that random coil peptide linkers form relatively compact assemblies, giving rise to relatively high energy transfer efficiencies. For example, while the long peptide linker used in CLY9 (71 amino acids) can span a distance of 252 Å when fully extended, the most abundant end-to-end distance is only 41 Å. A second issue that is sometimes misunderstood is that the energy transfer efficiency between fluorescent domains separated by flexible linkers reflects a broad distribution of distances and not a single distance. Because of the nonlinear dependency of E on distance, it is not correct to calculate an average distance based on the energy transfer efficiency. The apparent average distance will be underestimated for long linkers and overestimated for short linkers.

4 Quantitative Understanding of the Effect of Flexible Peptide Linkers on Effective Concentration In the previous section, we showed that the conformational distribution of two protein domains connected via Ser/Gly linkers can be accurately described by modeling the linkers as flexible chains with a persistence length of 4.5 Å [29]. The same model can also be used to calculate the effective local concentration of one protein domain with respect to another in a system in which the two protein domains are linked [27, 30, 31]. A clear demonstration of the predictive power of this model, and the importance of considering conformational distributions in the design of FRET-based sensor proteins, is our work on the Zn(II) sensor CALWY [29, 32]. This sensor is based on a serendipitous discovery that Zn(II) mediates an interaction between ATOX1 and WD4, two metal binding domains involved in Cu(I) homeostasis. A sensor system consisting of ECFP-Atox1 and WD4-EYFP was shown to bind Zn(II) at subnanomolar levels using equimolar amounts of both proteins (2 μM) [32]. Next, a series of ECFP-Atox1-linker-WD4-EYFP fusion proteins were


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Fig. 5 Variation of the linker length in CA-Ln-WY allows rational tuning of Zn(II) affinity in the picomolar to femtomolar range. (A) Structural model of CA-Ln-WY in the Zn(II) bound state. (B) Zn(II) titration for CFP-Atox1-linker-WD4-YFP proteins with 2, 5, and 9 GGSGGS repeats. Titrations were performed in 50 mM Tris, 100 mM NaCl, 1 mM DTT, 10% glycerol at pH 7.5 using various Zn(II) buffers. Also shown is a titration for a 1:1 mixture of CFP-Atox1 and WD4-YFP (2 μM each). Figures reprinted with permission from reference [29]. Copyright 2007 American Chemical Society

constructed in which the linker length was systematically varied between 2 and 9 GGSGGS repeats (CA-Ln-WY, with n = the number of GGSGGS repeats; Fig. 5a) [29]. This series of sensor proteins allowed us to systematically study the effect of the linker length on both the relative ratiometric change and the affinity for Zn(II), which was expected to increase due to the chelate effect. The Zn(II) affinity showed a consistent trend with Kd values ranging from 170 fM for the protein with the longest linker (CA-L9-WY) to 1.4 pM for the shortest linker (CA-L2-WY). Remarkably, the Zn(II)-induced intramolecular complex formation between Atox1 and WD4 resulted in a decrease in energy transfer for all sensor constructs (Fig. 5b). The emission ratio in the unbound state was strongly dependent on the linker length, whereas similar emission ratios were observed for the Zn(II)-bound state. This behavior can be readily understood by considering the conformational behavior of the linker. The distance between the C-terminus of Atox1 and the N-terminus of WD4 in the Zn(II) complex has been estimated to be 50 Å, based on the X-ray structure of an analogous Cd(II)-bridged Atox1–Atox1 homodimer complex [33]. Figure 6 shows the distribution of end-to-end distances of the

Fig. 6 Distribution functions showing the probability P(re ) of a certain end-to-end distance re for the peptide linkers used in CA-L2-WY, CA-L5-WY, and CA-L9-WY. Figure reprinted with permission from reference [29]. Copyright 2007 American Chemical Society

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peptide linker as predicted by the WLC model for linkers containing 2, 5, and 9 GGSGGS repeats. Even for the longest linker, the average end-to-end distance is smaller than 50 Å, resulting in a predicted net decrease in energy transfer upon formation of the Zn(II)-bound complex. The WLC model can also be used to predict the influence of linker length on the Zn(II) affinity. When both metal binding domains are not connected, the formation of the ternary CFP-Atox1-Zn(II)-WD4-EYFP can be described using the following equilibria: CA + Zn(II) CA-Zn(II) CA-Zn(II) + WY CA-Zn(II)-WY

s = KCA

[CAZn(II)] [CA][Zn(II)]

s KWY =

[CAZn(II)WY] [CAZn(II)][WY]

(5a) (5b)

[CAZn(II)WY] [CA][WY][Zn(II)] (5c) Binding of Zn(II) to the single, peptide-linked protein sensors may be described analogously, using the following equilibria: s s KWY CA + WY + Zn(II) CA-Zn(II)-WY K s = KCA =

(6a)

(6b)

(6c) Binding of Zn(II) to the first protein domain (CA) can be assumed to be the s l ). Binding of the second metal binding domain = KCA same in both cases (KCA (WY) is intramolecular for the linked protein (Eq. (6b)) and intermolecular for the l ternary system (Eq. (5b)). The intramolecular binding constant KWY is related to s s l the intermolecular binding constant KWY by KWY = ceff KWY where ceff is the effective concentration of the second protein domain after formation of the initial Zn(II) complex. The effective concentration for the formation of an intramolecular complex is proportional to the probability density p(re ) for the end-to-end distance, re , that the linker needs to span in the complex, according to Eqs. (7) and (8) [27, 30, 31].


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Fig. 7 (A) Calculated effective concentration (ceff ) as a function of linker length for different values of the end-to-end distance in the complex. (B) KD values predicted by the WLC model assuming various values for the distance between the C-terminus of ATOX1 and the N-terminus of WD4 in the Zn(II) complex. For comparison, KD values that were experimentally determined for various CA-Lx-WY proteins are shown as circles

p(re ) (7) 4πre2 p(re) (8) ceff = NAV where re , lp , and lc are in dm and NAV is Avogadro’s number. Figure 7a shows the dependence of the effective concentration as a function of linker length for several values of re . The model predicts that an increase in linker length can give rise to either an increase or a decrease in effective concentration, depending on the distance that the linker needs to bridge in the complex. The dissociation constant for complex formation (KD ) can be calculated using ceff and the formation constant (Ks ) for the ternary complex, which was previously determined to be 4.5 × 1015 M−2 using Eq. (9). p(re ) =

K l = ceff K s 1 1 KD = l = K ceff 4.5 × 1015 M−2

(9)

The experimental KD values agree very well with the calculated values predicted by the WLC model for an end-to-end distance of 50 Å (Fig. 7b). Thus, analysis of the distance distribution of the linker in terms of random coil behavior fully explains both the magnitude and the linker length dependence of the affinity enhancement.

5 Chelating Fluorescent Protein Chimeras as Efficient Zn(II) Sensor Proteins The example discussed above shows that the sensor properties of FRET sensors with long flexible peptide linkers can be rationalized and even be predicted [34].

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However, the CALWY sensors showed a relatively small change in emission ratio upon Zn(II) binding. Although the dynamic range can be improved by further decreasing the linker length, the WLC models also predict a significant decrease in Zn(II) affinity when using shorter peptide linkers. One problem encountered here (and in many other FRET sensor systems) is that the fluorophores are relatively far apart in the ligand-bound state, resulting in a low-energy transfer efficiency. In the ligand-free state, a distribution of conformations is present that has an average energy transfer efficiency that is also rather low, resulting in a small net change upon ligand binding. One approach to obtain sensor proteins with an intrinsically large difference in FRET between the two sensor states is to create complementary target binding sites directly at the surface of the donor and the acceptor fluorescent domains. This approach is particularly attractive for metal binding sensors. The factors that determine metal complex stabilities are relatively well understood [35], thus allowing successful creation of de novo metal binding sites on protein surfaces [36]. Metal binding to each of the two fluorescent domains should result in a complex in which the two fluorophores are at a fixed position and as close together as sterically possible. To demonstrate the feasibility of this approach we took advantage of a report by Jensen and coworkers who found that the interaction between ECFP and EYFP can be enhanced by the introduction of Zn(II)-coordinating residues (Ser208C and Tyr39His) at their ‘natural’ dimerization interface [37]. Wild-type GFP has a wellknown tendency to dimerize at high concentrations (Kdim = 0.11 mM) [38], forming a complex in which both β-barrels are in an anti-parallel orientation [39]. Jensen and coworkers found a weak Zn(II) affinity of 0.7 mM when mutant ECFP and EYFP were present at 10 μM, but we reasoned that an ECFP-linker-EYFP sensor should display a substantially higher affinity due to the high effective concentration of the two domains. Introduction of the Ser208Cys and Tyr39His mutations on both ECFP and EYFP in CLY9 yielded a zinc sensor (called ZinCh-9) that displays an impressive 4-fold change in emission ratio upon addition of Zn(II) [34]. Unexpectedly, Zn(II) titrations showed a biphasic Zn(II) sensitivity over a large range of Zn(II) concentrations between 10 nM and 1 mM (Fig. 8). The presence of high (KD = 100 nM)- and lowaffinity Zn(II) binding (KD = 50 μM) sites is inconsistent with the formation of an anti-parallel complex between ECFP and EYFP. Such a complex would be expected to have two equivalent Zn(II) binding sites, each containing one Cys208 and one His39 ligand. A second reason to believe that the Zn(II) complex is different from that observed for the GFP dimer is that formation of this complex is highly unlikely for sensor constructs with short linkers such as ZinCh-1, which was shown to form similar Zn(II) complexes as ZinCh-9. We therefore examined the Zn(II) binding properties of sensor variants that contained only the Ser208Cys mutation (CLY9S208C) or the Tyr39His mutation (CLY9-Y39H). CLY9-S208C showed the same high affinity for Zn(II) as ZinCh-9, but not the additional increase in emission ratio at higher Zn(II) concentrations (Fig. 9). CLY9-Y39H on the other hand showed very weak Zn(II) binding. These results are consistent with a model in which ECFP and EYFP interact in a parallel orientation that allows the two Cys208 residues to


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Fig. 8 Zn(II) titration experiments for ZinCh-9. (a) Emission spectra of ZinCh-9 in the presence of increasing amounts of Zn(II), normalized at the emission maximum of ECFP at 475 nm. (b) The EYFP/ECFP emission ratio of ZinCh-9 as a function of Zn(II) concentration shows a biphasic response. Data were obtained using Ba(II)-EGTA as a Zn(II) buffer for low Zn(II) concentrations (circles) and a non-buffered titration for high Zn(II) concentrations (squares). Figures reprinted with permission from reference [34]. Copyright 2007 Elsevier

Fig. 9 Comparison of the Zn(II) binding properties of ZinCh-9 (open circles), CLY9-S208C (closed circles), CLY9-Y39H (squares), and CLY9 (triangles) showing the EYFP/ECFP emission ratio as a function of Zn(II) concentration. Figure reprinted with permission from reference [34]. Copyright 2007 Elsevier

form a high-affinity Zn(II) binding site. Following the formation of the initial Zn(II) complex, the two His39 residues are pre-organized to form a second low-affinity binding site. To gain more insight into the changes that accompany each Zn(II) binding step, Zn(II) titrations were also monitored using fluorescence anisotropy measurements (Fig. 10). The fluorescence emission anisotropy observed using direct excitation of EYFP showed that the high-affinity Zn(II) binding results in an increase in anisotropy from 0.304 ± 0.004 to 0.328 ± 0.004. This increase in anisotropy indicates an increase in the apparent molecular volume of ∼100% and is thus consistent with complex formation between ECFP and EYFP. To learn about the changes in the relative orientation of the ECFP and EYFP chromophores, the change in anisotropy for EYFP emission was determined after excitation of ECFP. In the absence of Zn(II), the anisotropy is low (0.09), which is consistent with a random orientation of the EYFP domain with respect to the ECFP domain. Binding of Zn(II) to the

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Fig. 10 Zn(II) binding to ZinCh-6 monitored using fluorescence anisotropy. (A) Fluorescence emission anisotropy at 527 nm using 500 nm excitation (ryy ). (B) Fluorescence emission anisotropy at 527 nm using 420 nm excitation (rcy ). The solid line is a fit of the data using Eq. (5.11) with Kd1 = 0.35 μM and Kd2 = 50 μM. Figures reprinted with permission from reference [34]. Copyright 2007 Elsevier

high-affinity site results in a substantial increase in anisotropy to ∼0.19, indicating that a more fixed orientation between ECFP and EYFP is formed. Population of the second Zn(II) site results in a decrease in anisotropy to 0.13. This decrease in anisotropy is most likely due to a second conformational change, in which the transition dipole moments of ECFP and EYFP are oriented in a way that results in an increase in energy transfer efficiency due to a more favorable κ 2 , and, at the same time, a larger amount of depolarization (Fig. 11).

Fig. 11 Model of Zn(II) binding to ZinCh sensor proteins. Zinc binding to the two Cys208 residues stabilizes the formation of an intramolecular complex of ECFP and EYFP in a parallel orientation, resulting in a large increase in FRET. After binding of Zn(II) to the high-affinity site, the two His39 s are pre-organized to form a second, low-affinity Zn(II) binding site. Binding to this low-affinity site is accompanied by a small conformational change, resulting in a further increase in FRET. Figure adapted from reference [34] with permission. Copyright 2007 Elsevier

Assuming that the fluorescent domains adopt a conformation in which the two Cys208 residues are facing each other and the domains are in a parallel orientation, the distance between the fluorophores can be estimated to be ∼31 Å. The energy transfer efficiencies of 80 and 85% that are observed for binding of the first and the


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second Zn(II) ions then imply values of κ 2 of 0.19 and 0.27, respectively. Since κ 2 can vary between 0 and 4 it is clear that orientations of the transition dipole moments in these complexes are still far from optimal for energy transfer. Further optimization of the energy transfer in the complex form might be achieved using circular permuted forms of one of the fluorescent domains. However, rational prediction of these effects is difficult, particularly because the transition dipole moment of ECFP is still unknown.

6 Taking Advantage of ‘Stickiness’: FRET Sensor Proteins Based on Conformational Switching GFP and many of its color variants are known to dimerize at high (local) protein concentrations. The use of A206K or A206R mutants has therefore been advocated to prevent the formation of intramolecular interactions between fluorescent domains in FRET-based sensors [18]. Before we show how deliberate promotion of intramolecular interactions can actually be used to enhance the ratiometric response of FRET-based protease sensors, it is useful to first discuss an apparent contradiction between our findings and results obtained by Tsien and coworkers who reported a dissociation constant for the GFP dimer (Kdim ) of 0.11 mM [38]. The WLC model predicts effective concentrations in the millimolar range for the ECFP and EYFP domains when connected via a flexible GlyGlySer-linker, yet our studies clearly show the absence of significant interactions between ECFP and EYFP. The solution to this apparent contradiction may be found in the type of complex that can be formed. In the absence of a linker, GFP dimerizes in an anti-parallel orientation, at least according to the X-ray structure [39]. In our sensor proteins, the antiparallel orientation is strongly disfavored, because this orientation would result in an entropically unfavorable extended linker conformation. This explanation implies that Kdim for the formation of the parallel complex is significantly higher than the predicted effective concentration for this complex, which is ∼3 mM for the longest linker. Although wild-type ECFP and EYFP do not interact when connected by flexible SGGSGG linkers, we showed that subtle redesign of the dimerization interface by substitution of the hydrophilic Ser208 by a hydrophobic phenylalanine promotes intramolecular complex formation between the two fluorescent domains, resulting in a large increase in energy transfer (Fig. 12) [21]. The introduction of a second mutation at the dimerization interface, V224L, did not by itself enhance intramolecular interactions between ECFP and EYFP, but it did induce a further increase in energy transfer efficiency in the presence of the S208F mutation. Cleavage of the linker by a protease resulted in a 16-fold decrease in emission ratio (Fig. 13). The absence of any FRET after cleavage shows that the complex between these more ‘sticky’ variants of ECFP and EYFP is readily disrupted upon cleavage of the linker and no intermolecular complex is formed at the low micromolar concentrations used in these experiments.

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Fig. 12 Improvement of FRET-based sensor proteins by redesign of the dimerization interface between ECFP and EYFP. (A) In the classical sensor design the random coil behavior results in a distribution of conformations with a moderate energy transfer efficiency. (B) Introduction of S208F and V224L mutations promotes intramolecular complex formation, resulting in a substantial increase in energy transfer efficiency prior to protease cleavage. Figures reproduced with permission from reference [21]. Copyright Wiley-VCH Verlag GmbH & Co. KGaA

Fig. 13 (A) Emission spectra of CLY9 and various mutants before linker cleavage (excitation at 420 nm). (B) EYFP/ECFP emission ratios of CLY9 and mutant variants, monitored over time after the addition of proteinase K. Figures reproduced with permission from reference [21]. Copyright Wiley-VCH Verlag GmbH & Co. KGaA

Promoting intramolecular domain interactions in either the on or the off state of a sensor as we did in the protease sensor could be a more general approach to rationally increase the ratiometric response of FRET-based sensor proteins. Given the high sequence and structural similarities among the many color variants of GFP,


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these mutations are generally applicable. In fact, S208F and V224L are both present in YPet, suggesting that the improved FRET properties of YPet (and CyPet) are largely due to an increased ‘stickiness’ [20, 21]. To allow more rational application of these mutations, it will be useful to actually determine the dissociation constant for the S208F/V224L mutants. For now, the KD for this interaction can be estimated to lie somewhere in the range between 10 μM and 1 mM. A KD lower than 10 μM would result in complex formation in the absence of the linker, while a KD higher than 1 mM would be inconsistent with the intramolecular complex formation that is observed in the presence of the linker.

7 Conclusion and Outlook FRET between fluorescent protein domains is a powerful and generally applicable approach to translate a protein conformational change into a ratiometric fluorescent read-out for intracellular imaging. Rational design strategies to improve the ratiometric response of FRET sensors are important in order to broaden the scope of this technology, for example, to allow high throughput applications based on fluorescence plate readers and fluorescence-assisted cell sorting (FACS) [40]. Long, flexible peptide linkers consisting of GlyGlySer repeats are attractive building blocks for FRET-based sensors, because simple models exist that can quantitatively describe their random conformational behavior. We have shown how these models can be used to understand two important properties of FRET-based sensor proteins, energy transfer efficiency and the strength of intramolecular interactions between receptor domains. The concept of effective concentration provides a useful framework to understand and predict conformational preferences in multidomain proteins, such as FRET sensors. Redesigning the GFP dimerization interface was shown to be an attractive strategy to obtain FRET sensors with an intrinsically large change in emission ratio. Introduction of Zn(II) binding residues yielded a Zn(II) sensor with a 4-fold change in emission ratio, whereas the introduction of hydrophobic residues enhanced the ratiometric change of a protease sensor from 4to 16-fold. These examples illustrate that the distance dependence of energy transfer efficiency can be employed successfully in rational design approaches. Given the difficulty of controlling the precise orientation of protein–protein complexes, it will probably remain difficult to also include the orientational dependence of energy transfer into the rational design of FRET sensors. Instead, directed evolution approaches combined with high throughput screening methods such as FACS are probably best suited to optimize the orientation between donor and acceptor dipole moments. Acknowledgments I would like to thank all current and previous members of my group working on FRET-based sensor proteins. In particular, I would like to acknowledge Ir. Jan Vinkenborg, Dr Toon Evers, Dr Melissa Koay, and Dr Misha Golynskiy for critically reading this chapter. This work was supported by a Human Frontier of Science Program Young Investigator Grant (RGY0068-2006C)

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Fluorescence Imaging of Calcium Loading and Mitochondrial Depolarization in Cancer Cells Exposed to Heat Stress Olaf Minet, Cathrin Dressler, Jürgen Beuthan, Urszula Zabaryło, Rasa Zukiene, and Vida Midaziene

Abstract One main issue of thermotherapy is the stress response of mitochondria to heat. Thermotherapies function by inducing lethal heat inside target tissues. Actually spatial and temporal instabilities of temperature distributions inside target volumes require optimized treatment protocols and reliable temperature-control methods during thermotherapies. Since solid tumors present predominant targets to thermotherapy, on the one hand, the hyperthermic stress-induced effects on mitochondrial transmembrane potentials in breast cancer cells (MX1) were analyzed. On the other hand, the intracellular Ca2+ fluctuations in different cell types responding to heat stress were investigated. Heat sensitivities and stress reactions might be extremely different among different tissue species and tissue dignities; therefore, it is very important to investigate tissue-specific stress responses systematically. Even though this chapter contributes little information, only, to the enlightenment of systemic cellular heat stress mechanisms, it may support the fortification of basic knowledge about systemic stress responses. Using cytoplasmic and intramitochondrial fluorescent Ca2+ probes it was possible to compare hyperthermia-induced changes in the Ca2+ distribution between the cytoplasm and the mitochondria of normal and tumor cells and to examine the relationship between mitochondrial Ca2+ concentration and changes in the viability of these cell types upon hyperthermic treatment. We compared Ca2+ concentrations in cytoplasm and mitochondria in cancerous CX1 and MX1 cells with normal CHO cells after transfer from room temperature (25◦ C) to 37◦ C or 43◦ C. Sudden increase in the incubation temperature (from room temperature to 37◦ C) induced very different cytoplasmic and mitochondrial Ca2+ fluctuation patterns in normal CHO and CX1 and MX1 tumor cells. Estimating the CX1, MX1, and CHO cell viabilities upon hyperthermic treatment, we found that thermosensitivities increase in the sequence CX1590 nm long-pass filter) and the green fluorescence channel (λexcitation : 450–490 nm band-pass filter, λdetection : >515 nm long-pass filter) each. The exposure times for optimal fluorescence imaging were digitally measured by the AxioCam MR camera for each image (variation range: 268–2354 ms). Ca2+ indicators and MTG were imaged with an Axioplan 2 (Carl Zeiss, Germany) microscope equipped with a 100× water immersion objective, AxioCam MRc CCD camera (3900 × 3090 pixels). Rhod 2 was excited at 546+12 nm by a band-pass filter. The fluorescence light was detected by a long-pass filter

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>590 nm. Fluo 3 was excited at 450–490 nm by a band-pass filter. The fluorescence light was detected by a long-pass filter >515 nm. The fluorescence image was acquired by an AxioCam MRc camera, with the exposure time ranging from 268 to 2354 ms. Both microscopes were triggered by Zeiss AxioVision 4.5 software. Data processing and statistical evaluation: Ratios of the fluorescence amplitudes measured in the red and the green channel were evaluated for every image frame after correcting the amplitudes with the exposure times applied in each case. The average value of each temperature group was used for statistical assessment of red and green fluorescence intensities.

3 Results 3.1 Cell Viability The Alamar Blue assay shown in Fig. 4 clearly demonstrated that CX1 cells are more thermoresistant than MX1 and CHO, maintaining a viability of up to 100% in the temperature range from 42◦ C to 50◦ C, while the viability of MX1 cells at 45◦ C had decreased to approximately 80% and to about 30% at 50◦ C. The viability of CHO cells is similar to that of MX1 except that the two cell types show stimulatory effects at different sublethal stress temperatures. While the MX1 cell activity was stimulated at 40◦ C, CHO cells were slightly inactivated at 40◦ C but stimulatory effects were reactivated at 42◦ C. Finally, the viabilities of all cell types drastically decreased with stress temperatures rising above 45◦ C (Fig. 4). The obtained results indicated that the resistance of normal CHO culture cells to hyperthermic stress is not higher than the resistance of two tumor cell lines, and even much lower than that of CX1. This raised some doubts as to what extent the data on cultured cells may be extrapolated for the understanding of similar effects

Fig. 4 CX1, MX1, and CHO cell viabilities determined by Alamar Blue assay (BIOSOURCE International Inc., USA)

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on corresponding tissues (healthy versus tumor) since in general normal cells are much less stable in culture than are tumor-derived cells.

3.2 Visual Evaluation of Fluorescent Ca2+ Indicators Exposure times required for optimal fluorescence image acquisition were different for every image, ranging from 268 to 2354 ms; thus the fluorescence intensity had to be corrected mathematically. Using visual criteria, though, the staining quality, indicator compartmentalization, and cell structure were assessed. The real-time leak of the fluorescent indicator from the cell was visually inspected during hyperthermic treatment. All cell types used were stained with MTG in order to determine the mitochondrial localization and to prove that the Rhod 2 AM indicator is compartmentalized in mitochondria. Visual inspection was necessary because the equipment used did not permit an overlap of images and, therefore, was insufficient for that purpose (images not shown). In Figs. 5–7 the micrographs of MX1, CX1, and CHO,

Fig. 5 MX1 control cells stained with Rhod 2 and Fluo 3 at 37ºC

Fig. 6 CX1 control cells at 37ºC and stained with Rhod 2 and Fluo 3

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Fig. 7 CHO cells stained with Rhod 2 and Fluo 3 at 37ºC

respectively, at 37◦ C after 0, 30, and 60 min are given. It can be seen that even at physiological temperature there are some changes in the fluorescence area and intensity. An indicator leak is visible after 60 min in all cases, especially in CHO cells, although acetoxymethyl esters of the fluorescent indicators Rhod 2 and Fluo 4 cleaved by cell esterase are supposed to be membrane impermeable. The labeling method (indicator concentration, loading time, loading conditions) was optimized using visual criteria. The optimum concentration of both indicators was found to be 2 μM, the most appropriate incubation time was 30 min, and best incubation temperature was room temperature (25◦ C). One may note from Figs. 8 and 9 that in agreement with earlier observations [52], fluorescences of both cytosolic and mitochondrial Ca2+ probes were much lower in CHO cells than in both tumor cells; however, in Figs. 8 and 9 the curves of MX1 were slightly higher than those of CX1. The Fluo 3 and Rhod 2

Fig. 8 Mitochondrial and cytosolic Ca2+ indicator fluorescence in CX1, MX1, and CHO cells at 37◦ C


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Fig. 9 Red and green indicator fluorescences depending on timescale at 43◦ C in MX1 cell culture

fluorescences exhibited distinct changes with time depending on the stress temperatures applied. The biggest changes in Rhod 2 and Fluo 3 fluorescences appeared during the first 10 min after transferring cells from room temperature to 37◦ C (Fig. 8) or 43◦ C (Fig. 9). This may be explained by staining and measuring temperature differences: after staining cells at room temperature, the primary measuring temperature was 37◦ C. Since room temperature was around 25◦ C, a 12◦ C temperature rise was a sudden hyperthermic condition for the cells which had to take time to equilibrate cellular Ca2+ perturbations. Cells were also stained at 37◦ C, but the staining quality was then not good enough and leakage of indicators was higher during monitoring. CHO cells have less Ca2+ in mitochondria and in cytosol than do the tumor cells tested. The stabilized concentration of mitochondrial Ca2+ at 37◦ C and the stabilization time after change of temperature from room temperature to 37◦ C in MX1, CX1, and CHO cells are shown in Fig. 10. Cells of both tumor cell lines contained more Ca2+ in mitochondria than in cytosol. MX1 cells were more sensitive to heat than were CX1 – the rise in the Ca2+ concentration that followed the temperature shift by +12◦ C (from room temperature of 25◦ C to 37◦ C) was much higher (Figs. 8– 10) and the stabilization time (Fig. 10) was much longer in MX1 and CX1 cells when compared to CHO cells. MX1 cells were shown to be most sensitive to hyperthermia and contained the highest Ca2+ concentration compared to the other two cell types (Fig. 10). It took 40 min to stabilize mitochondrial Ca2+ after transferring MX1 cells from the room temperature of 25◦ C to 37◦ C. CHO cells contained the lowest Ca2+ concentration among the cell types tested and were less sensitive to hyperthermia than MX1. After 15 min the Ca2+ concentration inside CHO cells was stabilized. A clear oscillatory response in Fluo 3 and Rhod 2 fluorescences, as well as the oscillation patterns, depended on temperature and cell type. They exhibited the largest perturbations of mitochondrial Ca2+ after transfer from room temperature to 37◦ C.

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Fig. 10 Stabilized concentration of mitochondrial Ca2+ at 37◦ C and stabilization time after change of temperature from room temperature of 25◦ C to 37◦ C in MX1, CX1, and CHO cells

There were clear differences in the patterns of fluorescence changes for both Ca2+ probes induced by transfer to 37◦ C as compared to supraphysiological temperature of 43◦ C, and the difference was much larger with respect to the behavior of the mitochondrial probe Rhod 2, in particular for tumor cells. Hyperthermic stress at 43◦ C induced an initial mitochondrial Ca2+ ‘wave’ in all three cell types; however, it was more sudden and followed by a very steep decrease in Rhod 2 fluorescence in both tumor cell lines. It remains to be determined whether this decrease might be caused by MTP formation. Remarkably, Rhod 2 fluorescence at 43◦ C dropped to the lowest extent in the MX1 cells most sensitive to temperature, pointing at a possible causal connection between the mode of mitochondrial Ca2+ changes, MTP formation, and cellular resistance to heat. It is also noteworthy that the decaying oscillatory decreases in MX1 mitochondrial Ca2+ probe (Rhod 2) fluorescence, both at 37◦ C and 43◦ C, were correlated with increases in cytosolic Ca2+ probe fluorescence, indicating a periodic redistribution of Ca2+ ions between these two cellular compartments. Thus, the obtained data indicate cell-specific pattern of changes in Ca2+ compartmentalization as a part of short-term cellular response to heat shock.

3.3 Fluorescence Imaging of Mitochondrial Transmembrane Potential The heat responsibilities of mitochondria in MX1 breast carcinoma cells were investigated in a hyperthermic range between 40◦ C and 56◦ C. Spatial mitochondrial distributions were imaged by structure labeling with mitochondria-specific MitoTracker Green FM (MTG). The functional states of mitochondria were analyzed by membrane potential-dependent fluorescence labeling. The mitochondrial transmembrane potential (Ψ m ) sensor JC-1 was used. Depending on the stress temperature the cells were exposed to, the mitochondria were more or less depolarized, leading to an increase in green fluorescent JC-1


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monomers accompanied by a decrease in orange-red J-aggregates. This effect was rather weak under comparatively mild hyperthermic stress conditions between 40◦ C and 45◦ C, while under severe stress at 50◦ C or 56◦ C, the response was more pronounced and clearly documented by very low Ψ m . Mitochondria in MX1 cells consequently react to heat stress in Ψ m depolarizing with temperature-dependent extents. After labeling MX1 mitochondria with MTG more or less evenly, cytoplasmic distributions inside intact control cells (37◦ C) were observed. Because of comparatively high organelle numbers per cell, single mitochondri could not be distinguished from each other with the optical magnification applied (Fig. 11). Under mild stress conditions at 40◦ C, the mitochondrial fine structure was already diminished compared to the control. The 42◦ C group showed a clear aggregation of mitochondria. This process was even more obvious in the 45◦ C group, definitely supported by the changes in cell morphologies and sizes, since MX1 cells were smaller and round

Fig. 11 Mitochondrial fluorescence in heat-stressed MX1 cells after labeling with MitoTracker Green FM (Molecular Probes, Invitrogen, Germany). Panel (A) shows control cells incubated at 37◦ C, panels (B)–(F) show heat-stressed cells treated with the temperatures indicated at every panel for 30 min each

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after stress at 45◦ C. Treatments at more severe stress conditions (50◦ C or 56◦ C) resulted in pronounced diffuse pancellular fluorescence distributions, although cell morphologies again resembled the control cells as well as the 40◦ C and the 42◦ C stress groups. This observation supported the effect that treatment for 30 min at 45◦ C induced an active stress response when the cells shrunk and were rounding up, while higher stress temperatures did not permit active cell responses anymore (Figs. 11 and 12). Since MTG labeling did not give any information about functional states of heatstressed mitochondria, the MX1 cells were then labeled with the mitochondrial membrane potential sensor JC-1 in order to monitor heat stress-induced effects on Ψ m . The fluorescence micrographs in Fig. 12 clearly showed membrane potential correlated labeling of mitochondria in MX1 cells. With increasing stress temperatures at 40◦ C, 42◦ C, 45◦ C, 50◦ C, or 56◦ C, the m -dependent orange-red and green fluorescences in MX1 cells labeled with JC-1 were altered when compared with control cells. The orange-red fluorescence of J-aggregates at mitochondria with high Ψ m was imaged in both fluorescence channels, while green fluorescence with shorter wavelengths of monomer JC-1, of course, was imaged in the green channel only. After the mitochondria in the control group and the 40◦ C, 42◦ C, and 45◦ C stress groups exhibited orange-red fluorescence, the signal color in the red channel was changed in the 50◦ C and 56◦ C stress groups. Hyperthermic temperatures at 50◦ C or 56◦ C caused JC-1 to show deep red fluorescence (Fig. 12). A direct readout of fluorescence intensities imaged in both channels for every object was not possible since image acquisition times were digitally adjusted for optimal fluorescence imaging. Therefore the ratio of fluorescence amplitudes measured in the red and the green channel was evaluated (red/green ratio) for every object or every image pair, respectively. Average red/green ratios of the control group and the different heat stress groups were plotted against the temperatures at which MX1 cells were treated (Fig. 13). The red/green fluorescence ratios were shown to decrease with increasing stress temperatures in a nonlinear manner, because orange-red fluorescence intensities discontinuously decreased, while green fluorescence intensities increased and consequently the red/green ratios declined. The nonlinear interrelation between mitochondrial depolarization and stress temperature is presented by the exponential function shown in Fig. 13. Here the red-to-green fluorescence intensity ratios decayed rather rapidly in the

Fig. 12 (continued) Green (left) and red (right) fluorescence of heat-stressed MX1 cells after mitochondrial transmembrane potential-dependent labeling with JC-1 (Molecular Probes, Invitrogen, Germany). Panel (A) shows control cells incubated at 37◦ C, panels (B)–(F) show heat-stressed cells treated with the temperatures indicated at every panel for 30 min each. Exposure times employed for acquiring the images were the following: panel (A) 645 ms green, 268 ms red; panel (B) 580 ms green, 295 ms red; panel (C) 711 ms green, 633 ms red; panel (D) 571 ms green, 656 ms red; panel (E) 765 ms green, 1172 ms red; panel (F) 677 ms green, 2345 ms red


Fig. 12 (continued)

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Fig. 13 Average ratios of fluorescence amplitudes measured in the red and the green fluorescence channels. Red/green ratios indicate changes in the quantitative relation between orange-red fluorescent J-aggregates in MX1 cells with high Ψ m and green fluorescent JC-1 monomers in cells with low Ψ m . Standard deviations are given as mean square deviations

temperature range 40–45◦ C and switched over to a more decelerated decay at higher temperatures following an exponential curve progression as illustrated in Fig. 13. In Fig. 14 the fluorescence intensities in the green and the red channels were separated and illustrated as monochromatic 3D profiles. Generally, these results indicated that the amount of red fluorescent J-aggregates accumulating at mitochondria with comparatively high membrane potentials decreased with increasing stress temperatures opposite to the enhanced green fluorescence of J-monomers in cells with more or less depolarized Ψ m . In the control group, the red/green ratio was approximately 1.7. Under mild heat stress conditions at 40◦ C or 42◦ C, the red/green ratios decreased but slightly compared with those of the control cells. The ratios in these groups were definitely above 1, denoting that the orange-red signal intensities of J-aggregate fluorescence were higher than the green fluorescence signal intensities. With higher stress temperatures at 45◦ C and 50◦ C, the ratios drastically decreased to values below 0.7, because the red fluorescence intensities were weaker than the green fluorescence intensities. Exposing the cells to temperatures of 56◦ C resulted in pancellular green and red fluorescence distributions, whereas the green signal intensities were much higher than the red signal intensities, which represented only residual background fluorescence. Therefore, in the 56◦ C stress group, the average red/green ratio was again drastically decreased to reach a value of approximately 0.2, showing that the red fluorescence intensity was approximately 20% of the green fluorescence intensity measured in this experimental group. The red signal intensity decrease resulted from massive depolarization of Ψ m under exposure to 56◦ C heat accompanied by a destruction of structural organelles due to which the green fluorescence of JC-1 monomers inside severely damaged MX1 cells was enhanced. The fluorescence intensity distributions inside cells imaged in the red and the green fluorescence channels were derived as three-dimensional (3D) illustrations. Characteristic examples of 3D fluorescence intensity profiles are shown in Fig. 14.


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Fig. 14 Green and red fluorescence distributions in heat-stressed MX1 cells after mitochondrial membrane potential-dependent labeling with JC-1. Each of the regions of interest was divided into red and green data fractions for the green and the red fluorescence channels and

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Fig. 14 (continued) illustrated as 3D fluorescence distributions after their noise was reduced by a nonlinear digital total variation filter [71]. Panel (A) shows control cells incubated at 37◦ C, panels (B)–(F) show heat-stressed cells treated with the temperatures indicated at every panel for 30 min each. Two representative cells are shown in every panel


Fig. 14 (continued)

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Here, the fluorescence amplitudes were corrected with the respective acquisition times in order to compare the distributions of absolute signal intensities with each other. Each 3D image is z scaled to the maximum of fluorescence intensity. The red fluorescence intensity maxima (RFIM) were highest and nearly constant in the temperature range between 37◦ C and 42◦ C. From a stress temperature of 45◦ C, RFIM drastically declined, while the green fluorescence intensity maxima (GFIM) exceeded RFIM. This again indicated an amplified depolarization of mitochondria in response to hyperthermic stress.

4 Discussion In this review the heat stress responses from 40◦ C to 56◦ C, for 30 min each, of three different cell species (CX1, MX1, CHO) at the metabolic and mitochondrial level were questioned. Since Ca2+ presents a key factor of metabolic and signaling pathways in cells, the heat-induced Ca2+ fluctuations in these cell types were compared. In this subcoagulative temperature range, obvious mitochondria-involving stress responses were expected. As stress-sensitive indicators, mitochondrial and cytoplasmic Ca2+ fluctuations were analyzed by fluorescence microscopy after selective staining with fluorescent Ca2+ indicators. The network of Ca2+ transport into cells and inside cells is schematized in Fig. 1. Spectral properties of the indicators applied in our experiments are illustrated in Fig. 2. Hyperthermia has been proposed a potential therapeutic implementation in clinical cancer therapy [4]. The cellular mechanisms involved in heat stress responses and their impact on various subcellular structures so far have been described only for selected tissue models. Funk et al. studied heat stress-induced changes of mitochondrial morphologies in astrocytes and MDCK cells by video-enhanced contrast microscopy using a perfusion cell chamber system [23]. In their study the morphological alterations inside mitochondria exposed to moderate hyperthermic stress conditions were revealed to be reversible. Recovery of mitochondrial changes subsequent to heat treatment was also detected in microglial cells [27]. Metabolic investigations on microglial cells revealed dropping of the physiological ATP content by 60% 1 h after a 20-min heat shock at 45◦ C, indicating that heat stress leads to scaling down of energy resource ATP in this cell type [28]. In heat-shocked rats, failing of energy metabolism and ATP depletion were detected as the earliest cell-damaging factors of ischemic insult [72]. These data emphasize the interface actions of energy-supplying processes during cellular stress responses. The crucial involvement of Ca2+ in energy supply and stress-induced responses is out of question. Our results clearly documented MX1 cells to be the most thermosensitive among the cell types tested here. On the other hand, CX1 cells were shown to be the most thermoresistant type. Since MX1 and CX1 are both cancer cell species, the normal CHO cells were positioned right in the middle concerning thermosensitivity and therefore, no implication may be drawn predicating tumor or normal cells to be more or less thermosensitive than the others as compiled in Table 1.


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Table 1 Comparison of cancer cells MX1 and CX1 with normal CHO cells Feature

CHO

MX1

CX1

Tissue dignity Mitochondrial Ca2+ concentration Stabilization time of mitochondrial Ca2+ level after transfer from 25◦ C to 37◦ C (see Fig. 10) Thermosensitivity

Normal Low 15 min

Malignant High 40 min

Malignant High 30 min

Sensitive

Sensitive

Resistant

From a therapeutic point of view it certainly would be desirable if tumor tissues were more sensitive to heat than were healthy tissues. Unfortunately this is not the case. This fact imposes the obligation on scientists working on thermotherapy to study heat stress responses in as many cell species as possible with adequate care. Several subcellular structures have already been analyzed in MX1 cells after heat stress. In a previous study the F-actin cytoskeleton was shown to be thermally more sensitive than the plasma membranes, since F-actin fibers exhibited morphological alterations under comparatively mild stress at 40◦ C or 42◦ C, while plasma membrane morphologies were not affected under these conditions. Only temperatures higher than 42◦ C induced detectable morphological changes in plasma membranes [73, 74]. These results did not give any information about functionalities of investigated organelles. It was shown previously that the MX1 cell viability was not attenuated after stress at 40◦ C or 42◦ C. When exposed to 45◦ C or higher temperatures, the cell viabilities were increasingly diminished. Comparing these results it should be concluded that the reorganization of F-actin cytoskeletons in MX1 cells resulted from an active stress response under hyperthermic stress at 40◦ C or 42◦ C, which is supposed to be compensated during cellular recovery from sublethal heat stress. It was also demonstrated earlier that MX1 cells undergo necrosis during 30min treatment at 56◦ C, but not at 50◦ C or lower temperatures, because cells exposed to 50◦ C did not exhibit necrotic phenotypes [74]. Therefore, total depolarization of mitochondrial membranes in cells stressed at 56◦ C was an inevitable result of necrosis. Mitochondrial permeability transition in general is an early sign of the initiation of cellular apoptosis or necrosis, provoking a collapse in the electrochemical gradient across mitochondrial membranes [75–77]. Several studies have documented Ψ m to be a parameter sensitive to various kinds of environmental disturbances [75, 78–80]. Ψ m is established by the proton pumps powering the mitochondrial electron transport chain and is significantly involved in energy-providing processes and calcium metabolism [80]. In a feedback mechanism, calcium regulates the mitochondrial functions. Consequently, the mitochondrial calcium concentrations can give information about the current physiological or pathological state of the cell under investigation [82]. Knowledge about specific cellular responses to sublethal stress temperatures will contribute to improved thermotherapeutic protocols. In this connection, it is important to realize that there are no unequivocally definable borderlines between mild and severe heat stress or sublethal and lethal heat. Many factors may affect the thermosensitivity or thermoresistance of a target tissue, like cell species, tissue origin,

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cell cycle, developmental stages, and heat-exposure times. Possibly, there might be a rough borderline between lethal and sublethal heat stress [14], which again might be influenced by many intrinsic and environmental factors. As the central factory of cellular energy metabolism, mitochondria present cardinal targets of hyperthermic stress and other nonthermal stressors [23–25, 27]. Mitochondrial dysfunctions lead to a drastic decrease in cellular ATP levels and phosphorylation efficiencies [76, 77], thus triggering problems with energetic supplies. When permanent mitochondrial depolarization was detected in MX1 cells after heat stress, a nonlinear interrelation between depolarization dynamics and stress temperatures was revealed [83]. This response was reflected by an exponential decay function describing the average ratios of fluorescence amplitudes measured for the membrane potential sensor JC-1 in the red and the green emission channels (Fig. 13). The standard deviation, which is comparatively large in this case, should be reduced by increasing the number of investigated micrographs. In myocytes also, an exponentially proceeding depolarization of mitochondria was detected after application of FCCP [4-(trifluoromethoxy)phenylhydrazone]. The decrease in Ψ m -dependent tetramethylrhodamine ethyl ester (TMRE) fluorescence was measured in this study when spontaneous transient Ψ m depolarizations were observed [84]. These depolarizations were reversible and not permanent as in our investigations [83]. Apparently, mitochondrial depolarizations follow exponential dynamics. Nevertheless, this result supported the common statement that mitochondria take central positions during cell death [85]. The nonlinear response of MX1 mitochondria to hyperthermic stress corresponded very well with our results obtained using various microscopic techniques for investigating heat stress responses of different subcellular structures in MX1 cells [73]. In our previous studies also, nonlinear responses of plasma membranes and cytoskeletons in heat-stressed MX1 cells were observed [73, 74]. Applications of heat as therapeutic agent generally are conflicted with spatial and temporal variations in temperature distributions inside a target volume and surrounding structures [13]. It is also possible that different subcellular components in a certain cell species exhibit various heat sensitivities. Consequently, therapies inducing thermal destructions demand for highly thorough and specific intervention protocols concerning volume and location of diseased tissue, blood supply and perfusion dynamics, heat dose, application geometry and control techniques, as well as the patient’s physical condition. Only a well-balanced interplay of all relevant parameters may ensure a successful thermotherapy.

5 Conclusions It is known that mitochondrial Ca2+ levels are elevated 3–5-fold in most cells derived from hepatic tumor tissue and by a remarkable 70-fold in Morris 7777 hepatoma (a rapidly growing tumor) [3]. This finding is in line with the facilitation of MTP formation in cancer cells under hyperthermic stress. However, there is no direct


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evidence for causal connection between the amount of Ca2+ and the ability of certain cell types to survive mild heating. Sensitivity can be caused by mitochondrial permeability transition pore (MPT), which is the result of an increased Ca2+ concentration in mitochondria and oxidative stress. Mitochondria could rapidly accumulate Ca2+ ; therefore, these organelles are very sensitive to Ca2+ overload. If a hyperthermic effect is obtained due to mitochondrial Ca2+ overload and consequent MPT formation, the higher the mitochondrial Ca2+ concentration at physiological temperature, the faster and easier the Ca2+ threshold concentration is reached. However, this hypothesis is challenged by the fact that supraphysiological hyperthermia induces a very fast deenergization of the mitochondrial inner membrane, slowing down the Ca2+ uptake by electrogenic Ca2+ uniporter. Under hyperthermic treatment, the plasma membrane and the endoplasmic reticulum membrane also lose their barrier functions and thus exaggerate cellular ion imbalance, particularly increasing the danger of perturbations in Ca2+ homeostasis and causing Ca2+ overload [86–88]. Under these circumstances, uncoupled mitochondria can no longer fulfill their task of Ca2+ excess clearance, decreasing cellular safety guarantees in emergency situations. The results indicate that the tumor cell sensitivity to hyperthermia may originate from the mitochondrial Ca2+ concentration and any changes may have been caused by raised temperatures. 1. The viability of the cells after hyperthermic stress is different: CX1 cells are more thermoresistant than MX1 cells, they maintain a viability of 100% in the temperature range from 37◦ C to 50◦ C, while the viability of MX1 cells at 45◦ C decreases to approximately 80% and at 50◦ C to about 30%. The viability of the CHO cell line is similar to that of MX1. 2. The concentration of Ca2+ in mitochondria and cytosol of tumor and normal cell lines was compared. The CHO cells (normal) have less Ca2+ in mitochondria and in cytosol than do CX1 and MX1 cells and they take less time to equilibrate cell metabolism. 3. The time dependence pattern of changes in cytosolic and mitochondrial Ca2+ concentration was monitored. The biggest response was registered within 10 min after the tumor cells were transferred from room temperature to 37◦ C or 43◦ C, while the CHO cells showed a much smaller increase in mitochondrial Ca2+ concentration. Our investigations on mitochondrial responses to hyperthermic stress in MX1 breast cancer cells revealed a comparatively high thermosensitivity since mitochondrial morphologies and cellular distributions already changed when exposed to a mild stress temperature of 40◦ C (30 min) after MTG labeling. Morphological changes were not directly accompanied by Ψ m depolarizations, because JC1-labeled mitochondria exhibited only minor Ψ m reductions after stress in a temperature range between 40◦ C and 45◦ C when compared with control cells. More severe stress conditions at 50◦ C or 56◦ C induced complete mitochondrial

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depolarization detected by massive decrease in orange fluorescence intensities shifting to weak red fluorescence. Orange fluorescence was emitted by J-aggregates accumulated at highly polarized mitochondria. In cells containing depolarized organelles, JC-1 formed green fluorescent monomers. Heat-induced depolarization of mitochondria was monitored by the absolute fluorescence intensity ratios evaluated for the maximum fluorescence amplitude measured in the red and the green fluorescence image of every object (red/green ratio). Our results may support further developments in the field of thermotherapeutic dosimetry and irradiation planning. Nevertheless, each (cancer) cell type has an individual pattern of response to the stress challenge and mitochondrial involvement is also different and dependent on cellular context – the biology of heat still presents an open field of unresolved questions and all experimental data will contribute more or less essential pieces to the puzzle of systemic stress response mechanisms. Acknowledgments We express our thanks to Jens-Peer Kuska of the Interdisciplinary Center for Bioinformatics of Universität Leipzig for his kind assistance in image processing.

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Energy Transfer in Silica Nanoparticles: An Essential Tool for the Amplification of the Fluorescence Signal Sara Bonacchi, Damiano Genovese, Riccardo Juris, Ettore Marzocchi, Marco Montalti, Luca Prodi, Enrico Rampazzo, and Nelsi Zaccheroni

Abstract The careful design of dye doped silica nanoparticles in order to induce controllable energy transfer processes can yield very sophisticated species able to perform precious and complex functions. They can be therefore exploited in many fields of great economical and social importance, such as medical diagnostics, molecular biology, and solar energy conversion. In this chapter, we present the characterization of some functionalized silica nanoparticles with a particular emphasis on the discussion of the the energy transfer processes at the basis of their properties. Since a careful design is fundamental in the realization of more and more sophisticated materials, we also discuss the synthesis of these systems, in order to suggest new routes for the preparation of such valuable and versatile objects.

1 Introduction Electronic energy transfer is a crucial photophysical process, since it is at the basis of essential natural phenomena such as photosynthesis, as well as of widely spread artificial molecular devices; moreover, it is a very valuable tool for measuring distances at the nanometre level. For artificial systems, the possibility to take profit of energy transfer processes has enormously increased with the advent of supramolecular chemistry [1], and a new additional boost is expected to be observed with the very rapid growth of nanotechnology, in many different fields, ranging from medical diagnostics to solar energy conversion. In the context of nanotechnology, a great and increasing interest is devoted to nanoparticles [2]. Nanoparticles can be made of many different materials, including polymers, metals, semiconductors or a combination of them. Their versatility and different properties have already gained them many industrial applications in a wide range of fields, such as electronics, medicine L. Prodi (B) Dipartimento di Chimica ‘G. Ciamician’, Latemar Unit, Università degli Studi di Bologna, Via Selmi 2, 40126, Bologna, Italy e-mail: [email protected] C.D. Geddes (ed.), Reviews in Fluorescence 2008, Reviews in Fluorescence 2008, DOI 10.1007/978-1-4419-1260-2_5, C Springer Science+Business Media, LLC 2010

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and material sciences. One of the cut-edge fields in nanoparticle research is the enhancement of biological imaging for medical diagnostics and drug delivery. In this chapter, we will focus our attention on the potentialities offered by the use of energy transfer processes in dye-doped or dye-coated silica nanoparticles (DDSN or DCSN, respectively) [3–10], which conjugate a simple and low-cost preparation with the possibility to obtain sophisticated, but robust, multifunctional systems, especially when they are designed as ‘onion-like’, multilayer structures. The design of these systems is not trivial and it has to be carefully studied to be able to foresee the photophysical characteristics of the resulting materials. On the other hand, despite the complexity of these systems, their synthesis is relatively simple and very versatile. These features make them a very powerful tool to obtain very complex and precious functions from low-cost and easy-to-prepare nanoobjects. In the next paragraphs we will present the most common synthetic strategies to obtain dye-doped and dye-covered silica nanoparticles and discuss many examples, but we would like to start introducing the energy transfer process and its great scientific and applicative importance.

2 The Principles of Energy Transfer Processes To understand the potentialities offered by a proper use of energy transfer processes, it is important to recall a few basic theoretical elements; it is however not the scope of this contribution to provide a detailed overview of these processes, and we suggest to the interested readers to look up one in between the impressive choice of texts on this particular topic [11–13]. Energy transfer, which can be described according to Eq. (1), may occur via radiative or nonradiative mechanisms, but in all cases must obey energy conservation, i.e. the energy of ∗ A must be equal or lower than the one of ∗ D. ∗

ken

D + A −→ D + A∗

(1)

Radiative energy transfer, usually referred to as trivial mechanism, implies the emission of a photon from the donor ∗ D, a photon that is then absorbed by the acceptor A. In this case, no kind of electronic interaction between ∗ D and A is required and the acceptor molecule cannot influence the emission ability of the donor, but it simply intercepts the photon before its detection. This kind of mechanism is, therefore, of little importance for the design of molecular or nanostructured devices and will no longer be treated here. Nonradiative energy transfer processes on the contrary require some kind of electronic interaction between the two species. This interaction can be typically divided into two additive parts, the coulombic interaction (also called Förster mechanism) [14] and the exchange interaction (Dexter mechanism) [15]. The exchange mechanism can be represented as a double electron transfer reaction, i.e. the excited electron on ∗ D ‘ jumps’ into the LUMO of A simultaneously with

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the transfer of an electron from the HOMO of A into the corresponding orbital on D. The exchange resonance interaction occurs via overlap of the electron clouds and it requires physical contact between the interaction partners. According to Dexter mechanism, the rate constant for energy transfer obeys Eq. (2): ken = Ae−β(r−rc )

(2)

where r is the centre to centre distance between the donor and acceptor and rc is the sum of the van der Waals radii of the donor and acceptor species. The value of β is typically close to 1 Å–1 , and it can become lower when the donor and the acceptor are bound together through a conjugate bridge. It is important to stress here that the Dexter mechanism is therefore efficient only at very short distances (6–20 Å). The spin selection rules for this type of mechanism arise from the need to obey spin conservation in the reacting pair as a whole. This allows the exchange mechanism to be operative in many cases in which the excited states are spin forbidden in the usual spectroscopic sense [1]. In doped silica nanoparticles, this could be the only possible mechanism when the dye is a metal complex, such as the largely used Ru(bpy)3 2+ [16–18]. The coulombic resonance interaction (Förster mechanism), on the contrary, occurs via the electromagnetic field and does not require physical contact with the interacting partners. The basic mechanism involves the induction of a dipole oscillation in A by D∗ . In the frame of the Förster mechanism, the energy transfer rate constant can be accurately predicted according to Eq. (3):

ken

D κ 2 = τD r6

9000( ln 10) JF 128π 5 Nn4

(3)

where n is the refractive index of the solvent, D and τ D are the emission quantum yield and lifetime of the donor in absence of the acceptor, respectively and r is the distance between the donor and the acceptor. κ 2 is a geometric factor, which depends on the relative orientation of the dipoles and is usually assumed to be equal to 2/3, which is appropriate for a dynamic random orientation of the donor and the acceptor. JF is the overlap integral between the emission spectrum of the donor F(λ) (normalized such that F(λ)dλ = 1) and the absorption spectrum of the acceptor (λ) and can be calculated from Eq. (4). ∞ JF =

FD (λ)εA (λ)λ4 dλ

(4)

0

The distance R0 at which the energy transfer has a 50% efficiency is named Förster distance and can be calculated according to Eq. (5):

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R0 =

D κ 2 9000(ln 10) JF 128π 5 Nn4

(5)

As a consequence, Eq. (3) can be written in the form of Eq. (6) that clearly shows the dependence of the energy transfer processes (and, as a consequence, its efficiency) on the distance between the donor and the acceptor. The resonance energy transfer, RET, is then a through-space interaction and may take place over large distances (20–100 Å).

ken

1 = τD

R0 r

6 (6)

As far as the selection rules are concerned, RET is expected to be efficient when the radiative transitions connecting the ground and the excited state of each partner have high oscillator strengths. Thus, the typical example of an efficient Förster mechanism is that of singlet–singlet energy transfer between large aromatic molecules, a process used by nature in the antenna part of the photosynthetic apparatus [1]. Finally, it is worth mentioning that new theories, such as the one named NSET [19, 20], have been developed in case of energy transfer processes involving metal nanoparticles that could be of interest, in this context, for silica nanoparticles having a metal core.

3 The Power of Energy Transfer Processes Professors V. Balzani and F. Scandola, some years ago, in a pioneering work in the field of supramolecular photochemistry [1], had already described four different kinds of photochemical molecular devices (PMDs) based on electronic energy transfer processes and sorted them according to their main functions. We think that it can be very important to recall here these concepts since they can be very helpful to foresee new applications for silica nanoparticles. The first group includes the antenna systems, which present an enhanced light sensitivity obtained by an increase in the overall cross-section for light absorption. In typical supramolecular devices of this kind, the energy has to be conveyed to a single component that acts as the interface towards their applications. In simple (containing only one kind of dye) DDSNs used as fluorescent labels, the energy transfer process is not required, since the increase in the cross-section is simply originated by the increase in the number of dyes that are directly responsible for the fluorescence of the nanoparticle. In such systems, the occurrence of energy transfer processes could instead lead to undesired self-quenching phenomena, and it should be avoided. To this purpose the doping of the nanoparticles must be reduced, increasing in this way the average interchromophoric distance and, in agreement with Eq. (3), decreasing the rate of the energy transfer process. This decrease of the loading,


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however, has also the counter effect to lessen the cross-section for light absorption and, as a consequence, the brightness of the nanoparticle. Recently, Geddes and Lakowicz [21–23] proposed a new strategy aimed to increase the energy transfer rate based on the effect of plasmonic nanostructures. When the distance from a nanostructured metal (usually silver) is properly set, the radiative deactivation rate of the excited state of a dye becomes, in fact, higher. As a consequence of the enhancement of the fluorescence apparent quantum yield, the energy transfer process is expected to become more efficient (see Eq. (6)). However, when more complex systems are needed, like ‘onion-like’, multilayered structures, in which all the energy gathered in a single shell has to be conveyed to an adjacent one, the role of homo-energy transfer becomes essential. In this case, the energy absorbed by a dye should hop until it reaches the boundary of the neighbouring shell, and from here it should hop to the lower energy state of the dye that is entrapped in this last shell. To reach this goal, it is important to obtain a good balance between the inconvenient self-quenching, typical when many organic dyes of the same kind are very close to one another (usually a short distance phenomenon), and the homo-energy transfer, that should be, in turn, as efficient as possible. In order to achieve such a result, it is of great importance to know how the different dyes are located and distributed inside the nanoparticle, which we have recently started to study [9, 24]. Therefore, the importance of the antenna effect in DDSNs and DCSNs designed to work as sensors is crucial to obtain the so-called signal amplification effect. This kind of effect, which leads to a large increase in the sensitivity, and, as a consequence, to lower detection limits, can be obtained if the complexation of a single analyte is able to change the fluorescence properties of several neighbouring dyes, and this is not possible following the usual single receptor–single luminophore approach [8]. In multichromophoric structures, such as DDSNs and DCSNs, many different chemosensor units are present and, if no energy transfer can occur, they behave as they were ‘electronically isolated’, i.e. every dye changes its luminescence only when the connected receptor binds the target analyte [25]. On the contrary, when energy transfer is efficient, the electronic energy absorbed by each chromophore inside the matrix migrates towards an ‘electronic sink’, i.e. the lowest energy state, by a process that results in the amplified luminescence quenching [12] or enhancement [13]. This case can be schematized as thousands of bulbs electrically wired to thousands of switches connected to each other: acting on a single switch one changes the status of all the bulbs simultaneously. A second major effect is spectral sensitization, which is very important when the light absorption properties of a potentially photoactive (generally, luminescent) species do not permit efficient excitation in the desired wavelength range. This kind of phenomenon is crucial for many applications in different fields, such as, for example, the spectral sensitization of semiconductor electrodes in solar energy conversion. Although the use of silica nanoparticles is starting to attract the attention of researchers also working in the field of energy, the applications that more frequently takes profit of this kind of function are the ones requiring the preparation of systems possessing remarkable Stokes shift, which typically allows a dramatic

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increase in the signal to noise ratio in photoluminescence spectroscopy measurements. To obtain this goal, two different dyes, a donor and an acceptor, chosen in order to optimize the efficiency of energy transfer processes in agreement with Eq. (3), are inserted in the silica matrix in a controlled way to obtain suitable reciprocal distances. The light absorbed by the donor (usually a coloured species absorbing visible radiations) populates one of its excited states that is at a higher energy with respect to the emitting state of the acceptor, which can therefore be efficiently populated via energy transfer. This leads to the emission of the acceptor via excitation of the donor, yielding a significant bathochromic luminescent shift. Another kind of sensitization is based on the same process but it takes place between donors and acceptors that are distant from one another, and it is called remote photosensitization. In order to obtain energy migration over long distances, many subsequent energy transfer processes are needed, characterized by the exoergonicity of the first and last ones and the isoergonicity of the intermediate ones. This phenomenon is typical of components connected by long rigid bridges and in this case the energy transfer is vectorial, but it can also take place in non-rigid systems like photoactive soluble polymers or in solid systems (including silica nanoparticles) where the components are randomly distributed. In these last cases the process is obviously non-vectorial but it can be anyway extremely efficient (close to 1). The last kinds of systems carrying out a function based on energy transfer are the ones performing light energy upconversion, that is to say showing anti-Stokes luminescence. In this case the emitting moiety, the acceptor, presents the lowest excited state that is populated, in the first stage, via energy transfer from one donor and, before undergoing deactivation, is further raised to an upper excited level by a second energy transfer process. The two donors can be two different species or the same one transferring sequentially the two exciting photons. To obtain this very particular and precious function, many energetic, kinetic and spatial requirements have to be fulfilled and this is usually not easy to obtain. Silica nanoparticles can be, again, a very useful means to obtain such a result, thanks to their possible multichromophoric nature, great versatility and possibility to choose and tune inter-dye distances, although at the moment not many research efforts have been devoted in this direction.

4 The Synthesis of Luminescent Silica Nanoparticles Silica nanoparticles are particularly interesting nanomaterials also from a synthetic point of view: the great flexibility of their synthetic procedures allows a so-called modular approach that is an extremely valuable feature to obtain multifunctional materials. The most diffused and important synthetic strategies to prepare silica nanoparticles [26] are the Stöber [27] method and the reverse microemulsion one [28–30]. The chemical process at the basis of the silica nanoparticle formation is the same for both of them: the controlled hydrolysis of tetraethoxysilane (TEOS)


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molecules and their ammonia catalysed condensation. The two methods differ in the reaction environment of hydrolysis and condensation: in the first one it is an ethanol/water/ammonia solution, while in the second one it is a stable and macroscopically isotropic dispersion of a surface-active agent and water in an apolar solvent. In the microemulsion, the hydrolysis is confined inside the aqueous nuclei where TEOS and organic precursors diffuse during the hydrolysis reaction. In both cases it is possible to obtain nanoparticles with a diameter ranging from 15 to 200 nm and highly monodispersed, providing a careful control of the reaction conditions. The Stöber method allows the continuous and easy control of the nanoparticle dimension by a suitable choice of the concentrations and ratios of the components of the reaction mixture (TEOS, water and ammonia). In the microemulsion procedure, on the other hand, the main discriminating parameters are the kind of surfactant and the amount of water. In order to obtain a fluorescent material, in both synthetic strategies, it is necessary to add in the reaction batch a certain amount (usually ≤1% vs. moles of TEOS) of the needed fluorophore and the choice between the two methods depends on the solubility of the fluorophore itself; the microemulsion one, for example, is preferred when the emitting species is very soluble in water. The dye can be added (a space was laking) during the growth of the nanoparticles either already derivatized with an alkoxysilane group or without the introduction of any anchoring group to be simply physically entrapped in the structure. The latter choice allows its covalent linking on the surface or inside the particles. Proper functionalization of fluorophores is very often not trivial but, most times, it is an essential step for the preparation of dye-doped silica nanoparticles unaffected by dye leaching problems (in particular when a high level of doping is required). Since this is a crucial point, many synthetic efforts have been devoted in these last years to develop synthetic strategies able to yield pure luminescent silanes, suitable for both organic chromophores and metal complexes. Their pureness is also important since it is needed to avoid the inclusion of unreacted silanes that could interfere with the photophysical properties of these systems. The main synthetic procedures to obtain alkoxy-functionalized fluorophores include the hydrosilylation reaction and the use of commercial alkoxysilanes bearing useful reactive functional groups. Among the possible reagents we often prefer the triethoxy(3-isocyanatopropyl)silane since its reaction with fluorophore derivatives bearing an amine group is fast and quantitative. Furthermore, the formation of the ureidic group in the adduct often increases the solubility of lipophilic organic dyes in polar solvents (ethanol, water) to concentrations useful to the nanoparticle synthesis. During the preparation of these nanomaterials the condensation of trialkoxysilane derivatives is almost quantitative, but sometimes a purification step can be needed. There are a few efficient methods to separate the nanoparticles from the unreacted silane derivatives: centrifugation, ultrafiltration and dialysis are the most interesting ones and the choice among them depends on the properties of the sample. Centrifugation is generally proper for big nanoparticles (with a diameter of 100–200 nm and more) [31], or for those tightly stabilized by an efficient surface

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passivation able to prevent their aggregation via irreversible condensation effects between the outer siloxanic groups. Ultrafiltration is a particular kind of filtration originally conceived for protein purification. With this method the nanoparticle solution is forced under low pressure through a membrane presenting a controlled porosity that allows the separation of the unreacted molecules or oligomers from the nanoparticles. Together with dialysis this is the best technique in order to purify solutions of small nanoparticles, preserving their monodispersity and, when needed, changing the alcoholic environment of the synthesis in an aqueous one. We also showed [24, 32] that FlFFF (Flow Field-Flow Fractionation) is another powerful technique to size, sort and micropurify fluorescent colloids of nanometric dimensions, able to reduce their dimensional polydispersity and to remove the unreacted fluorophores. In order to study and obtain energy transfer processes in silica nanoparticles a few different strategies can be followed: (i) inserting a fluorophore in different concentrations and therefore at different intermolecular distances; (ii) inserting different dyes in the same nanoparticles; (iii) creating multishell particles, each differently doped; (iv) or introducing a metal core covered with one or more dye-doped silica shells. A common feature of all the different nanoparticles described so far is that their solubility, stability and reactivity or, at least, affinity towards other species depend on the nature of their surface. Therefore, the great importance to optimize chemical strategies to modify and decorate silica nanoparticle surfaces, also from an applicative point of view, is immediately evident. Some years ago, we developed a synthetic strategy aimed to covalently bind fluorescent silanes on the surface of small (15–20 nm of diameter) silica nanoparticles [33]. In this procedure, the silanes are refluxed for some hours together with a nanoparticle suspension in the presence of acetic acid, yielding nanoparticles presenting a thin layer of organic material on the surface. This strategy can be adapted to introduce almost any species and even more than one species at once, and their original solubility or photophysical properties can be tuned or preserved unchanged with a rational control of their amount and/or ratio. The uniformity and the stability of the surface covering can be controlled by changing the nature of the anchoring function. Triethoxysilane groups, in fact, are suitable for surface modifications but they do not allow the complete saturation of all the Si-OH sites at the surface that could therefore still present an undesirable reactivity. For this reason, the successive passivation with end-capping reactants such as (CH3 )3 SiCl could be sometimes necessary. As an alternative, monoethoxysilanes could be used instead of triethoxy ones, to obtain a more uniform covering, but this causes also a decrease in the binding efficiency.

5 Photophysical Properties of Luminescent Silica Nanoparticles As mentioned before, DDSNs and DCSNs are very versatile and synthetically easy platforms for the organization of a large number of photophysically active units.


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This makes silica nanoparticles very promising nanomaterials for the development of labels and sensors. They usually present quite complex photophysical characteristics, due to their multichromophoric nature. Dyes are often organized in high-density patterns on the surface of the nanoparticles, or inside them, or even in the different shells of multilayered structures. The fluorophores obviously experience different conditions inside the silica matrix and this induces photophysical processes that undergo delicate balances and that determine and tune the photophysical behaviour of the whole structure. In order to design and synthesize SNs presenting desired properties and functions, we think that it is fundamental to understand the behaviour of the dyes in these nanometric systems and much of the research efforts of our group have been devoted, in these last years, to the study of the position, communication and processes occurring between the fluorescent units. The synthetic strategies and the characteristics of these systems strongly depend on the kind of the SNs, and the choice between DDSNs and DCSNs is related to the specific application or function aim of the research. Therefore, we discuss hereafter some examples of both groups underlining their differences and potentialities. The first example of dye-doped silica nanoparticles dates back to 1992 [34, 35], and it was described by van Blaaderen. He used fluorescein isothiocyanate to obtain particles with a diameter of hundreds of nanometres. An extensive work has been made since then on this topic: many different fluorophores have been incorporated in nanoparticles ranging from 15 to 500 nm with excellent monodispersity. The occurrence of energy transfer processes between dye molecules in DDSN was demonstrated for the first time by our research group. Using fluorescein triethoxysilane derivative 1 (Fig. 1) we obtained DDSNs with a diameter of 20 nm [24], a size suitable for bio-analytical purposes, and reasonable for a colloidal system to foresee that each excited fluorophore could interact with a large fraction of the units bounded to the same nanoparticle. Through a very efficient microscale FlFFF-based method that allows the size sorting of the nanoparticles, we were able to obtain a batch of nanoparticle solutions characterized by slightly different diameters. In all the samples, the fluorophore quantum yield was independent of the nanoparticle dimension and it was about 50% of that of the free fluorophore in the same conditions. Moreover, despite the low rotational mobility of the fluorescein derivatives linked to the silica matrix, we found that nanoparticle fluorescence was almost completely depolarized (P< 0.02). These results, together with the existence of a single exponential component in the fluorescence intensity decay, suggested a nonhomogeneous density of the dye molecules in the silica matrix. In particular, the presence of more dense regions where self-quenching is the prevailing process could explain the quantum yield decrease, while less dense regions, where fluorophores are however close enough to take part in energy transfer processes, are responsible for residual depolarized emission. In order to explain such a behaviour we tried to elucidate the various steps of dye insertion during the particle formation, studying stepwise the process of formation of silica nanoparticles via the Stöber–van Blaaderen method and using the

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Fig. 1 Chemical structure of some of the molecules which have been used for the preparation of luminescent silica nanoparticles

newly synthesized pyrene triethoxysilane derivative 2 (Fig. 1) as probe [9]. Pyrene is an excellent choice to follow the formation process of the nanoparticles and to explore their inner structure. The photophysical properties of pyrene are, in fact, deeply affected by the local environment and concentration, in such a way that it has been extensively used for characterizing nanostructures as micellar systems. Its fluorescence quantum yield increases tenfold when segregated from oxygen, as happens inside the silica matrix, and this offers the possibility to evaluate the fraction of fluorophores inside the silica nanoparticles during the synthesis, following their growth in real time. Moreover, interactions between very close pyrene moieties upon excitation do not cause self-quenching but excimer formation. As a consequence, the evaluation of the ratio of the excimer and the monomer emission bands can give an indirect information on the local concentration of the dye inside the silica matrix. In our experiments, we correlated the absorption and fluorescence emission spectra of the DDSNs with the profile of the hydrodynamic volume growth obtained with DLS (dynamic light scattering) technique for two samples characterized by different amounts of pyrene triethoxysilane derivative (with respect to tetraethoxysilane). In particular, from the comparison between the excimeric and monomeric contributions, we obtained information about the local concentration of the dye during the different stages of the nanoparticle growth, evidencing the compartmentalization of


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Fig. 2 Schematization of the spontaneous organization of 2 in doped silica nanoparticles: two different emission bands corresponding to the monomeric and the excimeric form of pyrene can be detected (spectrum on the right)

the fluorophore molecules and the self-organized core–shell structure of the system (Fig. 2). Besides, we noticed that, while the dye inclusion in the nanoparticles was almost quantitative for both concentrations, the doping degree influenced both the final diameter and the initial speed of the first step of nanoparticle growth. Larger nanoparticles were in fact obtained in the presence of a higher concentration of the dye. The results obtained in the case of 2 suggest that also in the case of fluorescein derivative 1, a core–shell structure with preferential accumulation of the dye in the nucleus of the NPs is spontaneously formed. As schematized in Fig. 3 short-range interactions in the central highly doped region lead to quenching of the fluorescence, while migration of the excitation energy occurs in the external layer where interfluorophore average distance is larger. The strong dependence of the photophysical properties on the substructure of the nanoparticles, on the other hand, has also been demonstrated by Wiesner and co-workers with silica nanoparticles synthesized via the Stöber method and doped

Fig. 3 Schematization of the spontaneous organization of 1 in doped silica nanoparticles: homo energy transfer takes place in the outer shell while the molecules densely packed in the core are quenched

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with tetramethylrhodamine [36, 37]. Using exactly same amounts of reagents, but slightly different synthetic procedures, it is possible to obtain nanoparticles with a different inner architecture, from compact to expanded core/shell structures or even homogeneous systems. It has been shown that these diversities lead to a different photophysical behaviour of the dye. In particular, by means of fluorescence correlation spectroscopy (FCS) the brightness of the nanoparticles was compared to one of a single dye molecule revealing that core–shell nanoparticles present a peculiarly high absorption and emission efficiency. Anisotropy measurements evidenced, in this case, that there was no energy transfer between the dyes in the nanoparticles, but they absorb and emit photons independently from each other. To induce useful energy transfer processes a good strategy is to dope silica nanoparticles with two (or more) properly chosen different fluorophores. This is also a method to obtain functional multicolour systems to be used in multiplexed bioassays. In this context, anyway, the specific mechanism of energy transfer that is expected to operate must be considered. In the case considered by Förster of dipole–dipole interactions between the donor and the acceptor, typically important for organic dyes, energy transfer can take place with 50% efficiency even through distances as long as 5–6 nm. On the contrary, when orbital overlap is required, energy transfer occurs only between species in close proximity as in the case of Dexter mechanism. Average intermolecular distances suitable for this latter kind of energy transfer can be hardly achieved in doped silica nanoparticles. This is the reason why in SNs obtained by microemulsion processes, energy transfer between the two co-included metal complexes, Ru [tris (2,2 -bipyridyl) dichlororuthenium (II) hexahydrate] and Os [tris(2,2 -bipyridyl) osmium (II) bis (hexafluorophosphate)], was not observed [38]. The luminescence spectra recorded were, in fact, the simple sum of those of the two luminescent components and the emission pattern could be controlled linearly by varying the molar ratio of the two dyes during the synthesis. Tan and co-workers, anyway, demonstrated that this possibility of tuning the spectral signature of the NPs may find important analytical applications. If each individual batch of nanoparticles is coated with a different analyte ligand, it is possible to record a complex matrix composed of many signals, using a single excitation wavelength [39]. A global analysis of these data can give qualitative and quantitative information on the target analytes present in the sample. This concept can be further extended by increasing the number of luminescent species inside each nanoparticle and taking advantage of energy transfer processes. Replacing the ruthenium and osmium complexes with organic dyes, in fact, energy transfer can occur via dipole–dipole interactions and the relatively long distance between the donor and the acceptor becomes less critical. As discussed above, to maximize the efficiency of RET, the emission of each dye should largely overlap the absorption of the neighbouring one emitting at lower energy. One example is given by tri-luminophore-doped silica nanoparticles containing FITC-R6G-ROX in different ratios. These systems were prepared following the classic Stöber method that is to say co-condensing silanized dyes during the nanoparticle growth. In this case FITC acts as a donor for R6G which in turn acts


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as a donor for the final acceptor ROX. Again, by varying the molar ratio between the three dyes, the emission patterns can be even more distinct than in the previously cited example by the same authors, with a very large Stokes shift, that helps in overcoming problems arising from autofluorescence (common phenomenon in biological matrixes), scattering of the excitation light and Raman/Rayleigh bands. The systems described up to this point were designed with the aim of controlling, improving or combining the properties of molecular or supramolecular luminophores mostly in view of a possible application as nanosized luminescent labels. In this perspective, the inclusion of the dyes in the silica matrix offers the relevant advantage of insulating them from the external environment, preventing their interaction with species that could cause luminescence quenching or even chemical or photochemical degradation. In the next part of this paragraph we will show that, despite such a shielding effect, organic molecules included in silica nanoparticles are still accessible to small species such as metal ions. This permeability makes SNs a very versatile platform for the design of luminescent nanosensors. Energy transfer is in this context extremely important since it may allow the different molecular components to ‘communicate’ the transforming of the individual responses to the target recognition into a collective one, an effect that translated into analytical terms means higher sensitivity and lower detection limits. In order to demonstrate the possibility of amplifying the response of a fluorescence chemosensor by inclusion in SNPs, we synthesized a proof of principle system based on the dansylated 3-[2-(2-Aminoethylamino)ethylamino]propyltrimethoxysilane commercial receptor 3 (Fig. 1) [40]. The nanoparticles were synthesized with the Stöber method maximizing the density of the fluorophores with the purpose of allowing the occurrence of multicomponent cooperative photophysical processes. Interestingly, the addition of copper, cobalt and nickel ions induced a strong quenching of the fluorescence intensity even at nanomolar concentrations (Fig. 4).

Fig. 4 Changes in the fluorescence of silica nanoparticles doped with 3 upon addition of metal ions. The equivalents are referred to the concentration of 3 molecules considering a one to one stoichiometry for the complexation

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Fig. 5 Schematization of the energy transfer processes occurring in SNPs doped with chemosensor 3. Complexation of a single Cu2+ ion causes the average quenching of 13 fluorophores (only 5 are reported in the picture for simplicity)

The ability of Cu2+ , Ni2+ and Co2+ to quench the fluorescence of the dansyl fluorophore has been reported for polyaminic dansylated systems and has been explained on the basis of energy transfer processes involving the dansyl excited state and the metal ion (Fig. 5). The results obtained with our system, however, suggest that each copper ion, the species having the highest affinity, could quench up to 13 dansyl units, leading to strong signal amplification. This fluorescence response to the complexation showed by the nanoparticles is in our opinion extremely interesting since it proves that they are porous enough to let in small cations and that the proximity of the chemosensor units allows the communication of each receptor moiety with all the neighbouring fluorophores. This was not our first research work based on the covalent binding of the dansyl unit to SNs, since our earliest paper in the field dealt with silica nanoparticles coated with 4 (Fig. 1) [33]. We have studied, in fact, a few DCSN systems before starting to focus our attention on DDSNs and usually aiming to prepare efficient nanosensors. In this first work, we capped preformed silica nanoparticles (Ludox AS30) with dansylated 3aminopropyltrimethoxysilane units 4. Dansyl is a widely used fluorophore, in which the lower excited state has a charge transfer character involving the promotion of a lone pair electron of the amino group into a π antibonding orbital of the naphthalene ring. Since this process is strongly affected by the protonation of the amino group itself, it is not unexpected that 4 and 4H+ behave as different fluorophores that can be selectively excited and detected (Fig. 6). We used the nanoparticle surfaces to densely arrange the fluorophores, to produce a polydansylated model system in which the ratio of two different dyes (4 and 4H+ ) could be modulated through the degree of protonation. This system showed not only a dramatic response to protonation due to charge transfer interactions between fluorophores but also a strongly solvent-dependent behaviour. As schematized in Fig. 6, in a quite polar environment (chloroform/acetonitrile 5:1) as soon as 4 protonation began, an amplification quenching process of the dansyl units was evidenced together with a slightly delayed switching on of the fluorescence of the 4H+ units. A different situation was found in a less polar environment (pure chloroform), where the dansyl quenching


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Fig. 6 Left: changes in the photophysical properties of 4 upon protonation. Right: schematization of the processes occurring in silica nanoparticles coated with 4 in the case of the protonation of a high fraction of the dansyl units in CH3 CN (top) and CHCl3 (bottom)

amplification process was negligible. In this context, on the contrary, the dansylated nanoparticles behaved like an antenna system in which upon excitation of the 4H+ units the energy was efficiently transferred to the luminescent 4 moieties. Following this synthetic strategy, we have recently demonstrated the possibility to take advantage of the spatial organization and electronic communication between chromophoric units on the surface of silica nanoparticles for the development of a self-organized Zn(II) fluorescent chemosensor [41]. We used a triethoxysilane derivative of TSQ (6-methoxy-8-p-toluenesulfonamide), 5 in Fig. 1, to realize a multichromophoric network on the surface of preformed silica nanoparticles. TSQ is a widely used fluorescent chemosensor able to bind Zn(II) ions with a good selectivity. It is characterized by an off–on response due to an internal charge transfer (ICT) in the Zn(II)TSQ and Zn(II)(TSQ)2 complexes (Fig. 7). In our system, the off–on

Fig. 7 Left: changes in the photophysical properties of 5 upon complexation. Right: schematization of the processes occurring in silica nanoparticles coated with 5 at low zinc concentration

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fluorescence signal was amplified by the energy transfer process from the uncomplexed TSQ units to the luminescent Zn(II) complexes (Fig. 7). In a low Zn(II) concentration regime, this self-organized chemosensors showed a 50% increase in the response with respect to the reference system TSQ in the same conditions. The relatively low level of amplification can be explained by taking into account the Förster radius for the energy transfer (Eq. 4), which for the nanoparticle-based TSQ system can be estimated to be shorter than 0.5 nm, a value compatible with the distance between small trialkoxysilane derivatives grafted on the surface of a silica nanoparticle. Even if the sensitivity gain produced by the sole energy transfer process is quite limited, this system is, to our knowledge, the first example of an amplification effect in an off–on material of this kind. This phenomenon together with the enhanced affinity towards the substrate (the association constant increases of almost four orders of magnitude in the NPs), induced by the self-organized network on the surface of the nanoparticles, yields a great increase in the sensitivity of the system (Fig. 8) and suggest interesting hints for the development of new fluorescent chemosensors. Multichromophoric systems obtained by the organization of active units on the surface of preformed SNs are in this context more promising than those resulting from silica doping since they allow a higher local density of molecules and hence a stronger electronic communication. When the same TSQ derivative is in fact included in the silica matrix its fluorescence is still sensitive to the presence of zinc ions but, as Mancin and co-workers reported, no amplification effect could be observed because of the excessive distance between the receptor molecules [42]. Fig. 8 Fluorescence spectra of a solution of nanoparticles coated with 5 before (a) and after (b) the addition of zinc (1 × 10–6 M). The same addition to a solution of 5 with the same concentration of dye causes very small changes (from (c) to (d))

Although the occurrence of energy transfer processes is definitely important to achieve amplification effects, insulation of dye molecules in order to avoid it may become fundamental in the design of ratiometric sensors. One of the dyes present in these devices in fact is used as a reference and its fluorescence signal has to be independent from the state of the luminescent component responsible for the sensing. A possible design strategy for preventing energy transfer processes between


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different dyes was proposed independently by Wiesner and Mancin and it is based on multilayered structures where the reference dye is localized in the core and the sensitive agent dispersed in the external shell of the nanoparticles [4, 10]. Recently, a new method has been proposed for increasing the efficiency of energy transfer exploiting the effect of plasmonic nanostructures [21–23]. Silica in this system is often used as an insulating layer to avoid direct ‘contact’ of the fluorophores with the metal (usually silver) which would result in excitation energy dissipation. These hybrid systems are a clear example of how accurately nanostructures must be designed in order to funnel the excitation energy through the desired path. The efficiency of the energy transfer between the fluorophores, in fact, can be increased by the presence of the metal nanostructure if this is localized at an optimal distance, but if it is too close direct energy transfer to the metal takes place with a concomitant quenching of the fluorescence. On the contrary, when the distance from the metal is too large its presence becomes ineffective having no influence on the excited state deactivation.

6 Conclusions All the systems discussed above demonstrate how the careful design of energy transfer processes in silica nanoparticles can induce very sophisticated and controllable functions. Since the synthesis of SNs is still in its ‘infancy’, many new and powerful possibilities are constantly explored and introduced. The first important aim is to gain a very precise control of the density of the dyes inside (or at the surface of) the nanoparticle, and even if many progresses on the synthetic protocols and on the characterization of SNs have already been reached, in this context, a really total control has still to be achieved. Furthermore, the construction of more sophisticated materials, such as multilayered NPs, is opening up new challenges on the design of this kind of nanostructures, expanding to an extent not even thought possible till a few years ago, the possibility to induce and tune all desired photophysical processes. If researchers will succeed in this goal, new ambitious applications will be possible in many essential fields, such as medical diagnostics, molecular biology and solar energy conversion. Acknowledgment The financial support from MIUR (FIRB 2003-2004 LATEMAR, http://www.latemar.polito.it) and Fondazione del Monte is gratefully acknowledged.

References 1. V. Balzani and F. Scandola, Supramolecular Photochemistry (Ellis Horwood Ltd, Avon U.K., 1991). 2. L. Prodi, G. Battistini, L. S. Dolci, M. Montalti and N. Zaccheroni, Luminescence of Gold Nanoparticles. In Frontiers in Surface Nanophotonics: Principles and Applications. Ed. D. L. Endrews and Z. Gaburro (Springer, New York, 2007), pp. 99–120.

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Spectroscopic Characterization of Plasma – Chemically Functionalized and Fluorophore-Labeled Polymer Surfaces Katrin Hoffmann, Renate Mix, Joerg F. Friedrich, and Ute Resch-Genger

Abstract The potential of spectrofluorometry and fluorescence microscopy for the characterization and quantification of different functionalities like OH and NH2 groups at plasma-chemically modified polymer surfaces is assessed using traditional reactive dyes such as dansyl derivatives and a sophisticated VIS-excitable chromogenic and fluorogenic pyrylium label showing binding-induced spectral and intensity changes in absorption and emission. Aiming at an improved fluorometric surface analysis, based upon these measurements, several sources of uncertainty inherent to fluorescence measurements are illustrated ranging from environment-dependent dye absorption and emission features over spectral correction and nonspecific adsorption to the critical influence of label choice on the measured background. Solutions to these drawbacks are given thereby underlining the potential of fluorometry for surface analysis.

1 Introduction Many advances in the material, life, and analytical sciences are closely linked to surface chemistry. For example, analytical techniques such as the microarray technology, the majority of bead-based techniques, the miniaturization of tools for chemical analysis like the design of lab-on-a-chip systems, the development of new chemical, hybrid, and biosensors, and the design of fluorescent nano and microparticles for labeling and sensing applications require control of the wetting, adsorption, and/or binding properties of support or building materials [1–7]. Accordingly, engineering of surfaces on a molecular level is of ever-increasing importance and thus, the development of evaluated strategies toward a defined surface functionalization with different chemical groups [8–12]. For this purpose, not only chemical and U. Resch-Genger (B) Working Group Fluorescence Spectroscopy, Department I, BAM Federal Institute for Materials Research and Testing, Richard-Willstaetter-Str. 11, D-12489 Berlin, Germany e-mail: [email protected] C.D. Geddes (ed.), Reviews in Fluorescence 2008, Reviews in Fluorescence 2008, DOI 10.1007/978-1-4419-1260-2_6, C Springer Science+Business Media, LLC 2010

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photochemical approaches are used, but also plasma-chemical treatment techniques, e.g., for coating, finishing, or etching [13–17]. Similarly crucial in this context are straightforward, robust, cost-efficient, and evaluated analytical tools for the reliable characterization of the chemical nature and surface concentration of functional groups. Especially the accurate quantification of surface functionalities presents a major challenge. Traditional surface characterization techniques include Fourier transform infrared spectroscopy in the attenuated total reflection mode (ATR-FTIR), contact angle analysis, atomic force microscopy (AFM), X-ray photoelectron spectroscopy (XPS; without and in combination with chemical derivatization using XPS tags or labels), and lately also time-of-flight secondary mass spectrometry (TOF-SIMS) [18, 19]. Alternatives are the use of simple and sensitive fluorometric methods in conjunction with labeling techniques [19–23], and for selected applications aiming, e.g., at the calibration of other (relative) techniques, also the application of radioactive labels in conjunction with a scintillation analyzer [24]. In addition, also surface plasmon resonance (SPR) has been employed occasionally [25]. XPS, the most commonly used surface-sensitive technique, is costly and often fails at low concentration. In the case of ATR-FTIR, the comparatively large penetration depth of this technique reaching from 1.6 to 2.5 μm (depending on the used ATR crystal) can hamper the distinction between the surface groups and the bulk material. Label-free TOF-SIMS has proven to be a powerful tool to analyze complex surfaces like self-assembled monolayers, polymers, and protein films due to its high surface sensitivity, high mass resolution, and molecular specificity. However, it is a rather sophisticated and expensive technique to which many laboratories do not have access. Contact angle measurements provide a straightforward measure for surface wettability, yet this method, is not suited for the typically desired accurate determination of the density or the concentration of surface functionalities. SPR techniques are commonly limited to the study of surface-dependent binding or adsorption processes. The use of radioactive labels that require special safety precautions, underlies stringent regulations, and creates hazardous wastes, is increasingly unpopular. This situation – in conjunction with the huge toolbox of reactive fluorescent dyes for the derivatization of a broad variety of different functional groups [26] and the comparatively inexpensive and easy-to-use instrumentation – renders the application of fluorometric methods very attractive. Even though the monitoring and characterization of surface functionalities with the aid of surface-linked fluorophores have been frequently reported [27, 28], the potential of fluorometry to accurately identify and quantify reactive groups and intermediates at surfaces is still debated [27–30]. Critical factors determining the reliability and quantification potential of fluorometry are here, e.g., the sensitivity of the spectroscopic features of most chromophores to dye microenvironment, the lack of analytically exploitable binding-induced changes in the absorption and fluorescence properties of most labels, nonspecific dye adsorption and binding that both result in an enhanced background fluorescence, and fluorescence quenching phenomena [31, 32].

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This encouraged us to study different strategies for the labeling of surface functionalities on solid supports employing various types of fluorophores differing in the electronic nature of the emissive state and the occurrence of binding-induced spectroscopic changes. In a second step, the potential of fluorescence techniques, here conventional spectrofluorometry examining the features of the bulk films and spatially resolving confocal laser scanning fluorescence microscopy, for the characterization of surface functionalities is discussed. Aiming at an improved fluorometric surface analysis, potential sources of uncertainty of fluorescence measurements are illustrated and solutions to these drawbacks are given. For this purpose, we voluntarily chose a polymeric system, here polypropylene, that showed a certain porosity and a complex surface chemistry due to plasma-chemical modification.

2 Surface Modification 2.1 Surface Functionalization of Polypropylene A wide variety of functional groups can be elegantly generated at chemically inert polymer surfaces using plasma-chemical processes thus forming anchoring sites for chemical grafting of different labels or other functional molecules. For our studies on the suitability of fluorescence techniques for the characterization of polypropylene (PP) films (Goodfellow, UK; thickness of 100 μm), two different routes of surface modification have been used: (i) an oxygen plasma treatment, followed by a wet-chemical reduction process [33], that yields stable OH-terminated polymers with surface densities of about 10–14 OH/100 carbon atoms on the PP surface [34] and (ii) the plasma-initiated polymerization of monomers carrying functional groups in a pulsed plasma. The pulsed plasma regime results in polymer films with a high degree of retention of the functional groups. With this strategy, NH2 -terminated materials could be generated by deposition of allylamine layers onto the surface of PP [35].

2.2 Determination of Surface Functionalities with XPS XPS characterization of surfaces is typically performed in conjunction with the previous derivatization of surface functionality with substances containing heteroatoms (e.g., chlorine, bromine, fluorine, sulfur, nitrogen) which are absent in the untreated sample. In these derivatization reagents or so-called XPS tags a number of the heteroatoms as high as possible is desired. Subsequent XPS measurements of the incorporated heteroatoms provide information about the quantity of reacted functional groups per 100◦ C atoms in a 5–7 nm thick surface layer. Prerequisite for correct results are accordingly quantitative derivatization reactions (degree of surface group conversion of at least 90%) that should occur selectively with a known stoichiometry. These reactions are preferably performed in the gas phase, because

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additional uncertainties can arise from the dissolution of oligomeric or polymeric species at the surface in contact with solvents. Furthermore, the derivatized species have to be stable under XPS conditions. In the following, an example for the derivatization of plasma-chemically generated surface hydroxyl groups with trifluoroacetic anhydride (TFAA), see Fig. 1, is given [12]. For the direct comparison of XPS and fluorescence measurements, fluorescent reporters containing heteroatoms like sulfur atoms such as dansyl derivatives (SO2 moiety) or nitrogen atoms (from, e.g., isocyanate groups) were used to circumvent uncertainties due to different reactivities and molecular sizes of XPS and fluorescent labels [30, 32].

Fig. 1 Derivatization of surface OH-functions with the XPS tag trifluoroacetic anhydride (TFAA)

2.3 Fluorophore Labeling of PP Surfaces Functional groups like amino, hydroxyl, and thiol functionalities, e.g., at biomacromolecules, are often monitored with fluorescence techniques after subsequent derivatization with reactive chromophores [36–38]. Traditional reactive dyes are chromophores like fluoresceins and rhodamines as well as also cyanines that all reveal slightly structured, comparatively narrow, absorption and emission bands and a small Stokes shift which is almost matrix polarity insensitive [39]. A disadvantage of these labels is the strong spectral overlap between absorption and emission, which hampers the ease of separation of excitation from emission, reduces the efficiency of collecting the emission signal and favors concentration-dependent emission spectra due to reabsorption. Also, as compared to other dyes, the pronounced tendency toward the formation of nonfluorescent aggregates can be critical. An example for a different class of fluorescent labels presents dansyl chromophores. These fluorophores show broader and structureless absorption and emission bands and a larger matrix polarity-dependent Stokes shift indicative of intramolecular charge transfer (CT) transitions and CT states [40]. Disadvantageous for such CT-operated dyes, which are less prone to aggregation-induced self-quenching, can be the pronounced sensitivity of their spectroscopic features to the polarity and often also to the proticity of dye environment. Moreover, all these traditional fluorescent labels were developed and evaluated for the reaction with freely accessible functionalities of molecular systems such as biomacromolecules in solution and not for the detection of surface functionalities on solid supports where an influence of the surface on the reaction is likely. This can, e.g., affect the reaction kinetics and the reaction


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yield in the case of labels bearing sterically demanding substituents (e.g., sterical hindrance). In our group, various synthetic strategies, partly adopted from the wellinvestigated polyurethane chemistry, have been tested for fluorophore labeling of plasma-chemically hydroxyl-modified surface functionalities, as well as for the introduction of spacer molecules like toluylene-2,4-diisocyanate (TDI) or methylene-di-p-phenylene diisocyanate (MDI) (see Fig. 2). Using these concepts, the covalent and surface-separated attachment of amino-functionalized fluorophores (e.g., dansyl cadaverine, DNS-Ca, and dansylhydrazine, DNS-Hy) and various commercial reactive fluorophores (isothiocyanates, succinimidyl esters, sulfonyl chlorides) to plasma-chemically generated OH-functions was realized. The experimental conditions for the chemical modification of the surfaces are described elsewhere (e.g., [12, 32]). To modify plasma-chemically amino-terminated PP surfaces directly or via a glutaraldehyde spacer molecule, the well-known reactions of reactive dyes forming sulfonamides, thioureas (see Fig. 3a), or azomethines (see Fig. 3b) were used [41].

Fig. 2 Labeling of OH-functionalized and subsequently diisocyanate-modified PP surfaces with amino-functionalized fluorophores (A), conversion of diisocyanate functions to aminofunctionalized surfaces by hydrolysis (B), and different routes for surface labeling with common reactive fluorophores (isothiocyanates, succinimidyl esters, sulfonyl chlorides) forming thioureas (C), sulfonamides (D), or carboxamides (E)

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Fig. 3 Labeling of amino surfaces, generated by plasma deposition of allylamine layers, with amine-reactive probes forming sulfonamides and thioureas (A), and, after the introduction of a glutaraldehyde spacer, with amino-functionalized fluorophores forming azomethines (B)

All the traditional fluorescent labels have in common that the spectral position of their absorption and emission bands is barely affected upon reaction with functional groups. This renders the straightforward spectral discrimination between unreacted and reacted, here surface-attached chromophores, impossible. A distinction can only be achieved with chromogenic and fluorogenic labels that reveal drastic binding-induced changes in the spectral position of their absorption and emission bands and favorably also a strong binding-induced fluorescence enhancement and a change in fluorescence lifetime [42, 43]. These advantages compared to traditional labels encouraged us to also use a more sophisticated chromogenic and fluorogenic pyrylium label Py-1 to fluorometrically characterize amino-functionalized polymer surfaces for the first time (Fig. 4) [44].

Fig. 4 Schematics of the reaction of the fluorescent reporter Py-1 with amino functionalities at PP surfaces


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3 Characterization of PP Films with Commonly Used Surface-Sensitive Techniques The modification of a PP surface with an pulsed oxygen plasma can easily be monitored by Fourier transform infrared spectroscopy in the attenuated total reflection mode (ATR-FTIR) [12]. The FTIR spectra shown in Fig. 5 were recorded with a Nexus instrument (Nicolet, USA) using the ATR accessory (equipped with a diamond crystal, 45◦ angle of incidence, one reflection) or a germanium cell (‘Golden Gate,’ Specac, Kent, UK). The IR signal originates from a near-surface layer of the polymer film with information depth of about 1.5 μm using germanium and 2.5 μm for diamond [12]. Fig. 5 Changes in the ATR-FTIR spectra of PP films (grey line) due to O2 -plasma-chemical modification and a wet-chemical reduction process. Upon derivatization of the OH-functions with TFAA strong TFAA-related C=O (at ca. 1785 cm–1 ) and C-F bands at 1223 cm–1 appear

Figure 5 illustrates the effect of the plasma-chemical modification and the changes in the ATR-FTIR transmittance observed upon subsequent derivatization with TFAA, which exclusively reacts with OH-functionalities. An additional wet-chemical reduction process with diborane [45] was previously applied to the plasma-chemically treated substrates to transform all the different oxygencontaining surface species other than ether bonds into OH-functions. This procedure results in surface densities of about 10–14 OH/100◦ C [33, 34]. After derivatization of the hydroxyl groups with TFAA, a carbonyl band (1785 cm−1 ), caused by ester formation, appears. The signals at 1223 cm−1 are assigned to the valence vibrational band νC-F of surface-attached TFAA molecules. This signal is directly related to the concentration of surface OH groups [12]. The results of the XPS analysis of these samples are summarized in Fig. 6 . These data were recorded with a Sage 150 Spectrometer (Specs, Berlin, Germany) using a non-monochromatized MgKα or AlKα radiation (e.g., [12]). The XPS spectra illustrate various intermediate steps of the above-mentioned modification course applied to the untreated PP film (A). The spectral changes due to treatment with an oxygen plasma (B) and the subsequent reaction with the XPS tag TFAA (C) are displayed additionally. The results clearly confirm the successful attachment of the TFAA tag at the PP surface. Furthermore, the effect of the wet-chemical reduction process

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Fig. 6 XPS spectra of PP films as received (A), modified in an O2 -plasma (B), modified in O2 -plasma and derivatized with the XPS tag TFAA (C). Spectrum (D) reveals the XPS data obtained upon O2 -plasma-chemical modification of the PP films, wet-chemical reduction, and derivatization with TFAA

for the enhancement of the density of OH-surface functions is demonstrated by the increasing fluorine signal in spectrum (D) [12]. Measurements of contact angles can also provide useful information about the changes of the surface properties by plasma modification. Figure 7 shows contact angles with water obtained for dansylhydrazine (DNS-Hy)-modified PP films, using an automatic Contact Angle System G2 (Kruess, Germany). The substrates were pretreated with an O2 -plasma of various power and post-plasma treated with diborane resulting in different concentrations of OH groups at the surface.

Fig. 7 Water contact angles resulting from dansylhydrazine (DNS-Hy)-modified PP surfaces. Before labeling, the PP films were pretreated with an oxygen plasma using a plasma power of 10 W (left panel) and 200 W (right panel)

After application of a weak plasma power, only a low OH density, and therefore a low surface density of the DNS-Hy fluorophores, was achieved resulting in a water contact angle of 91.7 ± 0.6◦ (Fig. 7, left). Oxygen treatment with a stronger plasma power, however, results in a decrease in the water contact angle to 70.5 ± 0.6◦ (Fig. 7, right), thereby indicating an enhanced hydrophilicity of the surface due to the increase in concentration of surface-attached fluorescent labels. These observations are consistent with the results from ATR-FTIR and XPS measurements.


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4 Fluorescence-Based Characterization of Labeled PP Surfaces – Pitfalls and Troubleshooting In comparison to simple measurements in solution, the reliable use of fluorescence techniques in conjunction with labeling reactions for the characterization of target functionalities on solid supports imposes some extra challenges on the spectroscopist [19, 27, 32, 46]. The detection of surface functionalities on solid supports is complicated not only by the likely influence of the surface on the reaction between the surface functionality and the chosen label, as previously discussed, but also by the influence of the surface and the support on optical detection (e.g., scattering and/or autofluorescence) in the case of polymers [2]. In addition, commonly used steady-state fluorescence techniques are not per se surface-sensitive with a limited depth of penetration and observation like ATR-FTIR and especially XPS. Since the observed signals result from both the surface and the bulk material [11], signal contributions arising from background emission either from the polymer support itself [2] and/or from adsorbed fluorophores have to be taken into account to obtain exclusively the emission of the surface-attached fluorophores. The removal of unreacted dye molecules is especially challenging in the case of polymeric materials or other porous systems, where not only adsorptions but also diffusion of label molecules into the material can occur. This is particularly enhanced for materials, functionalized by surface modification techniques like corona treatment or low-pressure plasma treatment [11], that not only produce chemically complex surfaces but also seem to alter the polymer porosity [13]. Furthermore, similarly as, e.g., for biomacromolecules, matrix–dye and dye–dye interactions as well as the microenvironment-dependent spectroscopic behavior of dyes can result in modulations of the absorption and the emission properties as well as in fluorescence quenching [47]. Despite these limitations, the ease of use and the sensitivity of fluorometry with a limit of detection (LOD) in the order of ∼1 × 10−12 mol/cm2 for fluorophores on polymer surfaces [27] render fluorescence techniques in conjunction with fluorescence labeling of surface species (FLOSS [46]) attractive tools for the characterization of functionalized polymer surfaces [48]. As accurate fluorescence measurements always require reference or blank samples for the consideration of autofluorescence and scattering from the matrix or support. For each series of fluorescence measurements, reference samples were prepared from nonfunctionalized polymer films using the same sequence of reactions as employed to covalently attach fluorophores to the corresponding surface-functionalized solid supports. In the following sections, crucial factors are illustrated, which have to be taken into account in order to overcome some of the limitations inherent to fluorescencebased approaches for the characterization of surface functionalities.

4.1 Environment-Dependent Spectroscopic Properties Dye properties like the spectral shape and position of the absorption, excitation, and emission spectrum and especially the molar absorption coefficient and the

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fluorescence quantum yield can strongly depend on the fluorophore’s microenvironment (e.g., its polarity, proticity, pH, viscosity) [49]. Especially charge transfer (CT) dyes like dansyl fluorophores reveal a pronounced dependence of their spectroscopic features on microenvironment [27, 28, 50, 51]. To illustrate the matrix effect displayed by the dansyl moiety, we determined the spectral changes of a spacerlinked dansyl chromophore upon attachment to an OH-functionalized PP support. The fluorescence spectra of the films, shown in Fig. 8, were obtained with PP films fixed between two quartz windows. These films were measured in a 90◦ geometry with the excitation polarizer set to 0◦ and the emission polarizer set to 54.7◦ using a Spectronics Instruments 8100 spectrofluorometer. As follows from Fig. 8, the emission spectrum of both nonspecifically adsorbed and covalently attached dansyl fluorophores are significantly (∼50 nm) blue-shifted in comparison to the fluorescence spectrum of dansyl cadaverine in methanol [52]. The observed hypsochromic shift is indicative of a less polar dye environment at the surface of the dry films [53, 54]. The inset in Fig. 8 displays the full width at half maximum (FWHM) of the emission bands. The FWHM values were calculated from the spectra which had been converted to the energy scale, corrected for the instrument spectral responsivity, and fitted by a Gaussian fit. As to be expected, the fluorescence bands at PP surfaces are about 10% broader than the spectrum of the dansyl chromophore in solution. This reflects most likely the more inhomogeneous environment of the immobilized chromophores at the solid support.

Fig. 8 Normalized fluorescence spectra of dansyl cadaverine in methanol (symbols, from ref. [52]) and spectrally shifted emissions of nonspecifically adsorbed (solid line) and covalently attached dansyl chromophores (broken line). The fluorescence spectra of the films were recorded after excitation at 335 nm. The inset shows the full width at half maximum (FWHM) of the emission bands


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4.2 Spectral Correction Measured fluorescence signals present always a superposition of the signal of the fluorescent analyte and instrumental effects. Figure 9 exemplary displays the influence of the wavelength- and polarization-dependent spectral responsivity of the instrument on the shape and the intensity of the signals by comparing the measured instrument-specific spectra of dansyl cadaverine attached to a PP surface and the corresponding fluorescence spectra, corrected for instrument-related effects. The spectra were recorded with the excitation polarizer set to 0◦ and the emission polarizer set to 0◦ (parallel alignment) and 90◦ (perpendicular alignment). Fig. 9 Apparent emission anisotropy (left) and the effect of spectral correction (right) for a dansyl-labeled PP film measured with perpendicular (solid lines) or parallel (symbols) polarizers. The spectral positions of the emission maxima before and the red-shifted maxima obtained upon spectral correction are indicated by arrows. The excitation wavelength was at 330 nm

The uncorrected emission spectra recorded for both emission polarizer settings differ considerably in intensity and shape (see Fig. 9, left). The bump at ∼500 nm in the uncorrected spectrum obtained with an emission polarizer set to 90◦ arises from Wood anomalies (diffraction effects) of the monochromator gratings of the fluorometer. As a result of the spectral correction, bathochromic spectral shifts are observed and the apparent polarization dependence [55, 56] of the uncorrected fluorescence intensity distributions disappears. The matching emission spectra are indicative of very small emission anisotropy. The data, shown in Fig. 9, clearly demonstrate the need for the elimination of wavelength- and polarization-dependent instrumental effects from raw data by special correction procedures to avoid misinterpretation and to enable the reliable comparison of fluorescence spectra between, e.g., different instruments [57].

4.3 Reproducibility The reproducibility of fluorescence measurements relies critically on different factors. This includes, for example, undesired background fluorescence, photobleaching, the measurement geometry, and the type of sample that in turn determines the

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measurement geometry to be used. Measurements of transparent dilute dye solutions using cuvettes can be performed with a reproducibility of typically ≥98%. However, we found a significantly increased uncertainty for fluorescence measurements of solid supports with a routine fluorometer. Exemplary performed investigations with DNS-Hy-labeled PP films yield an uncertainty of ∼15% determined in the band maximum in spite of extensive washing and sample preparation procedures (see Fig. 10) [48, 55]. This enhanced uncertainty arises probably from a combination of inhomogeneous functionalization/labeling and other sample-related parameters such as, e.g., scattering and reflection, an inadequate elimination of background fluorescence as well as from difficulties in reproducible sample positioning for measurements in front-face geometry. Fig. 10 Relative uncertainty of fluorescence measurements of fluorophore-labeled polymer films derived from the repeatedly recorded excitation (left) and emission (right) spectra of a PP-TDI-DNS-Hy film (after Soxhlet extraction for 18 h). The excitation spectra were recorded at 480 nm, the emission was excited at 335 nm

4.4 Nonspecific Adsorption As previously discussed, in the case of fluorescence measurements of fluorophorelabeled solid supports, the resulting signals contain information on the surface and on the spectroscopic properties of the bulk material. The extent of the latter contribution depends on the wavelength-dependent penetration depth of the exciting light and on the wavelength-dependent collection efficiency of the emitted light. The size of these effects is determined by the choice of the excitation and emission wavelength, the scattering properties of the system, and the occurrence of reabsorption. To minimize undesired signal contributions, especially the emission arising from nonreacted fluorophores has to be taken into account and should be preferably reduced to a minimum. Therefore, reliable fluorescence measurements of functionalized polymer supports or other polymeric and porous materials require extensive washing and purification procedures. The typically drastic effect of purification by a Soxhlet extraction procedure exemplary shown in Fig. 11 follows immediately from


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Fig. 11 Fluorescence intensity of a PP film modified with TDI-DNS-Hy (left panel; upper set of curves) and the decreased intensity after Soxhlet extractions (18 h) with ethanol (left, lower set of curves). The right panel shows the corresponding mono-exponentially fitted time development in the relative integral fluorescence emission It /I0

the loss of the initial fluorescence signal of ca. 50% for plasma-chemically modified PP films reacted with dansyl labels. Contrary to spectrofluorometry that is not a surface-sensitive technique, more sophisticated fluorescence techniques like surface plasmon fluorescence spectroscopy [58], total internal reflection fluorescence microscopy (TIRF) [59], and confocal laser scanning microscopy (CLSM) [60] providing spatially resolved emission intensities can yield fluorescence data exclusively from the film surface. This encouraged us to examine fluorophore-labeled PP films with spectrally scanning CLSM, using an Olympus Fluoview 1000, offering a spectral resolution of 2.0 nm. To the best of our knowledge, this is the first example of CLSM measurements on plasma-chemically functionalized and fluorophore-modified polymer surfaces. The spatially x–y resolved emission spectra of a thin hydroxy-modified PP film bearing TDI-spacer-linked dansylhydrazine molecules strongly resemble the corresponding spectra recorded with a spectrofluorometer in a front-face geometry (see Fig. 12, left, panel A). Using a CLSM, however, additional information on the emission intensity as a function of z-position of a transparent material is accessible. The z-scan enables the distinction between surface-attached dye molecules and fluorophores diffused into deeper layers of the substrate as illustrated in Fig. 12B. This figure summarizes the intensity profile obtained from a CLSM z-stack of the labeled PP film (Fig. 12B, top) and the effect of a Soxhlet extraction procedure on such a profile (Fig. 12B, bottom). Directly after labeling and before an extended extraction procedure, an almost uniformly distributed signal, independent from the z-position within the film, is recorded (Fig. 12B). The measured intensity profile is attributed to dye penetration into the plasma-modified polymer film [61]. After Soxhlet treatment, the highest signal intensities clearly result from the plasma-chemically modified PP surface, indicating a stable surface immobilization of the dansyl fluorophores (Fig. 12B, bottom). This underlines two facts: the importance of crucial washing procedures and the potential of CLSM for sensitive surface analysis.

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Fig. 12 Emission spectra recorded with a spectrofluorometer (symbols) and with a CLSM (solid line) (A) and the visualization of the effect of a purification process applied to a thin plasma-chemically hydroxy-modified PP film (thickness ∼80 μm) labeled with TDI-spacer-linked dansylhydrazine (B). The intensity profiles (excitation with an Ar+ laser at 364 nm, Olympus Fluoview 1000) result from a CLSM z-stack, directly measured after dye labeling (top) and after Soxhlet extraction (ethanol, 18 h) of the PP film (bottom)

4.5 Correlation of Fluorescence Measurements and XPS Characterization Quantitative fluorescence analysis of labeled surface functionalities is still a challenging task, which is commonly based on calibration curves generated from known analyte concentrations in solutions [27, 30]. However, the sensitivity of the absorption and emission properties of a dye to its microenvironment can cause significant changes in the spectral position of the absorption and emission bands for surfaceattached species in comparison to chromophore molecules in solution, see also Fig. 8 [32, 62]. Moreover, the molar absorption coefficient and the fluorescence quantum yield can vary considerably. These changes are typically not directly accessible. Accordingly, such a calibration approach can result in notable uncertainties in quantification depending on the size of the environment-dependent changes of the signal-relevant spectroscopic properties. The lack of knowledge of the polarity of the dye microenvironment at surfaces that is required to find a solvent of very similar polarity for an improved calibration curve is critical. To overcome these limitations, the applicability of several surface analytical techniques as independent calibration tools has been tested as a basis for method validation [12, 19, 24]. This is exemplary shown in Fig. 13 for dansyl-labeled films. This figure displays the data from water contact angle measurements, the atomic content of sulfur atoms as determined by XPS, related to the amount of reacted functional groups per 100 carbon atoms in a 5–7 nm thick surface layer, and the results from fluorescence measurements of the same dansyl-labeled films. In all cases, the measured data are plotted as a function of the O2 -plasma power applied to generate surface OH-functions, which provides a certain control of functionalization density.


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Fig. 13 Correlation of commonly used surface-sensitive techniques XPS and water contact angle measurements with results from fluorescence measurements of a bulky film of TDI-spacer-linked dansylhydrazine (excitation at 350 nm)

It is noteworthy here that the commonly required additional chemical derivatization step prior to the XPS characterization (cf. Section 2.2, Fig. 1) was circumvented by utilizing the sulfur atoms of the SO2 moiety or the N atoms of the isocyanatecoupled dansyl fluorophores as internal XPS tags. As previously discussed, this is a prerequisite for an improved comparability of data. The comparison of the results derived from the different analytical methods reveals the same general trend, see Fig. 13. Functionalization in an O2 -plasma (continuous-wave mode, 100 W, pressure of 10 Pa), was estimated to yield fluorophore concentrations of about 0.3– 0.4 nmol/cm2 at the PP surfaces, assuming almost quantitative labeling reactions [11, 32]. Within the examined concentration range found between the limit of detection of ∼10−3 nmol/cm2 and dye concentrations >1 nmol/cm2 , where quenching of the emission was reported for dansyl chromophores [27], spectrofluorometry presents a complementary technique for the surface analysis of polymer films and supports. The suitable concentration range depends on the choice of label and its tendency toward aggregation which should be checked in solution prior to use. The results shown in Fig. 13 can also pave the road to energetically control the density of surface OH-functions by variation of the plasma conditions.

4.6 Chromogenic and Fluorogenic Labels To elegantly circumvent the problems related to enhanced background fluorescence due to nonspecifically adsorbed fluorophores at the surface or dye molecules incorporated into the matrix, fluorescent probes are desired that undergo changes in their spectroscopic features upon covalent attachment to functional groups [42, 43, 63]. Suitable molecules should signal the binding event through drastic changes in

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energy and in intensity in absorption and emission. Changes in fluorescence lifetime can also be favorable, although this parameter has been rarely used for the characterization of surface functionalities. The best known example for a label revealing binding-induced spectroscopic changes is the amine-sensitive label fluorescamine [43] extensively used in bio- and surface analysis [27, 64, 65]. Nonfluorescent fluorescamine is a light up probe, the emission of which is switched on only upon the binding event. However, the resulting emissive chromophore absorbs only at short wavelengths (excitation at 390 nm recommended). Principally more attractive are fluorescent reporters that can be excited in the visible (VIS) or near-infrared (NIR) region, thereby minimizing signal contributions from scattering and autofluorescence. This is the case for the chromogenic and fluorogenic pyrylium dyes used here [63, 66, 67]. As a result of the chemical reaction of these dyes with primary amino groups, their absorption and emission maxima shift hypsochromically and, depending on the system, the fluorescence quantum yield can strongly increase. Pyrylium dyes have been successfully exploited to characterize amino functionalities in biogenic amines [68] and proteins [42, 63, 69] in solution. However, to the best of our knowledge, up to now, no attempts have been reported so far to adapt this strategy to the fluorometric characterization of amino-functionalized surfaces. This motivated us to study the potential of the pyrylium VIS dye Py-1 for the labeling and the monitoring of surface amino groups of plasma-chemically modified PP films (cf. Fig. 4). The spectroscopic changes accompanying the reaction of Py-1 with amino-terminated polymer films compared with the control sample, obtained from nonfunctionalized PP films, are displayed in Fig. 14.

Fig. 14 Hypsochromic shifts of the excitation and emission spectra accompanying the reaction of Py-1 with amino-functionalized PP surfaces. Normalized spectra of the PP-Py-1 conjugate (open symbols) were excited at 503 nm and fluorescence spectra of the control (solid lines) at 621 nm. The corresponding excitation spectra were recorded at 602 nm and 665 nm, respectively. The inset shows the enhanced integral emission of a Py-1-reacted PP film and the corresponding control using the excitation/emission conditions 503/602 nm (grey bars) and 621/665 nm (black bars), respectively


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The results shown in Fig. 14 highlight the potential of Py-1 for fluorometric monitoring of amino functionalities on solid supports exploiting both the strong binding-induced spectral shifts in absorption and emission and the significant enhancement in fluorescence intensity. The observed increase in intensity also indicates a relatively nonpolar environment of amino-reacted Py-1, which is in agreement with the fluorescence studies on polymer films with surface-attached dansyl chromophores [27, 32, 38]. The pronounced changes in the spectral features initiated by the derivatization reaction enable a straightforward distinction between the unreacted pyrylium dye and the surface-linked pyridinium-type label molecules. Thus, the background signal can be strongly reduced by proper choice of the excitation wavelength. Moreover, as illustrated by the results from CLSM measurements, the spectral changes revealed by Py-1 provide an additional spatial resolution for the distinction between dye molecules attached to amino groups localized at the PP surface and fluorophores diffused into the deeper layers of the substrate [70]. In Fig. 15, the intensity profile from optical sections through an amino-functionalized and Py1-treated PP film obtained by CLSM is displayed. The intense signal from the functionalized surface clearly indicates an effective Py-1 reaction exclusively with plasma-chemically deposited amino surface functionalities. The binding-induced increase in fluorescence quantum yield of the pyridinium dye, in conjunction with

Fig. 15 Emission intensity profile as a function of z-position of an amino-modified polymer support after exposure to Py-1 (film thickness ca. 100 μm; excitation with an 1 mW HeNe laser at 543 nm)

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the possibility to measure against an almost dark background, even if nonreacted pyrylium label is still present, provides an outstanding basis for a fast and sensitive detection of NH2 groups at solid supports.

5 Summary The potential of spectrofluorometry and fluorescence microscopy for the characterization and the quantification of different surface functionalities like OH and NH2 groups was assessed using traditional reactive dyes such as dansyl derivatives and exemplary also a VIS-excitable chromogenic and fluorogenic label, the sterically non-hindered pyrylium label Py-1. In addition, several sources of uncertainty inherent to the fluorometric characterization of surface functionalities at solid supports and porous materials were illustrated, ranging from environment-dependent dye absorption and emission features over spectral correction and nonspecific adsorption to the critical influence of label choice on the measured background. The results clearly underline the advantages of chromogenic and fluorogenic labels that reveal strong binding-induced changes in energy and intensity in absorption and emission as a prerequisite for the circumvention of the simultaneous excitation of unreacted, nonspecifically adsorbed dye. In addition, the great potential of spatially resolved fluorescence microscopy for fluorometric surface analysis was demonstrated. Acknowledgments Financial support from the Federal Ministry of Economics and Technology (BMWi) and the German Ministry of Education and Research (BMBF) is gratefully acknowledged. We would also like to thank Mrs. R. Decker and Mrs. M. Spieles for accurate technical assistance.

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Fluorescent Labeling and Its Effect on Hybridization of Oligodeoxyribonucleotides Ramendra K. Singh and Shipra Agarwal

1 Introduction Biophysical labeling involves selective modification of biological objects with various types of labels capable of providing information on their structure, molecular dynamics, and mechanism of actions. Nowadays, biophysical labeling methods are being used to solve a number of structural problems. These include methods of spin, triplet, photochromic, electron scattering, Mossbauer and NMR, radioactive and fluorescent labeling [1]. The most widely used labels in investigations associated with oligonucleotides/nucleic acids are the radioisotopes on account of their high detection sensitivity. However, nowadays, fluorescent labeling is being explored as a potential alternative to radioactive labeling for oligonucleotides/nucleic acids. Fluorescent nucleic acids in particular are important tools in molecular biology, diagnostics [2], and structural studies [3]. In order to make oligonucleotides fluorescent, a wide range of chemical [4–6] and enzymatic [7, 8] methods have been developed. Since even a little change in the conformation of a nucleic acid can lead to a reduced specificity of hybridization with its target, it is a challenge for an organic chemist to develop nucleic acid analogues and procedures for different applications that do not disturb the functions of the nucleic acids investigated. Radioactive labels are the most frequently used ever since the inception of recombinant DNA techniques [9]. The radioactive isotopes called radionuclides are unstable and decay with the coincident emission of energetic particles and electromagnetic radiation. The extensively used radionuclides for labeling of oligonucleotides/nucleic acids are 32 P, 35 S, 125 I, and 3 H. These can be easily incorporated into nucleic acids by a variety of enzymatic and chemical methods. Once incorporated into oligonucleotides, these show high detection sensitivity and hence are labels of choice.

R.K. Singh (B) Nucleic Acids Research Laboratory, Department of Chemistry, University of Allahabad, Allahabad-211002, India e-mail: [email protected]


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Although the use of radionuclides for labeling of oligonucleotides/nucleic acids ensures high sensitivity of detection, it is associated with inherent disadvantages like cumbersome nature of detection (autoradiography/scintillation counting methods), isotopic instability, health hazards, and disposal problems [10]. These are, therefore, excluded from many clinical applications associated with oligonucleotides/nucleic acids. In contrast to radioactive probes, the non-radioactive probes eliminate the need of specific isotope facilities as well as radioactive waste management. The fluorescent probes are one of the most popular non-radioactive probes in cell biology. A fluorescent probe is a fluorescent chromophore designed to localize within a specific region of a biological specimen or to respond to a specific stimulus. In addition to the high detection sensitivity, the fluorescence process is sensitive to subtle changes in molecular environment. Therefore, the use of fluorescent probes is a powerful method to obtain information about the structure, the function, and the health of cells.

2 Fluorophores Fluorescent molecules represent another class of reporter groups which are being used for labeling of biomolecules particularly oligonucleotides and/or nucleic acids [11]. These are now largely replacing radioisotopes in biophysical labeling on account of the following reasons: 1. These molecules on excitation emit light in UV-Vis region and hence can be detected directly without requiring any procedural workup. 2. Using fluorophores means the problems associated with radioisotopic labeling, viz. instability, health hazards, and disposal problems, can be avoided [10]. 3. The phenomenon of fluorescence itself is dependent on local environment (solvent, pH, temperature, concentration, etc.) and hence can be used to probe molecular interactions [12], cellular functions, biochemical processes [13], etc. 4. Since a number of fluorophores with discernible spectra are available, it is possible to incorporate different fluorophores at different sites within the same molecule and detect them simultaneously [14]. Further, the advent of multiple labeling [14], fluorescence resonance energy transfer (FRET) [15], and introduction of new hardware and detection system ensure that fluorescence detection can be performed with almost the same level of sensitivity as radioisotopic detection. Hood et al. [16] introduced automated nucleic acid sequencing system using laser-based fluorescence detection and since then fluorescently labeled oligonucleotides/nucleic acids are being used in a variety of biological applications [17].

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2.1 General Characteristics of a Fluorophore There are certain minimum structural requirements which, along with chemical environment, determine whether a substance shall fluoresce or not. These are listed as follows: (a) Only those molecules which absorb in the UV-Visible region show fluorescence. In general, the greater the absorbance of a molecule, the greater its emission. Hence molecules with conjugated system show fluorescence. (b) The transition involved in this phenomenon may be either n–π ∗ or π –π ∗ . However, molecules involving π–π ∗ transition give better emission due to greater absorptivity and lower inherent lifetime associated with these systems. (c) In general, aromatic hydrocarbons (particularly fused aromatic systems) and heterocycles with one or more aromatic nuclei fused to heterocyclic rings show more prominent fluorescence properties, though fluorescence can also be observed in some conjugated aliphatic/alicyclic systems. (d) Substitutions do affect fluorescence properties of a molecule. Electronreleasing substituents have favorable effect on fluorescence, whereas electronwithdrawing substituents and heavy atoms have adverse effect on fluorescence. (e) Besides the above factors, structural rigidity may also influence fluorescent properties of a molecule. Therefore, the fluorophores routinely employed in oligonucleotide chemistry are generally planar and conjugated organic molecules with or without fused aromatic rings, like biotin [18], fluorescein [19–25], digoxigenin [26–28], dansyl chloride [29–32], mansyl chloride [33], 4,6-diamino-2-phenyl indole [34–36], tetramethyl rhodamine [37], pyrene [38–41], acridine [42, 43], ethidium bromide [44], naphthalene derivatives.

2.2 Synthesis of Fluorophores The fluorophores have been designed with properties like high solubility in polar solvents, easy delivery into cells, and imaging bio-macromolecules using chemical tagging. Five new fluorophores have been synthesized and out of these, two have been used for labeling of nucleosides and oligonucleotides for studying their effect on oligonucleotide hybridization [45]. These fluorophores have very high degree of fluorescence and have detection limit up to 10–9 mol/L. The fluorophores, viz. 6-(6-isobutyrylamino-1,3-dioxo-1H,3H-benzo[de]isoquinolin-2-yl)-hexanoic acid (1), 6-(6-dimethylamino-1,3-dioxo-1H,3H-benzo[de] isoquinolin-2-yl)-hexanoic acid (2), 6-(6-benzoylamino-1,3-dioxo-1H,3H-benzo [de]isoquinolin-2-yl)-hexanoic acid (3), 6-(6-amino-1-oxo-1H,3H-benzo[de] isoquinolin-2-yl)-hexanoic acid (4), and 6-(6-amino-1H,3H-benzo[de]isoquinolin2-yl)-hexanoic acid (5) have been synthesized and characterized by 1 H NMR,

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mass, 13 C NMR, UV spectroscopic methods and elemental analysis. Another series of compounds (I–V) were first synthesized (Scheme 1) as precursors to the fluorophores 1–5 (Scheme 2). Their fluorescence was measured in different organic solvents, viz. methanol, dioxane, methanol–water, and in normal aqueous solution in the presence of inorganic salts like NaCl, KCl, MgSO4 , and NaHCO3 . Their structures are shown in Fig. 1. The parent molecule 6-(6-amino-1,3-dioxo-1H,3Hbenzo[de]isoquinolin-2-yl)-hexanoic acid was prepared according to published procedures [46].

Fig. 1 Structures of fluorophores (1–5)

2.2.1 4-Nitroacenaphthene (I) Dissolve acenaphthene (24.0 g; 0.16 mol) in hot glacial acetic acid (192 mL) and cool gradually to 10◦ C to form crystalline magma. Add nitric acid (12 mL) slowly with constant agitation and cooling. The temperature should not rise above 15◦ C during addition. Allow the temperature of the reaction mixture to rise to 30◦ C–35◦ C in the second half of nitration. Cool the reaction mixture and pour the crude dark yellow precipitate into chilled water (200 mL). Wash the precipitate many times with cold water to make it fully acid free and then crystallize from glacial acetic acid to afford yellow needles of 4-nitroacenaphthene. Yield 17.9 g (57%). Rf 0.4 (hexane:benzene 7:3, v/v); UV (MeOH): λmax 255 and 370 nm. 1 H NMR (CDCl3 , TMS = 0.00): 3.34 (t, 4H), 7.39 (d, 1H), 7.55 (d, 1H), 7.72 (d, 1H), 7.93 (s, 1H), 8.44 (s, 1H); 13 C NMR (CDCl3 ): 34.9, 35.9, 119.2, 123.1, 128.0, 128.6, 129.9,


165

Fluorescence (Iin millions)

25 compound 1 compound 2 compound 3 compound 4 compound 5 compound 6

20

15

10

5

0 334

369

404

439

474

509

544

579

614

649

684

719

754

789

Wavelength (in nm)

Fig. 2 Comparative fluorescence of all fluorophores in MeOH

130.0, 131.6, 132.1, 133.8, 145.8; MS (EI) m/z: 199.06 (M+ ); C12 H9 NO2 C, 72.35; H, 4.55; N, 7.03; O, 16.06; found: C, 72.1; H, 4.65; N, 6.9. 2.2.2 4-Nitro-1,8-naphthalic Anhydride or 6-Nitro-benzo[de]isochromene-1,3-dione (II) Take 4-nitroacenaphthene (10 g; 0.05 mol) in glacial acetic acid (150 mL) and add sodium dichromate (60 g; 0.2 mol) gradually to it with stirring at 22◦ C–25◦ C. Incubate the mixture in boiling water bath for 5 h. Cool, filter the orange precipitate, and crystallize from hot glacial acetic acid to give colorless crystals of 4-nitro1,8-naphthalic anhydride. Yield 9.1 g (80%). Rf 0.5 (benzene); UV (acetone): λmax 255 nm. 1 H NMR (CDCl3 , TMS = 0.00): 8.12 (d, 1H), 8.60 (s, 1H), 8.91 (d, 1H), 8.98 (d, 1H), 9.05 (s, 1H); 13 C NMR (CDCl3 ): 120.5, 123.8, 126.0, 126.5, 126.6, 127.8, 128.1, 129.8, 131.4, 150.2, 162.0; MS (EI) m/z: 243.02 (M+ ); C12 H5 NO5 C, 59.27; H, 2.07; N, 5.76; O, 32.90; found: C, 59.12.1; H, 1.95; N, 5.92. 2.2.3 4-Amino-1,8-naphthalic Anhydride or 6-Amino-benzo[de]isochromene-1,3-dione (III) Dissolve 4-nitro-1,8-naphthalic anhydride (4.3 g; 0.0165 mol) in glacial acetic acid (15 mL), add iron turnings (20.0 g), and then reflux the reaction mixture gently in a boiling water bath for 1.5 h with occasional agitation. Follow the completion of the reaction using TLC on silica gel and a positive ninhydrin test for amino group. Filter the hot solution to separate the unreacted iron, pour the filtrate into cold water, and collect the product. 1 H NMR (CDCl3 , TMS = 0.00): 7.03 (d, 1H), 7.79 (d, 1H), 8.32 (d, 1H), 8.60 (s, 1H), 8.75 (s, 1H); 13 C NMR (CDCl3 ): 106.1, 114.0, 121.3,

Scheme 1 Synthesis of compounds I-V

166 R.K. Singh and S. Agarwal


167

123.6, 124.4, 125.0, 127.8, 129.8, 130.4, 145.7, 162.0; MS (EI) m/z: 213.04 (M+ ); C12 H7 NO3 C, 67.61; H, 3.31; N, 6.57; O, 22.51; found: C, 67.12; H, 3.45; N, 6.49. 2.2.4 4-Nitro-1,8-naphthalimido-N-caproic Acid or 6-(6-Nitro-1,3-dioxo-1H,3H-benzo[de]isoquinolin-2-yl)-hexanoic Acid (IV) Dissolve 6-aminocaproic acid (1.90 g; 0.0145 mol) in anhydrous pyridine (6.0 mL) and add 4-nitro-1,8-naphthalic anhydride (3.523 g; 0.015 mol) in small fractions at room temperature with constant agitation. Stir the reaction mixture at 60◦ C for 1.5 h. Cool the reaction mixture to room temperature, evaporate it in vacuo, and dry over anhydrous P2 O5 . 4-Nitro-1,8-naphthalimido-N-caproic acid (4.54 g) is obtained in 88% yield. Rf 0.4 (butanol:acetic acid:water 4:2:2, v/v); UV (MeOH): λmax 235, 348, and 460 nm. 1 H NMR (CDCl3 , TMS = 0.00): 1.29 (t, 2H), 1.56 (t, 2H), 1.59 (t, 2H), 2.23 (t, 2H), 3.20 (s, 1H), 7.88 (d, 1H), 8.13 (d, 1H), 8.20 (d, 1H), 8.36 (s, 1H), 8.66 (s, 1H); 13 C NMR (CDCl3 ): 24.8, 26.8, 28.7, 35.8, 42.0, 121.9, 125.9, 126.1, 127.0, 131.4, 132.1, 132.5, 136.0,139.3, 156.1, 165.9, 177.0; MS (EI) m/z: 356.1 (M+ ); C18 H16 N2 O6 C, 60.67; H, 4.53; N, 7.86; O, 26.94; found: C, 60.52; H, 4.41; N, 7.95. 2.2.5 4-Amino-1,8-naphthalimido-N-caproic Acid or 6-(6-Amino-1,3-dioxo-1H,3H-benzo[de]isoquinolin-2-yl)-hexanoic Acid (V) Dissolve 6-aminocaproic acid (1.90 g; 0.0145 mol) in anhydrous pyridine (6.0 mL) and add 4-amino-1,8-naphthalic anhydride (3.197 g; 0.015 mol) in small fractions at room temperature with constant agitation. Stir the reaction mixture at 60◦ C for 1.5 h. Cool the reaction mixture to room temperature, evaporate it in vacuo, and dry over anhydrous Na2 SO4 . Purify the product by column chromatography (eluent, 20% hexane in dichloromethane). Yield 3.913 g (80%). Rf 0.54 (CH2 Cl2 :EtOAc 8:2, v/v); UV (MeOH): λmax 260 and 430 nm. 1 H NMR (CDCl3 , TMS = 0.00): 1.29 (t, 2H), 1.56 (t, 2H), 1.59 (t, 2H), 2.23 (s, 2H), 3.20 (t, 2H), 6.79 (s, 1H), 7.55 (d, 1H), 7.82 (d, 1H), 7.93 (s, 1H), 7.97 (d, 1H); 13 C NMR (CDCl3 ): 24.8, 26.8, 28.7, 35.8, 42.0, 111.5, 116.3, 120.4, 125.9, 126.7, 127.0, 130.3, 137.7, 138.3, 151.6, 165.9, 177.0; MS (EI) m/z: 326.13 (M+ ); C18 H18 N2 O4 C, 66.25; H, 5.56; N, 8.58; O, 19.61; found: C, 66.12; H, 5.41; N, 8.75; G◦ : 122.71 kJ/mol; H◦ : –323.37 kJ/mol. 2.2.6 6-(6-Isobutyrylamino-1,3-dioxo-1H,3H-benzo[de]isoquinolin-2-yl)hexanoic Acid (1) Dissolve 6-(6-amino-1,3-dioxo-1H,3H-benzo[de]isoquinolin-2-yl)-hexanoic acid (50 mg, 0.15 mmol) in anhydrous pyridine (2 mL), cool the solution in ice bath, add isobutyric anhydride (3 mL), and remove the reaction mixture from ice bath. Stir

168


Scheme 2 Synthesis of fluorophores 1–5

the reaction overnight. After 24 h, chill the reaction mixture in ice bath and add cold water (2 mL). After 15 min, concentrate the reaction mixture, pour into water, filter, wash with ether, and crystallize the product. Yield 69%. Rf 0.3 (DCM); UV(MeOH): λmax 397 nm; 1 H NMR (CDCl3 , TMS = 0.00): 7.97–6.79 (d, 5H, Ar–H), 3.2–1.29 (d, 8H); 2.78 (d, H), 2.23 (s, 2H); 1.19 (s, 6H); 13 C NMR (CDCl3 ): 18.0, 24.8, 26.8, 28.7, 35.8, 42.0, 111.5, 116.3, 120.4, 125.9, 126.7, 127.0, 130.3, 137.7,


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138.3, 151.6, 165.9, 177.2; MS (EI) m/z: 396.17 (M+ ); C22 H24 N2 O5 C, 66.65; H, 6.10; N, 7.07; found: C, 66.25; H, 6.15; N, 6.9; G◦ : 28.42 kJ/mol; H◦ : –557.21 kJ/mol. 2.2.7 6-(6-Dimethylamino-1,3-dioxo-1H,3H-benzo[de]isoquinolin-2-yl)hexanoic Acid (2) To an ice-cold solution of 6-(6-amino-1,3-dioxo-1H,3H-benzo[de]isoquinolin-2yl)-hexanoic acid (6) (3.26 g, 10 mmol) in dry DMF (10 mL) add tributylamine (4.8 mL, 20 mmol) and stir the reaction mixture for 15 min. Add methyl iodide (1.8 mL, 28 mmol) to the chilled solution dropwise and allow overnight stirring at room temperature. After 24 h, add dry ether (100 mL) and place the reaction mixture in refrigerator for 2 h. It is then concentrated and extracted with DCM and water. Crystals are obtained in DCM fraction in 71% yield. Rf 0.8 (DCM:MeOH 9:1, v/v); UV(MeOH): λmax 268 nm; 1 H NMR (CDCl3 , TMS = 0.00): 7.97–6.79 (d, 5H, Ar–H), 3.2–1.29 (t, 8H); 2.23 (s, 2H); 2.85 (s, 6H); 13 C NMR (CDCl3 ): 24.8, 26.8, 28.7, 35.8, 42.0, 44.0, 116.8, 121.0, 126.6, 127.7, 128.5, 134.4, 138.6, 161.3, 165.9, 177.0; MS (EI) m/z: 356.16 (M+ ); anal. calcd for C20 H22 N2 O4 C, 67.78; H, 6.26; N, 7.90; found: C, 67.7; H, 6.19; N, 7.94; G◦ : 183.88 kJ/mol; H◦ : –330.91 kJ/mol. 2.2.8 6-(6-Benzoylamino-1,3-dioxo-1H,3H-benzo[de]isoquinolin-2-yl)hexanoic Acid (3) Dissolve 6-(6-amino-1,3-dioxo-1H,3H-benzo[de]isoquinolin-2-yl)-hexanoic acid (1.63 g, 5 mmol) in dry pyridine (25 mL), add benzoyl chloride (5.8 mL, 49.7 mmol), and reflux the reaction mixture for 1 h. Cool it gradually and add ice. Add 5% NaHCO3 (50 mL) solution after 1 h and extract with DCM and water. White crystals of the product are obtained in 83% yield. Rf 0.7 (DCM:MeOH 9:1, v/v); UV(MeOH): λmax 388 nm; 1 H NMR (CDCl3 , TMS = 0.00): 7.97–6.79 (d, 5H, Ar–H), 7.95–7.44 (m, 5H Ar–H); 3.2–1.29 (t, 8H); 2.23 (s, 2H); 13 C NMR (CDCl3 ): 24.8, 26.8, 28.7, 35.8, 42.0, 111.5, 116.3, 120.4, 125.9, 126.7, 127.3, 128.6, 130.3, 131.9, 133.5, 137.7, 138.3, 151.6, 165.9, 177.0; MS (EI) m/z: 436.15 (M+ ); anal. calcd for C25 H22 N2 O5 C, 69.76; H, 5.15; N, 6.51; found: C, 69.71; H, 5.0; N, 6.72; G◦ : 168.53 kJ/mol; H◦ : –377.32 kJ/mol. 2.2.9 6-(6-Amino-1-oxo-1H,3H-benzo[de]isoquinolin-2-yl)-hexanoic Acid (4) To a mixture of water and HCl (1.25 + 0.04 mL), add Zn (1.0 g) and HgCl2 (40 mg) and stir the mixture for 15 min. Decant the liquid and use this amalgamated zinc immediately. Amalgamated zinc (0.5 g) and conc. HCl (0.3 mL) are gently refluxed for 15 min. Mix 6-(6-amino-1,3-dioxo-1H,3H-benzo[de]isoquinolin-2-yl)-hexanoic acid (25 mg, 0.076 mmol), conc. HCl (0.6 mL), and absolute ethanol (1.7 mL) in a flask and add dropwise to the above solution over 5 min. After addition, reflux it for 30 min and decant the liquid from Zn metal pieces. Extract the reaction mixture with DCM and water. Recover the product in 51% yield. Rf 0.8 (DCM:MeOH 9:1,

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v/v); UV(MeOH): λmax 352 nm; 1 H NMR (CDCl3 , TMS = 0.00): 7.58–6.62 (m, 5H, Ar–H); 4.9 (s, 2H, Ar-H); 3.2–1.29 (t, 8H); 2.23 (s, 2H); 13 C NMR (CDCl3 ): 24.8, 27.1, 29.2, 35.8, 47.5, 53.7, 107.9, 119.3, 121.4, 123.6, 125.6, 127.1, 129.0, 134.4, 143.2, 166.4, 177.0; MS (EI) m/z: 314.15 (M+ ); anal. calcd for C18 H20 N2 O3 C, 69.21; H, 6.45; N, 8.97; found: C, 68.9; H, 6.59; N, 9.0; G◦ : 82.14 kJ/mol; H◦ : –309.01 kJ/mol.

2.2.10 6-(6-Amino-1H,3H-benzo[de]isoquinolin-2-yl)-hexanoic Acid (5) Compound 5 is prepared by repeating the above procedure and obtaining in 48% yield. Rf 0.6 (DCM:MeOH 9:1, v/v); UV(MeOH): λmax 340 nm; 1 H NMR (CDCl3 , TMS = 0.00): 7.47–6.39 (m, 5H, Ar–H); 4.06 (s, 4H, Ar–H); 2.36–1.29 (t, 8H); 2.23 (s, 2H); 13 C NMR (CDCl3 ): 24.8, 27.4, 29.8, 35.8, 53.9, 60.2, 109.3, 119.2, 120.6, 123.0, 124.5, 125.6, 126.2, 130.2, 132.0, 140.5, 177.0; MS (EI) m/z: 300.17 (M+ ); anal. calcd for C18 H22 N2 O2 C, 72.46; H, 7.43; N, 9.39; found: C, 72.5; H, 7.3; N, 9.4; G◦ : 204.73 kJ/mol; H◦ : –171.31 kJ/mol.

2.3 Fluorescence Studies on Fluorophores 1–5 In order to enhance the fluorescence of the parent compound 6, certain substitutions have been introduced at C-4 position of naphthalimide derivative using isobutyryl, dimethyl, and benzoyl groups. As evident from Fig. 2, molecule 3 reported herein has extended system of conjugation, hence it is expected to give better emission but it gave poor emission signals, probably because of poor nature of electron donation by benzoyl group. In the case of compounds 1 and 2, fluorescence signals get enhanced as these molecules donate electrons to naphthalimide ring. It is well supported by ample literature that electron-donating chromophores enhance the fluorescence. An interesting result is obtained when one of the oxo groups is reduced (compound 4). The fluorescence decreases drastically, whereas the fluorescence remains almost the same as that of the original compound 6 when both of the oxo groups are reduced (compound 5) albeit the color changes from greenish yellow to greenish red, i.e., emission shifts to higher wavelength. The decreasing order of fluorescence of these fluorophores is 1>2>6≈5>4>3. The fluorophores 1 and 2 are chosen (as both of these have higher degrees of fluorescence than others) for labeling of nucleosides and oligodeoxyribonucleotides and studying their effect on oligodeoxyribonucleotide hybridization. The fluorescence of 1 and 2 has been observed in MeOH:water (50:50) and dioxane. Fluorescence of 1 slightly decreased in MeOH than dioxane and MeOH:water (50:50) (Fig. 3). It showed maximum fluorescence in dioxane and minimum in MeOH, while in the case of 2, it showed maximum fluorescence in dioxane but minimum in MeOH:water (50:50) (Fig. 4). Polar fluorophores show large sensitivity to solvent polarity. Shift in emission wavelength in different solvents is because of the polar nature of compounds


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30 1 in MeOH:water 1 in dioxane

Fluorescence (in millions)

25

20

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10

5

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472

494

516

538

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670

692

Wavelength (in nm)

Fig. 3 Comparative fluorescence of fluorophore 1 in MeOH:water and dioxane


25 2 in MeOH:water 2 in dioxane

20

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5

0 350

380

410

440

470

500

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560

590

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wavelength (in nm)

Fig. 4 Comparative fluorescence of fluorophore 2 in MeOH:water and dioxane

1 and 2. Generally solvent effects occur whenever a fluorophore is dissolved in any solvent and is independent of the chemical properties of fluorophore. As the solvent polarity is increased, the emission spectra continue to shift to longer wavelengths. It can be seen in Figs. 3 and 4 that emission spectra have been shifted a lot from polar aprotic solvent to polar protic solvent. Fluorescence of 1 and 2 in inorganic media has also been recorded and results showed that fluorophore 1 exhibited higher degree of fluorescence in inorganic media than did 2 and hence it may be a better choice for biological systems. Their excitation and emission spectra are recorded separately (Figs. 5 and 6). The decreasing order of fluorescence of fluorophore 1 was in KCl>NaHCO3 >NaCl>MgSO4

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R.K. Singh and S. Agarwal 80


70

NaCl 2 MgSO4 2 NaHCO3 2 KCl 2 NaCl 1 MgSO4 1 NaHCO3 1 KCl 1

60 50 40 30 20 10 0 350

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390

400

Wavelength (in nm)

Fig. 5 Excitation spectra of fluorophores 1 and 2 at 397 and 368 nm in inorganic media 70


60 50 NaCl 2 MgSO4 2 NaHCO3 2 KCl 2 NaCl 1 MgSO4 1 NaHCO3 1 KCl 1

40 30 20 10 0 401

421

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461

481

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521

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561

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Fig. 6 Emission spectra of fluorophores 1 and 2 in inorganic media

solutions while that of 2 was in MgSO4 >NaHCO3 >KCl≈NaCl solutions. However, the fluorescence of both the fluorophores was lower in inorganic media than in organic media. Emission spectra of all these compounds are recorded and their photophysical properties are summarized in Table 1. For fluorescence recording, emissions are recorded at their excitation wavelengths for all samples at the same concentration. Thus, from the studies it can be concluded that out of the five fluorophores reported herein, 1 and 2 show better emission characteristics than 3–5. From the studies of comparative emission spectra, it can also be concluded that the compounds reported herein show better emission in organic solvents as compared to


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Table 1 Photophysical properties of fluorophores 1–5 Compounds

λmax (nm)

ε (mol/cm L × 105 )

λex (nm)

λem (nm)

Φ

1 2 3 4 5

397 368 384 352 337

1.12 1.05 0.73 0.61 0.99

397 368 388 352 340

446 470 465 425 390

0.52 0.47 0.12 0.29 0.36

inorganic solvents. Among organic solvents, protic solvents are found to have more prominent effect than non-protic solvents. The presence of inorganic ions shows better emission than that observed in pure aqueous solution.

3 Synthesis of Fluorescently Labeled Nucleosides and Nucleotides 3.1 Generation of Linker Molecule at Nucleosides For non-isotopic labeling of oligonucleotides or DNA, fluorescent molecules, chromophores, or chemiluminescent molecules can be incorporated by enzymatic [7, 8, 47–51] or chemical [4–6, 46, 52–57] methods. Due to high stability, in acidic and basic medium, of pyrimidine nucleosides than purine nucleosides, pyrimidine nucleosides are easy to modify. The imide functions on thymine analogues do not require additional protection before oligonucleotide synthesis. From a chemical point of view, working with the adenine and guanine nucleosides is very difficult because of lessened solubilities, pH sensitivity, and the need for additional blocking groups on the exocyclic amines. The C-5 position of pyrimidines is not involved in hydrogen bonding and faces outward in the groove of the helix in a double-stranded DNA. This provides considerable steric tolerance and makes the C-5 position ideal for attachment of linker arm and reporter groups with little effect on hybridization. The most convenient and common method of C-5 derivatization involves direct mercuration [58] or halogenation at C-5 position of unprotected 2 -deoxyuridine (dU) or 2 -deoxycytidine (dC) and subsequent alkylation by olefins in the presence of palladium [59]. Methods for covalent labeling of oligonucleotides may generally be classified into pre- or post-synthetic strategies. The former strategy incorporates a probebound compound into the oligonucleotide during its chemical synthesis [60–63]. The probe is attached to a monomer prior to the condensation step. The base may be already labeled or the label may be attached to a ‘linker arm’ after oligonucleotide synthesis. The latter strategy incorporates a reactive group and connects the finished DNA with the probe afterward, utilizing the reactive group.

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Prior to the synthesis of nucleosides, a five-carbon chain has been introduced at C-5 position of 5-bromouracil and 5-iodo-2 -deoxyuridine. Nucleoside of 5-bromouracil has been synthesized by first attaching 5-aminopentanol-1 at C5 position and then coupling it with ABR (β-D-ribofuranose-1-acetate-2,3,5tribenzoate). At the C-4 position of 2 -deoxycytidine, a 10-carbon chain using spermine as linker has been attached followed by amidite formation using 2-cyanoethyl-N, N-diisopropylchlorophosphoramidite. The linker arm has been attached at the C-5 position of 5-bromouracil by using Williamson’s synthetic procedure for ether bond formation. FMOC-protected 5-aminopentanol-1 is used as the linker arm (Scheme 3).

Scheme 3 Amino group protection of aminopentanol-1

3.1.1 5-(9-Fluorenylmethoxycarbonyl) Aminopentanol-1 To a stirred solution of 5-aminopentanol-1 (2 g, 19.2 mmol, 10% aqueous w/v), acetone (40 mL), and 30% solution (w/v) of sodium carbonate, add 9-fluorenylmethyl chloroformate (5.2 g, 20 mmol) dissolved in acetone (30 mL) dropwise over 20 min. Filter the white solid precipitate obtained after 1.5 h stirring, wash with ice cold acetone–water (50 mL), and crystallize the creamish colored product. Yield 5.06 g (80%). Pyrimidine nucleosides substituted at C-5 position constitute a class of biologically significant molecules, but substituents no larger than n-butyl groups are of interest as chemotherapeutic agents [64–67], while other derivatives have found applications in the biochemical and physiochemical study of biological macromolecules [68, 69]. For the attachment of linker molecule 5-aminopentanol-1 to uracil and deoxyuridine, first it is converted into its sodium alkoxide by adding it dropwise to Na metal. This sodium alkoxide facilitates the reaction by removing halogens as sodium salts from aryl halide, i.e., 5-bromouracil and 5-iodo-2 deoxyuridine, and an ether linkage is formed between base/nucleoside which is quite


175

stable in both acidic and basic media. Nucleoside of 5-bromouracil has been synthesized after attachment of 5-aminopentanol-1 at C-5 position using Vorbrüggen procedure [70].

3.1.2 5-[5-N-(9-Fluorenylmethoxycarbonyl)-aminopentanoxy] Uracil and 5-[5-N-(9-Fluorenylmethoxycarbonyl)-aminopentanoxy]-2 deoxyuridine (7) FMOC-protected aminopentanol (1.456 g, 5.14 mmol) is dissolved in DMSO (20 mL), added dropwise to Na metal (140 mg), stirred for 2.5 h, and divided into two parts. 5-Bromouracil and 5-iodo-2 -deoxyuridine (0.8 g each) are added to the first and second part, respectively, and the reaction mixtures are stirred for 18 h at room temperature. The dense violet colored solutions are poured into water saturated with NaCl (150 mL) and extracted with ethylacetate (40 mL × 3) and reduced to gum at 40◦ C. 5-[5-N-(9-Fluorenylmethoxycarbonyl)-aminopentanoxy] uracil and 5-[5-N-(9-fluorenylmethoxycarbonyl)-aminopentanoxy]-2 -deoxyuridine (7) are obtained and used without further purification.

3.1.3 5-[5-N- (9-Fluorenylmethoxycarbonyl)-aminopentanoxy] Uracil (2 ,3 ,5 -tri-O-benzoyl-β- D-ribofuranose) (8) Uracil-bearing side chain at C-5 position (675 mg, 1.37 mmol) is stirred with ABR (781.2 mg, 1.55 mmol) in acetonitrile (23.25 mL) and HMDS (0.26 mL, 1.24 mmol), TCS (0.158 mL, 1.24 mmol), and SnCl4 catalyst (0.22 mL, 1.86 mmol) in CH3 CN (7.8 mL) are added and stirred for 20 h at room temperature. Add DCM (25 mL) and extract the reaction mixture twice with saturated solution of NaHCO3 (aqueous). The organic layer is washed with saturated NaCl (aqueous), dried over anhydrous Na2 SO4 , and reduced to a small volume under vacuum. 3.1.4 5 - O-Dimethoxytrityl-4-N-(tris-4,9,13-triazatridecane-1-yl)-2 deoxycytidine (9) The spermine is attached to 2 -deoxycytidine and this reaction has been completed in several steps. At first, 2 -deoxycytidine is transiently O-protected with trimethylsilyl chloride in pyridine and the excess of silyl reagent is removed under diminished pressure. Then p-toluenesulfonyl chloride (2 molar equivalents) is added and the reaction is continued overnight at 50◦ C. After the workup, including 4-h treatment with ammonia in aqueous pyridine, crude 4-Np-toluenesulfonyldeoxycytidine is obtained. The latter is subsequently reacted with 4,4 -dimethoxytrityl chloride in pyridine to give, after silica gel column chromatography, the desired 5 -O-dimethoxytrityl-4-N-p-toluenesulfonyl-2 -deoxycytidine in 74% overall yield [71].

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Scheme 4 Synthesis of fluorescently labeled 2 -deoxyuridine phosphoramidite. (i) DMSO, 18 h stirring at room temperature; (ii) MeOH, NaOMe, 1.5 h stirring; (iii) 6-(6-isobutyrylamino-1,3dioxo-1H,3H-benzo[de]isoquinolin-2-yl)-hexanoic acid /p-nitrophenol/DCC in dioxane pyridine; (iv) acetonitrile, DIPEA, 2-cyanoethyldiisopropyl chlorophosphoramidite, 1.5 h stirring

The amine N,N -bis-(3-aminopropyl)-butane-1,4-diamine is attached at N-4 position of 2 -deoxycytidine after replacing p-toluenesulfonyl group from 5 -Odimethoxytrityl-4-N-p-toluenesulfonyl-2 -deoxycytidine. This linker arm provides


177

Scheme 5 Synthesis of fluorescently labeled uridine phosphoramidite. (i) DMSO, 18 h stirring at room temperature; (ii) ABR, HMDS, TCS, SnCl4 , CH3 CN, 20 h stirring; (iii) MeOH, NaOMe, 1.5 h stirring; (iv) 6-(6-isobutyrylamino-1,3-dioxo-1H,3H-benzo[de]isoquinolin-2-yl)-hexanoic acid/p-nitrophenol/DCC in dioxane pyridine; (v) acetonitrile, DIPEA, 2-cyanoethyldiisopropyl chlorophosphoramidite, 1.5 h stirring

178


Scheme 6 Synthesis of fluorescently labeled 2 -deoxycytidine phosphoramidite. (i) TMSCl in pyridine/p-TsCl, 10 h stirring, 60◦ C; (ii) pyridine-ammonia, 4 h stirring; (iii) DMTrCl in pyridine, 3 h stirring; (iv) spermine in pyridine, 19 h stirring at 70◦ C; (v) 6-(6-dimethylamino-1,3-dioxo1H,3H-benzo[de]isoquinolin -2-yl)-hexanoic acid/p-nitrophenol/DCC in dioxane pyridine; (vi) acetonitrile, DIPEA, 2-cyanoethyldiisopropyl chlorophosphoramidite, 1.5 h stirring


179

solubility both in water and organic solvents, which allows easy workup and later chromatographic purification. As evident from literature, polymethylene linkers have tendency to form coiled structures in aqueous media. This is of crucial importance as the reporter groups that are attached to a synthetic oligonucleotide will ultimately bind with other biomolecules (nucleic acids and/or enzymes) and, thus, would be able to interact in the grooves of DNA. 2 -Deoxycytidine hydrochloride (1.32 g, 5 mmol) is transiently O-protected with TMSCl (2.5 mL, 2 mmol) in anhydrous pyridine (20 mL). To this solution, add p-TsCl (1.9 g, 10 mmol) and continue the reaction overnight at 60◦ C. After workup in dichloromethane and aqueous saturated NaHCO3 , the organic phase is concentrated. The product is desilylated by treating with a mixture of pyridine and concentrated ammonia (15 ml each) for 4 h. Reaction mixture is concentrated. The crude 4-N-p-toluenesulfonyl-2 -deoxycytidine is redissolved in anhydrous pyridine (20 mL), DMTrCl is added (1.7 g, 5 mmol), and the reaction is allowed to proceed for 3 h. After workup in DCM and aqueous 0.5 M NaHCO3 , the organic phase is dried and concentrated. The pure 5 -O-dimethoxytrityl-4-N-p-toluenesulfonyl-2 deoxycytidine is obtained after silica gel column chromatography with DCM and methanol. This compound is again dissolved in anhydrous pyridine, spermine is added (10 equivalents), and the reaction mixture is stirred at 70◦ C for 19 h. After completion of the reaction, it is partitioned between water and DCM and the organic phase is concentrated under reduced pressure and the product is purified by silica gel column chromatography with DCM and methanol. Fluorescent molecules are covalently attached to nucleosides using spacers of different lengths. After the removal of FMOC group from the nucleosides, fluorophores are attached using DCC [72]. The carboxyl groups at fluorophores are activated by p-nitrophenol and an amide bond is formed between fluorophore and linker arm with the help of DCC. Fluorescently labeled nucleosides are obtained in good yield, i.e., 70% after purification on silica gel column chromatography. These compounds are characterized by 1 H NMR, mass, 13 C NMR, UV spectroscopic methods and elemental analysis.

3.2 Attachment of Fluorophore at Linker Arms of Nucleosides The FMOC group at the linker arm in uridine and 2 -deoxyuridine and benzoyl groups at sugar in uridine are removed by treating the protected nucleosides with sodium methoxide at room temperature for 1.5 h. Add DCC (515 mg, 2.5 mol) to fluorophores 1 and 2 (311 mg, 1 mmol each) dissolved in dioxane (4 mL), dry pyridine (0.2 mL), and PNP (140 mg, 1 mmol). The dicyclohexylurea precipitates out after 2.5 h. The precipitate is filtered out and each nucleoside (1 mmol) (7, 8, 9) suspended in DMF (5 mL) is added to the supernatant, separately. To these reaction mixtures, TEA (1 mL) is added, stirred, and left overnight. The reaction mixtures are filtered and dried in vacuo. The products formed are identified by TLC due to the presence of fluorescence and purified chromatographically.

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The suitably protected bases are finally converted to their 3 phosphoramidites using known procedure [73]. The appropriate 3 -cyanoethyl phosphoramidites are prepared by reaction of these bases with 2-cyanoethyl-N,Ndiisopropylchlorophosphoramidite using DIPEA. The products 9, 10, and 11 can be purified on silica gel column chromatography in yield varying between 75% and 85%. These labeled phosphoramidites have been used for the synthesis of pre-modified oligonucleotides [74].

3.3 Synthesis of Labeled Phosphoramidites (10, 11, and 12) The fluorescently labeled nucleosides are finally converted to their respective phosphoramidites. Labeled nucleosides (1.5 mmol of each) are dried under reduced pressure for 5 h and dissolved in anhydrous acetonitrile (7.5 mL) separately and mixed with diisopropylethylamine (1.2 mL, 6 mmol). Now add 2-cyanoethyldiisopropyl chlorophosphoramidite (0.71 mL, 3 mmol) dropwise to the solution and stir for 1.5 h at room temperature. The reaction mixture is worked up with saturated aqueous solution of NaHCO3 and extracted three times with DCM containing 1% TEA. The combined layers are dried with anhydrous Na2 SO4 and evaporated till dryness. The product is purified by silica gel column chromatography in ethylacetate and hexane containing 1% TEA. The fractions containing 3 -O-phosphoramidite are collected and lyophilized. The synthetic procedures are outlined in Schemes 4, 5, and 6. Spectral data for compound 10: 1 H NMR (CDCl3 ): 1.05 (d, 12H), 1.24 (s, 9H), 1.29 (m, 4H), 1.57 (m, 4H), 1.66 (t, 2H), 2.18 (t, 2H), 2.22 (d, 2H), 2.97 (m, 2H), 3.2 (t, 4H), 3.57 (s, 1H), 3.66 (d, 2H), 3.9 (t, 2H), 3.96 (s, 1H), 5.85 (s, 1H), 6.87.97 (m, 18H); 13 C NMR (DMSO): 20.1, 23.7, 23.8, 25.1, 25.7, 26.4, 26.6, 28.3, 30.4, 31.7, 34.3, 36.3, 40.8, 41.6, 42.3, 43.1, 55.8, 56.0, 60.5, 63.5, 77.4, 78.9 ,89.1, 101.8, 114.6, 117.7, 125.8, 126.0, 127.3, 128.4, 129.0, 129.4, 131.0, 133.3, 134.6, 135.3, 143.0, 143.8, 152.5, 159.5, 164.4, 165.9, 168.9, 174.7, 184.7; UV (MeOH) λmax 310; 31 P NMR (CDCl3 ) 146; anal. calcd for C65 H84 N7 O13 P C, 64.93; H, 7.04; N, 8.15; O, 17.30; found: C, 64.54; H, 7.00; N, 8.0; O, 18.53. Spectral data for compound 11:1 H NMR (CDCl3 ): 1.05 (d, 12H), 1.24 (s, 9H), 1.29 (m, 4H), 1.57 (m, 4H), 1.66 (t, 2H), 2.18 (t, 2H), 4.28 (s, 1H), 2.97 (m, 2H), 3.2 (t, 4H), 3.57 (s, 1H), 3.96 (s, 1H), 5.93 (s, 1H), 6.8-7.97 (m, 18H); 13 C NMR (DMSO): 20.1, 23.7, 23.8, 25.1, 26.4, 28.3, 30.4, 31.7, 34.3, 36.3, 40.4, 42.0, 43.1, 55.8, 56.0, 63.8, 65.6, 66.7, 72.7, 74.3, 82.5, 89.1, 101.8, 111.5, 114.6, 116.3, 117.7, 120.4, 125.9, 126.0, 126.7, 127.0, 128.4, 129.0, 129.4, 130.3, 135.3, 137.7, 138.3, 143.0, 151.6, 152.5, 159.5, 164.4, 165.9, 174.7, 181.7; UV (MeOH) λmax 310; 31 P NMR (CDCl3 ) 146; anal. calcd for C67 H82 N7 O14 P C, 64.88; H, 6.66; N, 7.90; O, 18.06; found: C, 64.66; H, 6.5; N, 7.3; O, 18.03. Spectral data for compound 12: 1 H NMR (CDCl3 ): 1.05 (d, 12H), 1.29 (t, 2H), 1.41 (t, 2H), 1.59 (t, 2H), 1.67 (m, 4H), 2.22 (d, 2H), 2.55 (t, 8H), 2.65(t, 2H), 2.85 (s, 6H), 2.97 (m, 2H), 3.2 (t, 4H), 3.57 (s, 1H), 3.91 (t, 1H), 6.8–7.9 (m, 18H); 13 C NMR (DMSO): 14.0, 20.1, 22.4, 23.7, 25.7, 27.1, 29.6, 31.5, 33.6, 34.3, 36.3,


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40.5, 44.0, 46.6, 49.2, 55.8, 56.0, 60.5, 63.5, 77.4, 78.9, 89.1, 98, 114.6, 116.8, 117.7, 121.0, 126.0, 126.6, 127.7, 128.4 , 128.5, 129.0, 129.4, 131, 134.4, 135.3, 138.4, 138.6, 143.0, 158, 159.5, 161.3, 164, 166.2, 174.7; UV (MeOH) λmax 322; 31 P NMR (CDCl ) 146.3; anal. calcd for C H N O P C, 66.38; H, 7.48; N, 3 70 94 10 10 11.06; O, 12.63; found: C, 66.2; H, 7.5; N, 11.0; O, 12.53.

3.4 Fluorescence Studies on Labelled Nucleosides and Their Phosphoramidites The fluorescence of labeled nucleosides and the respective phosphoramidites has been recorded in MeOH and dichloroethane, respectively. Excitation wavelengths are fixed at 397 nm for uridine and deoxyuridine and 368 nm for deoxycytidine. Labeled uridine showed maximum fluorescence, while labeled 2 -deoxyuridine showed minimum fluorescence (Fig. 7). Phosphoramidite of labeled deoxycytidine showed maximum fluorescence, while phosphoramidites of labeled uridine showed minimum fluorescence (Fig. 8). In general, phosphoramidites have shown higher sensitivity than labeled nucleosides. 45

fluorescence (in millions)

40

Uridine deoxyuridine deoxycytidine

35 30 25 20 15 10 5 0 300

328

356

384

412

440

468

496

524

552

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608

636

664

692

wave length (in nm)

Fig. 7 Fluorescence of labeled nucleosides uridine, deoxyuridine, and deoxycytidine

4 Synthesis of Fluorescently Labeled Oligonucleotides For post-synthetic modifications on oligonucleotides, a six-carbon chain of hexamethylene diamine is introduced at 5 -end of two oligonucleotide sequences, oligo-1: DMT-AGT GGG TTA AGA and oligo-2: DMT-TCT TAA CCC ACT, and then fluorophores having six-carbon chain are attached to them. Out of five

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50

Uridine

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30

20

10

0 400

422

444

466

488

510

532

554

576

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664

686

wavelength (in nm)

Fig. 8 Emission spectra of labeled phosphoramidites of uridine, deoxyuridine, and deoxycytidine

synthesized fluorophores, the fluorophores 1 and 2 have been chosen for labeling oligodeoxyribonucleotides due to higher degree of fluorescence. Two sequences complementary to each other labeled at their 5 -end, two singly labeled DNA duplexes and one doubly labeled DNA duplex (Fig. 9), have been prepared using normal hybridization protocols. All oligonucleotides are purified by HPLC and hybridizational studies are done to calculate the melting temperatures of duplexes. The fluorescence studies have been done in phosphate buffer and electrophoretic mobility and fluorescence checked on native and denaturing PAGE (polyacrylamide gel electrophoresis).

4.1 Labeling of Oligodeoxyribonucleotides Using Post-synthetic Modification Approach The following sequences are synthesized following the standard β-cyanoethyl nucleosides phosphoramidite chemistry: oligo-1: DMT-AGT GGG TTA AGA and oligo-2: DMT-TCT TAA CCC ACT, using LCAA-CPG as solid support. The oligodeoxyribonucleotides have been synthesized on Pharmacia LKB Gene Assembler Plus on 0.2 μmol scale using standard protocols. The CPG-attached 12mer DMTd(AGTGGGTTAAGA) is placed into a vial and the resin is washed with anhydrous HPLC grade dioxane (3 mL × 3). Carbonyldiimidazole (50 mg) dissolved in dioxane (1.0 mL) is added to it and the reaction is allowed to proceed for 30 min at room temperature with occasional shaking. After this, the solid support is washed well with dioxane (3 mL × 3). The 5 -activated oligodeoxyribonucleotide

N

NHCOCH(CH3)2

O

d(TCTTAACCCACT) Oligo 2

Complementary sequence

Oligo 1

d(AGTGGGTTAAGA) N

N(CH3)2

O

Oligo 3

d(TCTTAACCCACT)

d(AGTGGGTTAAG A) Oligo 4

Complementary sequence

(CH2)5CONH(CH2)6NHCOO

Fig. 9 Fluorescently labeled oligodeoxyribonucleotides with their complementary sequences

O

O

(CH2)5CONH(CH2)6NHCOO

Fluorescent Labeling and Its Effect on Hybridization of Oligodeoxyribonucleotides 183

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is then allowed to react with hexamethylenediamine (1.0 mL, 0.2 M) dissolved in dioxane:water (9:1) in the same reaction vessel. The reaction is allowed to proceed for 20 min at room temperature and finally, the support is washed with dioxane and methanol. Similarly, the other oligodeoxyribonucleotide DMT-d(TCTTAACCCACT) is also derivatized at its 5 -end to bear a primary amino group. These derivatized oligodeoxyribonucleotides are cleaved from their support by treating with concentrated ammonia (25%, 4 mL) for 1 h at room temperature. The support is filtered out and the aqueous ammonia solution is heated at 55◦ C for 10 h. Ammonia is removed under vacuum and the oligodeoxyribonucleotides are then purified by reversedphase HPLC using C18 Lichrosphere column and gradient buffer CH3 COONH4 , 0.1 M, pH 7.1 (A) and CH3 CN (B) having a flow rate of 1 mL/min and a linear gradient of 0% to 25% B in 25 min. The absorption is monitored at 260 nm for peak detection. The retention time observed for d(AGTGGGTTAAGA) (10 OD, 280 μg) and d(TCTTAACCCACT) (11 OD, 363 μg) is 9 and 8.2 min, respectively. The desired fractions are pooled and lyophilized. p-Nitrophenyl esters [46] of fluorophores can be prepared by dissolving the fluorophores (0.054 mM) separately in dry dioxane (2.0 mL) containing dry pyridine (0.2 mL), TEA (0.2 mL), and p-nitrophenol (7.56 mg, 0.054 mM) followed by the addition of dicyclohexylcarbodiimide (27.8 mg, 0.135 mM) and stirring for 2.5 h. Dicyclohexylurea is removed by filtration to obtain p-nitrophenyl ester of 6-(6-isobutyrylamino-1,3dioxo-1H,3H-benzo[de]isoquinolin-2-yl)-hexanoic acid and 6-(6-dimethylamino1,3-dioxo-1H,3H-benzo[de]isoquinolin-2-yl)-hexanoic acid (yield 65%). 5 -Alkylamino oligodeoxyribonucleotides (6.5 OD and 8 OD for oligo 1 and oligo 2, respectively) are dissolved separately in 250 μL solutions of a mixture of 0.1 M Na2 CO3 /NaHCO3 (pH 9.0) and to these solutions are added 500 μL solutions of p-nitrophenyl ester of fluorophores dissolved in a mixture of 1.0 M Na2 CO3 /NaHCO3 buffer (pH 9.0):DMF:H2 O (5:2:3. v/v) and the reaction mixtures are vortexed and wrapped with aluminum foil to prevent light exposure. The synthetic procedure is outlined in (Scheme 7). The reaction mixture is stirred for 8 h at room temperature in dark. The labeled oligodeoxyribonucleotides are then passed through Sephadex G-25 column using ethanol–water (8:2, v/v) as eluent. The filtrates containing fluorescently labeled oligodeoxyribonucleotides are collected, concentrated, resuspended in water (1.0 mL), and purified by reversed-phase HPLC using C18 Lichrosphere column and gradient buffer CH3 COONH4 , 0.1 M, pH 7.1 (A) and CH3 CN (B) having a flow rate of 1 mL/min and a linear gradient of 0% to 25% B in 25 min. The absorption is monitored at 260 nm for peak detection. The retention time observed for labeled d(AGTGGGTTAAGA) is 16.5 min and for d(TCTTAACCCACT) is 15.8 min (Fig. 10). The desired fractions are pooled and lyophilized. Finally, 150 μg (6 OD; 0.03 μmol) and 165 μg (7 OD; 0.035 μmol) of labeled oligodeoxyribonucleotides could be obtained. Analysis of ESI-MS of labeled d(AGTGGGTTAAGA) and labeled d(TCTTAACCCACT) was m/z = 4285.53 and 4034.67, respectively.

Scheme 7 Fluorescent labeling of oligodeoxyribonucleotides. a. 3% Trichloroacetic acid in dichloromethane; b. carbonyldiimidazole dissolved in dioxane; c. hexamethylenediamine in dioxane:water; d. liquor ammonia, 10 h at 55◦ C, e. p-nitrophenyl ester of fluorophore dissolved in Na2 CO3 /NaHCO3 buffer (pH 9.0):DMF:H2 O 5:2:3, v/v

Fluorescent Labeling and Its Effect on Hybridization of Oligodeoxyribonucleotides 185

186


Fig. 10 RPHPLC profile of oligodeoxyribonucleotides (A) d(AGTGGGTTAAGA) and (B) d(TCTTAACCCACT); 1, oligodeoxyribonucleotide with 5 -DMT group; 2, oligodeoxyribonucleotide with linker arm;3, fluorescently labeled oligodeoxyribonucleotide. Purification was done using C18 Lichrosphere column and gradient buffer 0.01 mM NaH2 PO4 •2H2 O, 0.01 mM Na2 HPO4 , and 1.0 m NaCl, pH 7.2

4.2 Labeling of Oligodeoxyribonucleotides Using Pre-modification Approach For the synthesis of pre-modified oligonucleotides, two 11-mer oligonucleotides, DMT-d(GTGGGTTAAGA) and DMT-d(CTTAACCCACT) attached on LCAACPG (synthesized separately following the above procedure), are taken and fluorescently labeled phosphoramidites (compounds 10 and 12) are attached at the 5 -end of these oligonucleotides [75]. 2 -Deoxyuridine-3 -O-phosphoramidite activated by tetrazole to form 2 -deoxyuridine-3 -O-phosphomonotetrazolide is added onto the column, as it undergoes rapid nucleophilic substitution reaction with hydroxyl group of CPG-bound oligonucleotide and forms an internucleotide phosphite–triester linkage. The coupling yield of this step is 98.9% as detected by trityl analysis. This phosphite–triester is then oxidized to phosphotriester using iodine in THF, 2,6-lutidine, and water. This fluorescently labeled oligonucleotide is delinked from the LCAA-CPG support by ammonia treatment and purified on RP-HPLC (retention time 16 min), after removal of ammonia and phosphate protecting groups, using 0.1 M triethylammonium acetate and acetonitrile to get labeled oligonucleotide A, d(U∗ GTGGGTTAAGA), as shown in (Scheme 8).


187

Scheme 8 Synthesis of pre-modified labeled oligonucleotide

The above procedure is repeated with support-bound DMT-d(CTTAACCCACT) and fluorescently labeled 2 -deoxycytidine-3 -O-phosphoramidite to synthesize fluorescently labeled oligonucleotide B, d(C∗ CTTAACCCACT). It is also purified on RP HPLC showing retention time as 15.6 min (Fig. 11).

4.3 Hybridization and Fluorescence Studies on Labeled Oligodeoxyribonucleotides The following four duplexes for fluorescence and thermal denaturation studies have been prepared. Duplex I has fluorescently labeled d(AGTGGGTTAAGA), Oligo 1, and its complementary strand d(TCTTAACCCACT), Oligo 2, while duplex II has fluorescently labeled d(TCTTAACCCACT), Oligo 3, and its complementary strand, d(AGTGGGTTAAGA), Oligo 4. Duplex III has both labeled strands, Oligo 1 and Oligo 3, while duplex IV has both unlabeled strands, Oligo 2 and Oligo 4 (Fig. 12). The duplexes are formed by mixing the two strands, each having a concentration of 0.0015 μM, in the hybridization buffer. The solutions are heated at 90◦ C for

188


Fig. 11 RPHPLC profile of pre-modified oligodeoxyribonucleotides A and B (16 and 15.6 min, respectively). Purification was done using C18 Lichrosphere column and gradient buffer 0.01 mM NaH2 PO4 •2H2 O, 0.01 mM Na2 HPO4 , and 1.0 m NaCl, pH 7.2

FLUO I

LINKER

d(AGTGGGTTAAGA)

Duplex I Tm = 39°C

d(TCACCCAATTCT)

FLUO II

LINKER

d(TCTTAACCCACT)

Duplex II Tm = 41°C

d(AGAATTGGGTGA)

FLUO I

LINKER

d(AGTGGGTTAAGA) d(TCACCCAATTCT)

d(AGTGGGTTAAGA) d(TCACCCAATTCT)

Fig. 12 Duplexes I–IV and their Tm values

LINKER

FLUO II

Duplex III Tm = 34°C

Duplex IV Tm = 36°C


189

5 min and then cooled to room temperature for 10 min before measuring the melting temperatures. The melting temperature is determined as the maximum of the firstderivative plots of the melting curves obtained by measuring absorbance at 260 nm against increasing temperature (0.5◦ C/min) from 15◦ C to 70◦ C (Fig. 13). 0.135 Duplex III Duplex IV Duplex I Duplex II

Absorbance

0.115 0.095 0.075 0.055 0.035 0.015 15

20

25

30

35 40 45 Temperature (°C)

50

55

60

Fig. 13 Tm graph of duplexes I–IV

All melting temperatures are reported within an uncertainty of ±1◦ C, as determined from multiple experiments. The thermal denaturation studies showed that the singly labeled duplexes I and II melted at a slightly higher temperature (Tm 39◦ C and 41◦ C, respectively) than the unlabeled duplex IV (Tm 36◦ C), whereas the doubly labeled duplex III melted at slightly lower temperature (Tm 34◦ C). The stability of the double helix is largely a function of the type and concentration of counter ions present. Repulsive forces between neighboring phosphate anions tend to unwind the helix in the absence of stabilizing counter ions. Mono and divalent cations that bind to the phosphate groups neutralize this electrostatic repulsion and stabilize the helix. The expansion or the compression of the duplex is a function of cation concentration and has been detected by a number of techniques. Relative helix stability can also be determined by examining changes in the helix–coil transition temperature, Tm , of the double-stranded polynucleotides. As the DNA melts and the two strands coil randomly, the bases unstuck and lead to a cooperative increase in the absorption intensity at 260 nm. This hyperchromicity, monitored as a function of temperature, is a measure of helix stability. This optical assay allows the determination of the stability of DNA in relation to temperature, pH, ionic strength, and added small molecules, and in a variety of polar and non-polar solvents. It has been shown that Tm varies logarithmically with ionic strength; the higher the concentration of phosphate-binding counter ions, the greater the energy required to denature the helix and the higher the Tm . The fluorescence studies of all oligonucleotides and duplexes are done in the hybridization buffer (0.01 mM Na2 HPO4 , 0.01 mM NaH2 PO4 •2H2 O, 1.0 M NaCl,

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R.K. Singh and S. Agarwal 35


30

oligo A oligo B

25 20 15 10 5 0 400

420

440

460

480

500

520

540

560

580

600

620

640

Wave length (in nm)

Fig. 14 Emission spectra of pre-modified labeled oligonucleotides

pH 7.1), each having a concentration of 0.0015 μM. The fluorescence of premodified labeled oligonucleotides (A and B) is scanned in the range of 400–650 nm at room temperature (Fig. 14), while that of post-modified labeled duplexes is scanned in the range of 300–700 nm at room temperature (Fig. 15) and at varying temperatures from 15◦ C to 70◦ C as well (Fig. 16). Oligonucleotides A and B are excited at their excitation wavelengths 397 and 368 nm, respectively. Excitation wavelengths are fixed at 397 and 368 nm for duplex I and duplex II, respectively, and the duplex III is excited once at 397 nm (III) and then at 368 nm (III ). The emissions are recorded at 475 nm. Pre-modified oligonucleotides A and Bhave shown an appreciable degree of fluorescence where oligonucleotide A has higher degree than oligonucleotide B. The study shows that fluorescence of 6-(6-dimethylamino-1,3-dioxo-1H,3Hbenzo[de]isoquinolin-2-yl)-hexanoic acid, 2, increased 1.5 times when used in combination with 6-(6-isobutyrylamino-1,3-dioxo-1H,3H-benzo[de]isoquinolin-2-yl)hexanoic acid, 1 (III in Figs. 15 and 16) as compared to its fluorescence observed in the case of duplex II at the same excitation wavelength, i.e., 368 nm. Further the fluorescence of 6-(6-isobutyrylamino-1,3-dioxo-1H,3H-benzo[de]isoquinolin2-yl)-hexanoic acid decreases to half when used in combination with 6(6-dimethylamino-1,3-dioxo-1H,3H-benzo[de]isoquinolin-2-yl)-hexanoic acid fluorophore at its excitation wavelength 397 nm in comparison to its fluorescence in duplex I. An interesting observation was that the fluorescence of labeled Oligo 3 increased when it formed a duplex II with its complementary sequence. This has been supported clearly by fluorescence measurement at varying temperature where the fluorescence was drastically decreased beyond its melting point, i.e., 41◦ C. However, the fluorescence was almost unchanged in the case of duplexes I and III.


191

Fig. 15 Fluorescence of duplexes excited at their excitation wavelengths at 25◦ C (III shows excitation of duplex III at 368 nm and III shows excitation at 397 nm) 1.5 1.4 1.3 1.2 Fluorescence (in millions)

1.1

Duplex Duplex Duplex Duplex

1 0.9

I II III at 397 III at 368

0.8 0.7 0.6 0.5 0.4 0.3 0.2 0.1 0 15

20

25

30

35

40

45

50

55

60

65

70

Temperature (in C0)

Fig. 16 Spectra of fluorescence of duplexes with increasing temperature from 15◦ C–70◦ C at their excitation wavelength (III shows excitation of duplex III at 368 nm and III shows excitation at 397 nm

4.4 Identification and Electrophoretic Mobility of Labeled Oligodeoxyribonucleotides These labeled duplexes are easily identified on 20% non-denaturing (A and B, Fig. 17) and 20% denaturing (6 M urea) (C, Fig. 17) polyacrylamide gel electrophoresis (PAGE) [76] due to their fluorescent nature. DNA was loaded in

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Fig. 17 Polyacrylamide gel electrophoresis analysis of duplexes I and II. a 20% Native PAGE; b 20% (6 M urea) denaturing PAGE; A is duplex II, B is duplex I, and C is duplex III. Oligonucleotide labeled with compound 2 moved faster on PAGE (A, Fig. 17) and also fluoresced more than oligodeoxyribonucleotide labeled with compound 1 (B, Fig. 17)

6× TBE buffer with 0.25% bromophenol blue and 30% glycerol and gel was run in 1× TBE. The labeled oligodeoxyribonucleotides showed less electrophoretic mobility than unlabeled oligodeoxyribonucleotides, probably because of higher molecular weight.

5 Conclusion Thus, we have developed highly stable and sensitive fluorophores and their fluorescence is not adversely affected by the ions normally present in biological systems during hybridization of oligonucleotides. These molecules also increase the hybridizational affinity of labeled oligonucleotides. The labeled oligonucleotides were easily detected on PAGE. The further course of study involves in situ hybridization and detection of duplexes using fluorescence properties of the fluorophores herein.

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New Method for Determining Histamine Rate in Halieutic Products Alphonse Tine and Stéphy E. Douabalé

Abstract This work focuses on the development of a new method that is a much more reliable and reproducible determination of histamine in fish products. Although low-dose histamine plays a beneficial role in the nervous system as a neurotransmitter molecule, it is one of the leading causes of food poisoning due to fish in most countries. For example, in the United States, it comes second only to tobacco. In fact, histamine is highly toxic; the absorption of a certain amount of histamine can cause allergic reactions causing stomach pain, nausea, vomiting, headache, and itching. Thus the effects of absorption of histamine can be very serious for humans and lead to death. Most poisonings listed are linked to the consumption of certain fish species such as Scombridae (tuna, bonito, mackerel). However, the fish species of other families of healthy fish may be contaminated by microorganisms because of mishandling: First products safely and become the seat of a high concentration of histamine. It is therefore imperative to measure the level of histamine in all fish products prior to export or import. The best current method for determining histamine is the Association of Official Analytical Chemists (AOAC). However, the Laboratory of Photochemistry and Analysis (LPA), Department of Chemistry, Faculty of Science and Technology have proposed a new method which is much more reliable and reproducible. This work was published at the World Intellectual Protection Organization (WIPO) under No. WO 2006/018673 A1 since February 2006 with a deposit of European Patent No. B52105/EP.

A. Tine (B) Laboratoire de Photochimie et d’Analyse, Faculté des Sciences et Techniques, Université Cheik Anta Diop, Dakar, Senegal e-mail: [email protected]; [email protected]


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1 Introduction The histamine molecule was identified only in the early 20th century. In 1916, Guggenheim and Loffler were among the first to isolate that molecule in nature [1]. However, food intoxications attributed to it date far back in history. Indeed, the oldest case of histamine intoxication was reported by Henderson in 1828 [2] following Bonite (Scomber pelamis) consumption. Although histamine was still unknown then, the incriminated fish does not leave any room for doubting the nature of histamine intoxication. Histamine is then one of the primary causes of fish-related food intoxication in most countries. In the United States, it ranks second right under ciguatera [3]. The largest histamine intoxication occurred in Japan in 1973 with 2656 cases following mackerel consumption (Trachus japonicus) [4]. The bulk of work realized on histamine up to this day shows the importance of the molecule in several varied areas of studies like toxicology, neurology, pharmacology, gastroenterology, ophthalmology, biochemistry, and analytic chemistry. Despite the abundance of results, the research of histamine dosage methods is still going on as the sought cost and precision have not been found yet. In this early 21st century, the world population has reached 6 billion and the food resources are not equally available in quantity and quality through the globe. However, for a good diet, food must be diversified with as less nutriments as possible. Nutritionists’ works indicate that fish, meat, and certain vegetal such as soya are the main food protein providers. The WFO is advising a protein-rich diet, knowing that the latter play an important part in the constitution and the functioning of the human body [5]. In addition, the percentages of amine acids that are essential allow us to assert that fish is a major source of proteins [5]. Fish is a worldwide researched food, hence the rules and policies set by the different states of regional communities and WHO [5]. The objective is to protect the consumer. In fact, when it is not well conserved, food can be a public health issue. A bad conservation of seafood will cause alteration and therefore it can result in serious health consequences for consumers. Histamine results from the decarboxylation of histidine, a natural acid amine. That transformation is catalyzed by histidine decarboxylase, an enzyme that is present in certain microorganisms such as Klebsiella oxycota, Klebsiella pneumoniae, Morganella morganii [6–9]. Let us bear in mind that several families host these microorganisms. Among the latter, we will name species like Scombridae and Scomberesocidae families (tunny, bonite, and mackerel) and Coryphaenoides (mahi-mahi) that are major vectors of histamine intoxication [6, 7, 10]. Even worse, sea species in the other hazardless fish families can be contaminated by microorganisms named before by careless handling. The hazardless products then become the receptacle of a strong histamine concentration. That is why it is recommended to follow strict health regulations for the manipulation of halieutic foods [11]. Although histamine concentration in living fish is very slight, thanks to an internal autoregulation [4], it is increased in the postmortem period and without freezing [6, 11, 12]. Health measures are then to be taken. Thus, in some countries; the tolerated ratio of histamine-consumed product is under 50 ppm. The control of histamine rate in food products remains then an important quality test.

New Method for Determining Histamine Rate in Halieutic Products

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All international exchanges are under a regulation destined to preserve public health. Therefore, it is necessary to have accredited laboratories implementing approved dosage methods to control histamine content particularly in sea products. Up to date, several processes of histamine quality and quantity determination have been elaborated. We have enzyme, radio enzyme, electrochemical, chromatographic, photometric, and fluorimetric methods mainly. The choice of a good method is based on several criteria, among which we can name security, specificity, cost, speed, reliability, and limit of detection [15]. Unfortunately, among the current methods; many failures are to be pointed out. The obstacles will vary depending on the method used. For that reason, a study carried on by Rogers and Staruszkiewics on the different dosage methods highlights the failures of each one [16]. – Thin layer chromatography presents a too high detection limit (20–50 ppm). It is then called semiquantitative method. – Biologic methods are also less quantitative; they use for instance the muscle fiber contraction of an animal of little sensitiveness to histamine. Moreover, they require guineas’ animals. – Radio enzyme methods use radio isotopes, which must be handled very carefully. – Enzyme methods are very long because of the duration of the incubation and multiple extractions, all the more as enzymes are not easily available and their purification is tiresome. – Immunologic methods require antibodies and enzymes with very long incubation periods. – Spectrophotometers by optic density measure (electronic observation) are not very quantitative. They involve risks of erroneous results due to their weak domain of calibration. – Electrochemical methods require sophisticated oxygen electrode sounds with use of an enzyme that is not commercially available. – High-performance liquid chromatography poses problems in quantification when the derivation is made in pre-column, while a derivation post-column will generate important variations of absorbance or identity of fluorescence. – Fluometric method is by far the most performing. Histamine not being fluorescent, the fluorescence comes from its complex formed with orthophthalaldehyde (OPA). That method seems to be more precious than the others. In fact, to produce interference between two compounds, they must present the same wavelength of absorption, excitation, and emission. One other advantage of fluometric method is its sensitiveness which allows obtaining very low limits of detection. Thus, compared with the other existing methods, fluometric route dosage is by far the most used method as it is the most sensitive. However, there are major inconveniences related to this method: • Measurement is done by a fluorimeter and not by a spectrofluorimeter. • Kinetics of formation of OPA–histamine complex in basic medium cannot be obtained by fluorimeter. • Acid medium in which fluorescence is measured has proved to be an inhibitor of the intensity of fluorescence of the complex for some values of pH. As

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• • •

a consequence, the calibration domain was reduced; the results obtained become then less reliable, all the more as the difference between two distinct concentrations is less clear. Reaction medium is not optimized according to OPA and pH concentrations. Reaction temperature is not optimized. Statistic results (limit of detection and recovery rate) are not very conclusive in acid medium. This method is not easy to use for the determination of histamine in the medical medium (on human being) where the rate is generally low.

•

This study aims at elaborating a new fluorimetric method based on the kinetic formation of the OPA–histamine complex in basic medium where we find the highest sensitiveness and the widest calibration domain. For that, we will study the effects of pH (NaOH and HCl) of NaCl, and of OPA on the fluorescence of the complex as well as the effect of temperature on the stabilization of OPA–histamine complex in basic medium. We will then compare our results to those obtained by the Official Association of Analytic Chemistry (OAAC) or Lerke and Bell method.

2 Experimental Study 2.1 Products Used Dihydrochlorate of histamine (98%, m/m), histamine (96%), orthophthalaldehyde (OPA, 99%, m/m), and methanol of spectroscopy quality were purchased at Sigma Aldrich (Taufkirchen, Germany) and used without further modification. Hydroxide of sodium (97%, m/m), hydrochloric acid (36%, m/m), acetate of sodium and acetic acid were purchased at Labosi (Oulchy Le Chateau, France). Trichloroacetic acid (TCA 99%, m/m) came from Janssen Chimica (Belgium) and phosphoric acid (85%) from Fluka Chemica (Suisse). Amberlite resin GC-50 (H) type I was delivered by Prolabo (France). Distilled water was used to prepare all the necessary aqueous solutions.

2.2 Instrumentation All fluorescence measurements were taken at ambient temperature with spectrofluorimeter of Kontron brand, SFM25 model, connected to IBM personal computer model Aptiva. Uncorrected spectra of fluorescence were obtained with K-wind 25 software and treated with Microcal Origin software version. We used a Velp R Scientifica magnetic shaker endowed with thermostatic heating system; to record our electronic absorption spectra, we used spectrophotometer UV-Visible of Jasco brand model 7800 associated with Philips monitor of type 80 and a Jasco printer, model PTL-3965. We prepared our buffer liquid with pH-meter of Hanna


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Instruments brand model HI 190 N; a Moulinex grinder was necessary to homogenize the fish samples. A Startorius 0.1 precision balance was used to weight the products. Besides, five polished faced quartz cells were used to measure the fluorescence, while two faced quartz cells were necessary for absorption measures. All cells had 1 cm of optical path. The glasses were constituted of two chromatographic columns, phials, beaker, and straws of several dimensions. A 20 μL micro straw of Pipetman brand and cones were necessary for sampling.

2.3 Experimental Process 2.3.1 Preparation of Solutions Stock solutions of histamine (1 × 10−3 M) were directly prepared in water, while OPA ones (1 × 10−3 M) required a minimum of methanol (1/10, v/v) to make dissolution easier. Serial dilutions have been done to obtain standard working solution. All solutions were protected against light with aluminum paper and kept in a fridge. The mother solutions of HC1 (1 N) and NaOH (1 N) were prepared with distilled water and used with or without preliminary dilutions depending on the specificity of each case.

2.3.2 Preparation of Samples Preparation of sample as well as extraction and purification of histamine were conducted using the Lerkle and Bell method. Changes were made on the level of condensation and the measurement of fluorescence. Actually, at this stage, buffer phosphate solution pH 11.5 was also used.

2.3.3 Measurement of Fluorescence The measurement of fluorescence consisted in preparing OPA–histamine complex in a reactor containing distilled water, OPA, NaOH, and histamine. Sometimes, a quantity of NaCl salt was introduced in this reactor. At any rate, we would take care of introducing either OPA or histamine just before starting the timer as it was at that moment that the reaction would start. To study the effect of pH, we would add measured quantities of HCl or NaOH in the reactor. The shaking and heating time would vary between 0 and 5 min corresponding to the reaction and condensation time. A proportion of that blend was sampled, put in the tube, and taken to the spectrofluorimeter to record the kinetics of the complex. After each measurement, the reactor was washed and rinsed before taking another reaction’s blend. The excitation wavelength is fixed at 350 nm; the interval frequency of the fluorescent spectrum was set between 350 and 550 nm. For kinetics of fluorescence, the excitation wavelength was set at 350 nm and the emission frequency at 430 nm. Kinetics allowed us to measure the stability of the fluorescent complex, and the repetitive kinetic measures

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we had taken according to the concentration of histamine allowed us to establish the straight line calibration. All fluorescence measures were corrected in relation to the signal of the solvent. The results have been expressed at average values of repeated measures. The software Microcal Origin, version 6.0, was used for the statistic treatment of data.

3 Experimental Process 3.1 Optimization of Repetitive Dosage 3.1.1 Verification of the Effect of NaOH Concentration on the Formation of OPA–Histamine Complex by Repetitive Measurement We took OPA–histamine equimolar for which kinetics has been studied at different concentrations of NaOH (Graph 1). We noticed that all the graphs seem to have the same origin. At very slight concentrations of soda equaling 10–4 M, the intensity of fluorescence hardly increases or is slightly increased, and the level at the end of the reaction is very low. For soda concentrations around 10−3 M, the intensity of fluorescence increases rapidly as shown by the steady slope observed before reaching the final level. That level marking the end of the reaction of complex formation is reached very fast, which shows that there is an increase in the speed of the reaction. At more important soda concentrations, above 10−2 M, the reaction duration is longer, so the reaction becomes slower with a significant decrease in fluorescent

Graph 1 Effect of NaOH concentration (mol L−1 ) on formation kinetics of OPA–histamine complex


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intensity. Thus, strong concentrations of soda have an inhibiting effect on the formation of OPA–histamine complex (Graph 2). The catalysis by NaOH is then more efficient for concentrations of 2 × 10−3 M (Graph 1).

40 35 30

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Graph 2 Effect of NaOH concentration (mol L−1 ) on fluorescence of OPA–histamine complex

3.1.2 The Effect of Temperature, Formation Kinetics of OPA– NaOH Complex We have examined the effect of heating time on the kinetic formation of OPA– histamine complex. Kinetic graphs have shown that for the same concentration, the intensity of fluorescence corresponding to the maximum of the lowest level of complex increases with the heating time (Graph 3). So, the heating time exalts the fluorescence of OPA–histamine complex. Here also, we observe a decrease of the complex reaction and formation depending on time and heating. Kinetic graph obtained in Graph 10 corresponds to a strong excess of OPA in the cell. In order to obtain a calibration graph in the slightest concentrations of histamine, we decreased the OPA concentration and plotted kinetic graphs after shaking and heating at 50◦ C (Graph 4). We found intensities of fluorescence relatively constant for each concentration and more so at low levels of concentration. Then we plotted the straight line of calibration from the maxima of fluorescence corresponding to each lowest level depending on the concentration of histamine (Graph 5). We obtained a coefficient of correlation of 0.999, close to unity, which explains the accuracy of our measurement. The straight line of calibration is characterized by a very high slope (21.15 × 107 ) and a very vast calibration domain.

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A. Tine and S.E. Douabalé pH = 11,48; [OPA] = 10–4 M; [Histamine] = 0,1.10–4 M Kinetics = f(Heating Time) 30 Heating Time (a): 15 s (b): 30 s (c): 60 s

25

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5 Volt. 250 Fact. 1

0 0

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120 180 240 300 360 420 480 540 600 Time (s)

Graph 3 Effect of heating on kinetic formation of OPA–histamine complex in basic medium pH 11.5 [OPA] = 10−4 M [histamine] = 0.1 × 10−4 M

OPA 10–5M/NaCl 0,1M/NaOH 10–3M/water; If = f([histamine]) 200 (7) 175

(6)

150

(5)

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[Histamine]*107 (M) 8.76 (7) (6) 7.41 6.02 (5) (4) 4.58 3.10 (3) (2) 1.57 (1) 0

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Graph 4 Combined effects of temperature and shaking on formation kinetics of OPA–histamine complex


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OPA 10–5M/NaCl 0,1M/NaOH 10–3M/water; If = f([histamine]) 200 175 150

If

125 100 R = 0,99939 75 slope = 21,15.107 50 Volt. 350 Fact. 5

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Graph 5 Calibration of histamine in presence of [OPA] = 10−5 M, [NaCl] = [NaOH] = 10−3 M, from formation kinetics of OPA–histamine complex at 50◦ C during 3 min

We notice that in the reactional medium, we have added NaCl to be in compliance with a possible dosage of histamine extracted from a sea product for instance. Therefore, there is ground for discussing the effect of NaCl on the fluorescence of the complex.

3.1.3 The Effect of NaCl Concentration on Fluorescence of OPA–Histamine Complex in Basic Medium Histamine is extracted from fish by hydrochloric acid (HCl 0.5 N). Let us bear in mind that histamine does not produce any emission. However, the complex OPA– histamine that occurs in the basic medium is fluorescent. As a consequence, the histamine sample to be dosed will contain NaCl by neutralizing hydrochloric acid with soda before being in alkaline medium. Therefore, we have found it necessary to examine the effect of NaCl on the intensity of fluorescence of OPA–histamine complex in an alkaline medium. We then determined the variation of the fluorescent intensity of the complex according to the NaCl concentration. The choice of the interval of concentration is justified by the concentration of NaCl obtained after neutralizing with Lerke and Bell method. The analysis of chemical species gives 0.2 M concentration for NaCl in the reaction medium at alkaline pH. Graph 6 shows that the concentration of NaCl would not have a significant effect on the intensity of fluorescence of the complex, at least in this alkaline medium between 0 and 0.6 M of soda.

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A. Tine and S.E. Douabalé [OPA] = [Histamine] = 10–4 M; pH = 11,48 160 140 120

If

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Volt. 250 Fact. 1

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Graph 6 The effect of NaCl concentration on fluorescence of OPA–histamine complex in basic medium

3.1.4 The Effect of pH on Fluorescence of OPA–Histamine Complex in Basic and Acid Medium Effect of HCl on Fluorescence of the Complex The effect of hydrochloric acid on the spectrum of emission of fluorescence of OPA– histamine complex in basic medium is shown in Graph 14. We can see a lowering of the peak as basicity decreases. As soon as there is an excess of acid, the peak vanishes totally. This proves that in acid medium, OPA–histamine complex cannot be detected through fluorescence, at least not around 430 nm at a low voltage of 200 V (Graphs 7 and 8). Thus, in acid medium, fluorescence of the complex is totally inhibited. Actually, the intensity of fluorescence of the peak decreases rapidly depending on the quantity of acid added. However, when we increase the voltage to 350 V, we can detect two different types of complexes. The first one emits at 430 nm and the second one at 510 nm (Graphs 8a and b). By maintaining the emission wavelength at 430 nm, we recorded spectra of excitation and detected two peaks. The first one at 346 nm, corresponding to the excitation of the emission peak at 430 nm, and the other one at 386 nm, corresponding to the excitation of the emission of the complex whose peak is at 510 nm (Graph 9a and b). We have also drawn the spectrum of emission by fishing the excitation at 386 nm (Graph 10a). We observed the apparition of an isosbestic point around 510 nm. This proves the existence of a balance between both the complexes. By continuing acidification, we notice that the intensity of both peaks decreases at the same time


205

Graph 7 Lowering of emission intensity of fluorescence of OPA–histamine complex as soda is neutralized by hydrochloric acid [OPA] = [histamine] = 10−4 M

when we obtain neutrality between acid and soda (Graph 10b). This then proves that hydrochloric acid inhibits the fluorescence of both complexes. However, to be in the same conditions as in Lerke and Bell method, we optimized the effect of the OPA concentration. Actually, the conditions fixed by Lerke and Bell method were chosen for an OPA concentration around 10−3 M. If at low OPA concentrations (10−4 M) no fluorescence has been noticed, we notice a slight increase of fluorescence of the complex at 430 nm depending on the concentration of OPA that stabilizes around 10−3 M (Graph 11). However, when studying the intensity of fluorescence in acid medium in relation to pH, we see that the maximum intensity of fluorescence corresponds with a very narrow interval of pH (Graph 12). This low interval can cause fatal errors if we are not working in buffer medium. Thus, we plotted the calibration graph of the intensity of fluorescence in relation to the concentration of histamine in optimal conditions of OPA in acid medium (Graph 13). We have found a coefficient of correlation of 0.994 and a slope of 2 × 107 . If that graph is accurate, the calibration domain is very low. This low calibration domain should not give us conclusive results for the determination of low levels of histamine from sea products; all the more as we use Lerke and Bell method to determine the level of histamine in the food left by intoxicated patients. This low domain of calibration allows us to work in the alkaline medium.

NaOH Effect on Fluorescence of the Complex In studying pH effect, we observed that the intensity of fluorescence is maximal around a pH between 11 and 12, precisely to pH equaling 11.48 (Graph 14). In taking into account all the previous optimal conditions, we plotted a calibration curve from fluorescent intensity corresponding to the levels of the complex versus histamine concentration (Graph 15). We obtained a very high slope (2.38 × 108 ) with a coefficient of correlation of 0.999 proving the precision of our measurements.

206

A. Tine and S.E. Douabalé [HCl]

200 180

(1)

Relative Fluorescence

160 140

(2)

430

120

[NaOH]

[NaCl]

(1)

0 ; 2,27x10–4 M ; 0,097 M

(2)

0 ; 1,50x10–4 M ; 0,095 M

(3)

0 ; 0,75x10–4 M ; 0,094 M

(4)

0 ; 0,37x10–4 M ; 0,092 M

(5)

0 ;

0

; 0,090 M

100 (3) 80 60

a

(4)

40

Volt. 350 Fact. 1

20 (5) 0 400

450

500

550

Wavelength (nm)

600

λexc= 346 nm

B (4)


40

[HCl]

430 30

(4) (5)

20

10

[NaOH]

[NaCl]

; 0,37x10–4 M ; 0,092M ; 0 ; 0,090M

0 0

(6) 0,37x10–4 M ;

0

; 0,093M

(5)

(7) 1,46x10–4 M ;

0

; 0,093M

(6) (7)

(8) 4,26x10–4 M ;

0

; 0,093M

(8)

510

b 0

Volt. 350 Fact. 1 400

450

500

Wavelength (nm)

550

600 λexc= 346 nm

Graph 8 a and b Spectra of emission of OPA–histamine complex at high voltage (350 V)

Comparison of Experimental Results Obtained in Acid and Basic Medium In comparing our obtained results in alkaline medium (our own method) with those obtained in acid medium (Lerke and Bell or OAAC method), we noticed that the calibration domain is ten times higher in alkaline medium, with a coefficient of correlation much closer to unity (Graphs 16 and 17 and Table 1).


207

346 200

(1) (2)

(1) (2) (3) (4) (5) (6) (7) (8)

Relative Excitation

160 (3) 120

(4) (5)

80

[HCl] 0 ; 0 ; 0 ; 0 ; 0 ; 0 ; 0 ; 0 ;

[NaOH] 7,14x10–4 M 6,3x10–4 M 4,65x10–4 M 3,85x10–4 M 3,05x10–4 M 1,5x10–4 M 0,37x10–4 M 0

(6)

40

250

300

[NaCl] 0,099M 0,099M 0,097M 0,097M 0,096M 0,095M 0,094M 0,093M

a

(7)

Volt. 350 Fact. 1

(8) 0 200

; ; ; ; ; ; ; ;

350

400 λexc= 510 nm

Wavelength (nm)

346 386 (7)

20

Relative Excitation

(8) (9) 15

b

(10) (11) [HCl] (7) 0 (8) 0 (9) 0,74x10–4 M (10) 2,88x10–4 M (11) 4,26x10–4 M

10

5

0 200

250

300 350 Wavelength (nm)

400

; ; ; ; ;

[NaOH] 0,37x10–4 M 0 0 0 0

; ; ; ; ;

[NaCl] 0,094 M 0,093 M 0,093 M 0,091 M 0,090 M

Volt. 350 Fact. 1

λexc = 510 nm

Graph 9 a and b Spectra of emission of OPA–histamine complex at high voltage (350 V)

The analysis results from straight line calibration are then more precise in alkaline medium than in acid one. The statistic results showed precisely that our method was improved in comparison with Lerke and Bell one. In fact, if the detection limit is 0.25 ng/mL for the method in basic medium, it is only 2.5 ng/mL when proceeding by Lerke and Bell method. Also, the recovery is between 89 and 114% for our method. It is between 17 and 217% for the other one and is consequently a very wide recovery. So, the statistic results showed that the analysis method in basic medium is precisely better than the one in acid medium.

208

A. Tine and S.E. Douabalé (2) 200 (3)

(4)

a (1)


160

(1) (2) (3) (4) (5) (6) (7)

120 430 80 (5) 40

[HCl] 0 0 0 0 0 0 0

; ; ; ; ; ; ;

[NaOH] 8x10–4 M 6,30x10–4 M 4,65x10–4 M 3,05x10–4 M 1,50x10–4 M 0,75x10–4 M 0,37x10–4 M

; ; ; ; ; ; ;

(6)

400

Volt. 350 Fact. 1

510

(7)

0

450

500

550

Wavelength (nm)

600

650

λexc= 386 nm

b

45 (7)

40

(8) (9)

35 Relative Fluorescence

[NaCl] 0,1 M 0,1 M 0,096 M 0,096 M 0,095 M 0,094 M 0,094 M

[HCl] (7) 0 (8) 0 (9) 1,46x10–4 M (10) 4,26x10–4 M

30 430

25

(10)

; ; ; ;

[NaOH] [NaCl] 0,37x10–4 M ; 0,094M 0 ; 0,093M 0 ; 0,094M 0 ; 0,090M

20 15

(7)

10

510

(8)

5

Volt. 350 Fact. 1

0 400

450

500

550

Wavelength (nm)

600

650

λexc= 386 nm

Graph 10 a Spectrum of emission of the complex OPA–histamine at high voltage (350 V) b Spectrum of emission of the complex OPA–histamine at high voltage (350 V)

If our results obtained in basic medium are much more convincing only in acid medium, we notice that pH domain corresponding to the maximum intensity of fluorescence is very narrow (Graph 7), hence, the usefulness to work in a pH buffered solution equaling 11.48.


209

pH = 0,99; Heating and shaking

60 50

If

40 30 [Histamine] = 10–4 M

20 10

Volt. 250 Fact. 1

0 0

2

4

6

8

1

0 1

2

4 [OPA]x10 (M)

Graph 11 Effect of OPA concentration on fluorescence of OPA–histamine complex in acid medium

70 [Histamine]=1,1.10–4 M

60

[OPA]=1,46.10–3 M

50

If

40 30 20 10

Volt. 250 Fact. 1

0 0

1

2

3

4

5

6

7

8

9

1

1 0

pH

Graph 12 Effect of pH on fluorescence of OPA–histamine complex

1

1 2

1 3

4

210

A. Tine and S.E. Douabalé 60 50 pH = 0,82

If

40 30 20

R = 0,99399 Slope = 0,20978x108

10 Volt: 400 Fact: 10

0 0

5

10

15

20

25

[Histamine] x 108 (M)

Graph 13 Calibration of the complex OPA–histamine in acid medium according to Lerke and Bell method

Graph 14 pH effect on fluorescence of OPA–histamine complex ([OPA] = 1.1 × 10−4 M and [histamine] = 1.1 × 10−4 M)


211

[OPA] = 10–5 M, to 50°C 60 50 R = 0,99965 slope= 2,38x108

If

40 30

pH = 11,48 20 10

Volt: 400 Fact: 10

0 0

5

10 15 [Histamine] x 108 (M)

20

25

Graph 15 Histamine calibration in presence of [OPA] = 10−5 M in NaOH medium at pH 11.48 (a):[OPA] = 10–5 M –3 (b):[OPA] = 1,46.10 M 60 (a)

50

pH = 11,48

R = 0,99965 Slope = 2,38x108

If

40 30

pH = 0,82 20 (b) 10

R = 0,99399 Slope = 0,20978x108 Volt: 400 Fact: 10

0 0

5

10 15 [Histamine] x 108 (M)

20

25

Graph 16 a and b Calibration of OPA–histamine complex in basic medium (a) and in acid one (b)

3.2 Optimization with Phosphate Buffer 3.2.1 Preparation of Phosphate Buffer Solution Neutralization of eluat acid for the condensation reaction is a reaction between strong base and strong acid (NaOH and HCl). Such reaction always causes a sudden

212

A. Tine and S.E. Douabalé Relative domain of calibration

Graph 17 Comparative diagram of domains of calibration of OPA–histamine complex in acid (pH 0.83) and basic (pH 11.48) mediums

100 80 60 40 20 0

pH 0,82

pH 11,48

Table 1 Comparison of statistic results obtained in two mediums

Method Our method (basic medium) Lerke and Bell method (acid medium)

Relative domain of calibration in If

Limit of détection (ng/mL)

Recovery (%)

Coefficients of correlation

1–100

0.25

90–114

0.9996

0–10

2.5

17–217

0.9940

lapse of pH next to the equivalent point between acid and base. Then it is difficult to be in optimal conditions of pH in neutralizing only eluat acidity by soda. That is why we used buffer solution. That solution is obtained from phosphoric acid. We know that phosphoric acid corresponds to three equilibria which are the following: Equilibrium 1: H3 PO4 + H2 O H2 PO4 − + H3 O+ pKa1 = 2.15 Equilibrium 2: H2 PO4 − + H2 O HPO4 2− + H3 O+ pKa2 = 7.20 Equilibrium 3: HPO4 2− + H2 O PO4 3− + H3 O+ pKa3 = 12.38 In neutralizing phosphoric acid by soda, we can have buffer solution corresponding to the equilibrium 3. In fact, this equilibrium allows us to obtain buffer solution covering pH equal to 11.5. The extent of this buffer domain covers an interval containing pH value of work (11.5). Then we have the following formula for weak concentrations of PO4 3− and HPO4 2− . pH = pKa + log

[PO4 3− ] [HPO4 2− ]

On the other hand, we know that hydrochloric acid concentration of eluat is 0.2 N. To minimize the contribution of this acid in buffer solution, we must use strong concentrations of PO4 3− and HPO4 2− . In these conditions, pH of buffer solution responds to the relation


pH = pKa + log

213

aPO4 3− aHPO4 2−

If we know that the activity a = yc, then the activity of basic type can be written as aBase = γBase · PO4 3− and the acid type one is written as aAcid = γAcid · PO4 3− Therefore relation (2) can be written as pH = pKa + log

aBase [PO4 3− ] + log aAcid [HPO4 2− ]

The report of basic form activities by acid form being known and equal to aBase = 0.447 aAcid pH formula can be written as

pH = 12.38 + log 0.447 + log

[PO4 3− ] [HPO4 2− ]

at 25◦ C (4)

And after calculation, the final formula can be written as

pH = 12.03 + log

[PO4 3− ] [HPO]4 2−

Concerned with having pH equal to 11.48 at the moment of dosage, we prepared buffer solution of pH 11.5 (1 M). For that, we mixed 330 mL of phosphoric acid solution 1 M to 670 mL of soda solution 1 M. Thanks to pH meter and stirring rod we adjusted pH to 11. 5 in using soda solution 1 M. The experience showed that in mixing 2 mL of eluat histamine contained in HCl 2.0 N solution with 5 mL of buffer solution, we obtained a mixture whose pH must be around 11.48. It is in this solution in the presence of well-defined concentration of OPA that we made the condensation reaction of histamine to obtain OPA–histamine complex.

214


3.2.2 Standard Straight line Calibration and Calculation of Detection Limits and Quantification We found linear correlation between the intensity of fluorescence corresponding to the maxima of complex kinetics and histamine concentration in considering all optimal conditions. Straight line calibration had a slope of 18.717 × 107 with coefficient of correlation of 0.998, close to unity (Graph 18). Then, we can use basic buffer solution instead of NaOH solution for OPA– histamine complex formation. The only disadvantage is to have a viscous solution because of the concentration of buffer solution (117). That is the reason why the detection limit is weak in relation to the one obtained with NaOH. Further studies are being done to diminish the viscosity of buffer solution. From this straight line of calibration, we found a few statistical results: limit of detection (LD) and limit of quantification (LQ). For that, we determined standard deviation corresponding to successive attempts of the intensity of fluorescence for solvent (see Table 4). We found standard deviation σ = 1.683124. From that value, we determined limit of detection (LD) and limit of quantification (LQ). The limit of detection (LD) is calculated as being the concentration obtained from the intensity of fluorescence corresponding up to three times the standard deviation on the line of calibration, with LD = 0. 05 × 10−7 mol/L corresponding to 555 ng/L or 0.55 ng/mL This value is higher than 0.25 ng/mL obtained in acid medium because of viscosity. The limit of quantification (LQ) is defined as being the concentration corresponding to the intensity of fluorescence equal to ten times the standard deviation (10σ ).

OPA 10–5M/ phosphate buffer pH 11,5 / HCl; If = f([histamine]) 90 80 70 slope = 18,71.107 r = 0,99787

60 If

50 40 30 20 10

Volt. 400 Fact. 5

0 0,0

0,5

1,0

1,5

2,0

2,5

3,0

3,5

4,0

[histamine] x 107 (M)

Graph 18 Standard straight line calibration of OPA–histamine complex in presence of OPA and phosphate buffer pH 11.5


LQ =

10σ 10β

215

1/α = 1.5 ng/mL

Detection and quantification limits, with very low rate of 1 ng/mL, are among the best methods of histamine dosage that have been published till date. Fortified by these results, we found a protocol for the extraction and determination of histamine all the more as the cover level is contained between 96.87 and 110.83%. In fact, this method gives very reliable results and is less expensive. This protocol would be very useful for an industrial utilization, which is easy to realize with few reagents.

3.2.3 Determination of the Recovery by Our Experimental Method To determine the cover level of dosage, we realized measures of fluorescence of known concentration of histamine standard equaling 7.9 × 10−8 . For that, concentration has been made at 50◦ C in phosphate buffer solution in the presence of OPA 10−5 . From the intensity of fluorescence obtained, we could determine the corresponding concentration with standard straight line calibration. For that very solution, we made repetitive measures after having previously emptied out the cell. Let the recovery be between 96.87 and 10.83%. This shows that the used method is very good, much better than the official one where cover level varies between 17 and 217%.

3.3 Applications: Determination of Histamine Rate in Fish To illustrate our protocol for the determination of histamine rate contained in dried fish sample, The experiment was made in two stages. The first stage consisted in determining histamine rate right after purchase (sample 1), and the second stage a few days after keeping the sample in freezer (sample 2).

3.3.1 Determination of Histamine Rate in Sample 1 We extracted histamine from this sample in conformity with the protocol described above. Thus, we compared excitation and emission spectra of histamine extracted from sample with those coming from standard in the very solvent (Graph 19). We obtained a difference in nonexistent wavelengths for each type of spectrum, confirming then the presence of histamine in our sample. Thus, we determined the histamine rate of the sample from standard additional straight line and histamine calibration curve (Graph 20). Coefficients of correlation corresponding to two straight lines are around 0.999, close to unity, which indicates the good precision of our measures. A good extrapolation of the y-axis at the origin of the standard additional line on the calibration line gives us histamine concentration contained in the reactor. With the concentration of 1.39 × 10−7 M found, the calculation gives histamine rate equal to 54 ppm when using the protocol formula.

216

A. Tine and S.E. Douabalé 70 2

1

60 50

Emissions

Excitations

If

40 b

30

a

20 10 0 200

250

300

350 400 wave length λ (nm)

450

500

Graph 19 Excitation and emission spectra of OPA–histamine complex: authentic histamine (a and b), extracted from fish sample (1, 2): in presence of [OPA] = 10−5 M and phosphate buffer pH 11.5

3.3.2 Determination of Histamine Rate in Sample 2 With the same process, we realized standard calibration line (in relation to the standard histamine concentration) and another additional standard (Graph 21). We found also coefficients of correlation to be around 0.999 for the straight lines, indicating the precision of our measures. The use of these curves allowed us to extrapolate and deduce histamine concentration equaling 1. 4 × 10−7 M in the reactor, which always corresponds to 55 ppm rate when using the formula of the protocol. OPA 10–5M/ phosphate buffer pH 11,5 / HCl; If = f([histamine]) 90 80

slope = 21,37.107 r = 0,99858

70 60 If

50 40

slope = 20,11.107 r = 0,99861

30 20

Volt. 400 Fact. 5

10 0 0,0

0,5

1,0 1,5 [histamine] x 107 (M)

2,0

2,5

Graph 20 Standard addition (above) and calibration (below) of histamine in presence of [OPA] = 10−5 M and phosphate buffer pH 11.5


217

OPA 10–5M/ phosphate buffer pH 11,5 / HCl; If = f([histamine]) 90 80

slope = 21,37.107 r = 0,99858

70 60 50 If

slope = 20,11.107 r = 0,99938

40 30 20 10

Volt. 400 Fact. 5

0 0,0

0,5

1,0

1,5

2,0

2,5

3,0

3,5

4,0

[histamine] x 107 (M)

Graph 21 Standard addition (above) and calibration (below) of histamine in presence of [OPA] = 10−5 M and phosphate buffer pH 11.5: verification of the reproducibility

This 54 ppm histamine rate is identical to the one obtained at the first measurement of sample 1. Then, this result proves there is no evolution of histamine rate when we preserved the cold sample. This result proves also the reproducibility of our method.

4 General Conclusion Our present work composed of four parts leads to the following results: The first part made up of our preliminary studies in absorption and in fluorescence allowed us to confirm the stoichiometry 1:1 of the reaction between orthophthalaldehyde (OPA) and histamine. The calibration curves obtained in absorption and in fluorescence allow us to determine histamine rate corresponding to the concentrations of 10−5 M and above. To have weak detection limits, we made curves of calibration between the intensity of fluorescence corresponding to the maxima of kinetics of complex formation and the histamine concentration. After having searched optimal conditions (optimal temperature, optimal pH 11.48), we obtained stability of OPA–histamine complex, and correlations corresponding to the obtained lines of calibration were 0.99, close to unity. In that very basic medium, we had a high domain of calibration. Detection limits were 0. 99 ng/mL, much more reliable than at the time of our preliminary work. So, we showed that the determination of histamine rate was more probing and more reliable in basic medium than in acid one (Lerke and Bell conditions or Official Association of Analytical Chemistry). In fact, we showed the existence of two types of complex in acid medium and a very weak domain of calibration.

218


However, the study of pH on fluorescence of the complex showed us a very narrow interval of pH in basic medium. Then, we had to do the same work in pH 11.48 buffer solution. In that medium, considering optimal conditions, we also found linear correlations between intensities of fluorescence corresponding to the maxima of kinetic complex and histamine concentration. Coefficients of correlation around 0.999 show the precision of our measures. The comparison of statistical results of our measures with those obtained by Lerke and Bell method showed that our method was far more reliable, more precise, and more reproducible than official methods. The difficulties met for the determination of histamine rate by official method are then solved and our protocol can be valuably used, in a more precise and concise manner, for the determination of histamine rate as well in industrial mediums as in hospitals.

References 1. W. Lorenz, E. Neugebauer; Fluorometric assays, curent techniques of histamine determination; 2 (1990) 9–30. 2. P.B. Henderson; Case of poisoning from the bonito (Scomber pelamis); Edinb. Med. J., 34 (1830) 317–318. 3. C. Den Brinker, C. Rayher, M. Kerr; Investigation of Biogenic Amines in Fermented Fish and Fish Products; Ed. 1, Public Health Division, Victorian Government Department of Human Services (2002). 4. S. Taylor, Histamine food poisoning: toxicity and clinical aspects; Crit. Rev. Toxicol., (1986) 91–128. 5. H.H. Huss; La qualité et son évolution dans le poisson frais; document technique sur les pêches – 348, Organisation des Nations Unies pour l’Alimentation et l’Agriculture (FAO); Rome, 1999. http://www.fao.org/documents/show_cdr.asp?url_file=/DOCREP/003/ V7180F/V7180F00.HTM 6. S.H. Kim, K.G. Field, M.T. Morrissey, R.J. Price, C.I. Wie, A.N. Haejung; Source and identification of histamine-producing bacteria from fresh and temperature-abused albacore; J. Food Prot., 64, No 7 (2001) 1035–1044. 7. E.I. Lopez-Sabater, J.J. Rodriguez-Jerez, A.X. Roig-Sagues, M.A.T. Mora-Ventura; J. Food Prot., 57 (1994) 318. 8. K.V. Basavakumar, I. Karunasagar; Indian J. Microbiol., 32 (1992) 75. 9. P.K. Vijayan, P.K. Surendran, K.K. Balachandran; (Symp. on Tropical Marine Living Resources, Cochin, India, Jan. 12–16, 1988), J. Mar. Biol. Assoc. India, 31 (1989) 202. 10. S.L. Taylor, J.E. Stratton, J.A. Nordlee; J. Toxicol., 27 (1989) 225. 11. L. Lehane, J. Olley; Histamine fish poisoning revisited; Int. J. Food Microbiol., 58, No. 1–2 (2000) 1–37. 12. C.M. Osborne, P.J. Bremer; Application of the Bigelow (z-value) model and histamine detection to determine the time and temperature required to eliminate morganella morganii from seafood; J. Food Prot., 63, No. 2 (2000) 277–280. 13. T. Hibit, M. Senda; Enzymatic assay of histamine by amperometric detection of H2 O2 with a peroxidase-based sensor; Biosci. Biotechnol. Biochem., 64, No. 9 (2000) 1963–1966. 14. P.A. Lerke, M.N. Poscuna, H.B. Chin; J. Food Sci., 48 (1983) 155. 15. W. Lorenz, E. Neugebauer, B. Uvnäs, M.A. Beaven, M. Ennis, G. Granerus, J.P. Green, J.J. Keyzer, P.T. Mc Bride, P.F. Mannaioni, F.L. Pearce, J. Watkins; K. MUNICH Consensus Development Conference On Histamine Determination, Current techniques of histamine determination, 2 (1990) 81. 16. P.L. Rogers, W. Staruszkiewicz; J. AOAC Int., 80, No. 3 (1997) 591–602.

Spectroscopy of DNA–Actinomycin Complexes Nikolai Vekshin

Abstract Complexes of actinomycin D (AMD) and 7-amino-actinomycin D (7AAMD) with model hairpin oligonucleotide HP1 and various types of DNA in aqueous solutions were investigated by steady-state, polarized, time-resolved and stopped-flow fluorimetry. Prompt non-stacking binding of the actinomycins inside HP1 was observed. No energy transfer from nucleotides to 7AAMD in the complex was detected, most likely because of the absence of stacking intercalation. The data confirm the non-stacking binding of 7AAMD or AMD to HP1 and DNA in aqueous solutions. Strong competition between AMD and 7AAMD for a cavity in HP1 was observed. The decrease in the 7AAMD emission after addition of DNA to the 7AAMD/HP1 complex indicates that actinomycins can be redistributed from HP1 to DNA. The biological significance of this finding is that hairpin oligonucleotides can serve as molecular carriers, transferring actinomycins to DNA. Addition of 7AAMD to the caffeine clusters leads to its sorption on the cluster surface. Photo-excitation of 7AAMD leads to its de-sorption from the surface into the aqueous phase and emission of a quantum. Fluorescence of 7AAMD in the presence of caffeine clusters is quenched by dinitrophenol more weakly than without clusters due to decreased stereo-availability of the antibiotic to the quencher. Addition of 7AAMD-caffeine complexes to DNA leads to a long-wavelength shift in the excitation spectrum and an increase in the fluorescence intensity along with a shift of the fluorescence spectrum to the short-wavelength area. This fact reflects redistribution of the antibiotic from the caffeine surface to the hydrophobic areas inside DNA.

1 Introduction Actinomycins are widely applied because of their antibiotic and anti-tumor activity. Actinomycins are among the most effective antibacterial and antitumor drugs, N. Vekshin (B) Institute of Cell Biophysics, Pushchino, Moscow region, 142290, Russia e-mail: [email protected]


219

220

N. Vekshin

which are widely used for the chemotherapy of sarcomas, lymphomas, melanomas, rhabdomyosarcomas, genital malignancies, etc. [1–4] Moreover, actinomycins decelerate ageing by prolonging the life cycle [5]. Well known among actinomycins is actinomycin D (AMD), consisting of 2-aminophenoxazin-3-one chromophore and two cyclic pentapeptide lactones (Fig. 1).

Fig. 1 Actinomycin D. Here R is N in the case of AMD and R is NN2 in the case of 7AAMD

The biological activity of AMD is attributed to its ability to bind to DNA and thus to inhibit the RNA polymerase reaction and protein biosynthesis [6–8]. It is known that AMD displays cytotoxicity to cancer cells at very low concentrations, but its clinical use is limited by side toxic effects and a bad penetration into cells. The widespread application of AMD requires determination of the molecular mechanism of the AMD/DNA complex formation. It was generally accepted that AMD intercalates into duplex DNA, preferring GpC sites [6, 9]. An alternative binding mechanism at non-GpC (i.e., ApT/TpA) sites is also possible [10]. In the solid phase, the stacking intercalation of phenoxazone chromophore between nucleotides was shown in crystals by X-ray analysis [9] and in films by fluorescence energy transfer [11]. In aqueous solutions, actinomycins can bind to DNA in two ways: (a) stacking-like intercalation, at high concentrations and (b) binding in non-stacking positions, at low concentrations [11, 12]. The highest binding constant for actinomycins was observed for hairpin-like sites or loops [11–15]. Instead of AMD, a fluorescent analog, 7-amino-actinomycin D (7AAMD), is usually used at low concentrations to study the properties of actinomycin/DNA complexes [11–17]. Fluorescence is much more sensitive than absorption. That is why 7AAMD instead of AMD is used. The excitation spectrum of 7AAMD bound to DNA exhibits a long-wavelength (“red”) shift in comparison to free 7AAMD in water, whereas the emission spectrum shows a short-wavelength (“blue”) shift [12, 13]. The biggest difference in the

Spectroscopy of DNA–Actinomycin Complexes

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fluorescence intensity of 7AAMD in DNA vs. 7AAMD in water was found on the “red edge” of excitation and “blue edge” of emission [12]. It has been shown that 7AAMD can form a compact complex with hairpin (single-stranded) oligonucleotides with high binding constants [14, 15, 17]. In particular, 7AAMD binds strongly to the 8.6 kDa hairpin HP1 d(5 AAAAAAATAGTTTTAAATATTTTTTT-3 ). The binding is accompanied by an increase by more than an order of magnitude in the fluorescence intensity due to changes in the lifetime and spectral shifts [12, 14–16]. The sequence of HP1 is also compatible with the formation of a homo-dimer, a mismatched duplex. In principle, hairpins can form both types of structures (they can be distinguished by native PAGE). In [12–17] it was shown that at very low (physiological) concentrations (1 μM and lower) actinomycins can bind to hairpin oligonucleotides and to hairpin-like sites or loops in DNA. The highest binding constant was ∼107 M−1 [17]. However, the reason for such strong interaction was unknown. It was also unclear why the binding takes place at a non-stacking position [11, 12]. It is important to determine the features of AMD and 7AAMD binding to HP1 and DNA at low concentrations. Also, it would be interesting to examine the possibility that actinomycins can be redistributed from HP1 to DNA. Such redistribution can be used in medicine for effective transport of antibiotics to DNA. The advantage of the present investigation over the previous ones is the combination of methods: steady-state, polarized, phase-modulating, and stopped-flow fluorimetry. Also, a new approach for determining the location and orientation of 7AAMD in HP1 and DNA by means of the efficiency of energy transfer (using excitation spectra) was applied.

2 Emission of 7AAMD in Oligonucleotides and DNA The excitation spectrum of 7AAMD bound to DNA exhibits a red shift in comparison to free 7AAMD in water, whereas the emission spectrum shows a “blue” shift (Figs. 2 and 3). 7AAMD emits mainly from untwisted sites of DNA. That is why the highest intensity of fluorescence is observed in the case of denaturated DNA (Figs. 2 and 3). 7AAMD can serve like a good probe on untwisting, denaturation, and fragmentation of DNA. In the case of the 7AAMD/HP1 complex, the intensity of the 7AAMD fluorescence is very high (Table 1). The reasons for this are not only increases in the lifetime and quantum yield of 7AAMD after binding to the hairpin but also a red shift in the 7AAMD excitation band [11, 12]. Besides, a blue shift of the emission band takes place [11–16]. The fluorescence intensity of 7AAMD depends on the type of DNA to which it is bound. Table 1 presents data for lambda, calf thymus, and salmon sperm DNA. The fluorescence of 7AAMD reaches a plateau in 5 min after addition of

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Fig. 2 Emission spectra of 7AAMD (2 μM) in the presence of native thymus DNA (1), denaturated thymus DNA (2), and in the buffer (10 mM cacodylate, 150 mM NaCl, pH 7) (3).λex = 570 nm

Fig. 3 Excitation spectra of 7AAMD (2 μM) in the presence of native thymus DNA (1), denaturated thymus DNA (2), and in the buffer (10 mM cacodylate, 150 mM NaCl, pH 7) (3).λem = 620 nm

Table 1 Fluorescence intensity of 7AAMD in the presence of various nucleic acids. The 7AAMD concentration was 1 μM. The buffer was 20 mM Tris-HCl (pH 7.5). The concentration of all types of DNA, yeast t-RNA, HP1, and HP1A was 0.014 mg/ml. Excitation was at 570 nm and emission was at 610 nm. The presented intensity values are the averages of five measurements Sample

Intensity (rel. un.)

HP1 + 7AAMD HP1 + 7AAMD + 10 μM AMD HP1A + 7AAMD Salmon sperm DNA + 7AAMD Calf thymus DNA + 7AAMD Lambda DNA + 7AAMD Lambda DNA + 7AAMD + 80 μM AMD Yeast t-RNA + 7AAMD Buffer + 7AAMD

2610 160 250 740 560 505 205 200 130


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the DNA aliquot. The highest 7AAMD emission intensity was observed for salmon sperm DNA, indicating that this DNA preparation contains many loops and breaks. Relatively short native λDNA has a minimum of hairpin-like sites and breaks. The hairpin-like sites, loops, and breaks appear in the long double helix due to mechanical damage and thermal fluctuations. Replacing guanine in HP1 with adenine (HP1A) leads to an abrupt decrease in the binding affinity for actinomycins (Table 1). The presence of guanine in HP1 is essential for formation of the hairpin-like structure. This is why the presence of guanine nucleotide in HP1 is essential for 7AAMD binding. It is important to emphasize that there is no “specific interaction” between guanine and 7AAMD. Guanine itself is not an obligatory basis for binding of actinomycins [10–12, 18]. In fact, 7AAMD and AMD can effectively interact with oligonucleotides independently of the presence or absence of a GpC sequence [10, 18]. The lifetime of 7AAMD in the complex with HP1 is 1.7 ns. In the case of complexes with DNA, the 7AAMD lifetime is similar: 1.5–2.1 ns for all types of DNA used. However, the fluorescence intensity of 7AAMD in HP1 was four to five times higher than that in DNA (under the same conditions; Table 1). This means that the population of 7AAMD molecules in DNA is quenched. It can be suggested that such static quenching of 7AAMD takes place in the double helix by neighboring nucleotides (it is well known that nucleotides are strong quenchers, since they have very intense vibration modes). In the presence of yeast t-RNA, the fluorescence intensity of 7AAMD was almost the same as in the case of free 7AAMD. Thus, 7AAMD is unable to bind to yeast tRNA, since this rigid poly-nucleotide has too small a hairpin cavity. This means that the existence of hairpin-like sites itself is not sufficient for effective binding. Other conditions are the size of a cavity or loop and flexibility of oligonucleotide tails. The free ends of the HP1 hairpin exhibit a high flexibility. Actinomycins can easily penetrate into open dynamically arising cavities inside a flexible hairpin structure. In the case of DNA, long loops and big hairpin-like or untwisted sites are necessary for the fast actinomycin binding. The 7AAMD fluorescence intensity depends on the presence of hydrophobic cavities in the DNA structure.

3 Negligible Energy Transfer from DNA to 7AAMD Energy transfer from photo-excited nucleotides to a dye is one of the main features, indicating stacking intercalation in complexes [19, 20]. The stacking position of a dye between nucleotide chromophores in DNA provides a favorable condition for effective energy transfer. Energy transfer can be detected, for instance, as the appearance of the nucleotide band in the excitation spectrum of the emitting dye [19–21]. Total efficiency (Qda ) of energy transfer from nucleotides to dye molecules was calculated with the equation [19, 21]

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[d]Qda = Fd εa [a]/Fa εd [d] where εd is the extinction coefficient of the donor (in the case of DNA, it is the average ε of the nucleotides at 260 nm), εa is the extinction coefficient of the acceptor at its absorption maximum, [d] is the concentration of the donor, [a] is the concentration of the acceptor (the average concentration of nucleotides), Fd is the intensity of the donor band in the excitation spectrum (Fd equals the difference between the total intensity at 260 nm and the contribution from the acceptor at this wavelength), and Fa is the intensity of the acceptor band (Fa equals the total intensity at the acceptor excitation maximum minus the contribution from nucleotides at the same wavelength). The number of nucleotides (N), donating the energy per one acceptor molecule, was estimated according to the equation N = Fd εa /Fa εd The intercalating ability of 7AAMD in comparison with those of ethidium bromide (EB), a typical intercalating dye, and Hoechst 33342, a typical nonintercalating dye, was estimated. EB induces denaturation and hyperchromism of native DNA [19]. That is why EB is a strong cancer-inducing agent. On the contrary, Hoechst (for instance, H33342 and H33258) induces DNA condensation [22]. It is interesting that Hoechst is known as an anticancer drug [22]. In the UV region, EB exhibits an intensive absorption band at 284 nm, Hoechst one at 340 nm, and 7AAMD one at 260 nm. The absorption band of EB overlaps to some degree with the nucleotide emission spectrum. The fluorescence quantum yield of poly-nucleotides in aqueous solutions at room temperatures is very low [19, 20]. Thus, Dexter’s exchange energy transfer [20] from nucleotides to this dye could be expected only in the case of EB intercalation into DNA. Förster’s dipole– dipole energy transfer [20] can hardly be expected here. The Hoechst absorption band overlaps sufficiently with the nucleotide emission spectrum. Therefore, both types of energy transfer could take place for Hoechst. In the case of 7AAMD, mainly Dexter’s energy transfer can occur. To determine the efficiency of the total energy transfer, it is necessary to determine the fluorescence intensity of a dye, sensitized by nucleotides, and the intensity of a directly excited dye (it is also necessary to determine the nucleotide/dye relationship and absorbance) [19, 20]. The fluorescence intensity of EB increases after its binding to DNA. A strong nucleotide band at 260 nm was detected in the excitation spectrum of EB in DNA [12, 19]. The same observation was made for EB in HP1 [11, 12]. Hence, effective energy transfer takes place, which indicates stacking-like intercalation of EB. The nucleotide band in the EB excitation spectrum appears due to exchange energy transfer to EB from 1–2 neighboring nucleotides [12, 19, 21]. The calculated number of nucleotides (N) donating the excitation energy per one dye molecule in HP1 for the EB/HP1 complex is 1 (Table 2), i.e., one nucleotide donates the energy to one dye molecule. The efficiency (Qda ) of energy transfer from nucleotides to the


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Table 2 Efficiency of energy transfer from nucleotides to dyes. The intensity of the dye excitation band (Fa ) was taken as 100%. Fd is the intensity of the donor nucleotide band in the acceptor dye excitation spectrum, Qda is the maximal total energy transfer efficiency from nucleotides to dye, and N is the number of nucleotides donating the energy to one dye molecule. The nucleotide/dye ratio [d]/[a] was 20. Emission of Hoechst was detected at 460 nm, of EB at 610 nm, and of 7AAMD at 640 nm Complex

Fd (%)

Qda

N

7AAMD/HP1 Hoechst/HP1 Ethidium/HP1 7AAMD/λDNA Hoechst/λDNA Ethidium/λDNA

∼0 0 95 0.03 0 –

∼0 0 0.16 0.003 0 0.15

∼0 0 1.0 0.06 0 1.3

dye for the EB/HP1 complex in aqueous solution is 0.16. Similar N and Qda values were obtained previously for a complex of EB with phage DNA [21]. On the contrary, no nucleotide band at 260 nm was detected in the excitation spectrum of the Hoechst/HP1 complex. The N and Qda values are equal to zero (Table 2). There is no energy transfer in this non-intercalative complex (but a considerable increase in the own Hoechst emission after its binding to HP1 was observed). The absence of energy transfer indicates that the distance between Hoechst and the nucleotide chromophores are large and/or that the orientation is not coplanar. Similarly, no nucleotide band was detected at 260 nm in the excitation spectrum of the 7AAMD/HP1 complex. The N and Qda are approximately equal to zero (Table 2), i.e., there is no effective energy transfer. This fact is a potent proof against stacking of 7AAMD in HP1. Indeed, this fact is direct evidence that the chromophore of 7AAMD is located far from the nucleotide hetero-cycles and/or has another orientation. Since only one 7AAMD molecule can bind to a single HP1 molecule, N and Qda should be concentration independent. In the case of the complex of 7AAMD with DNA, the Qda value was 0.003 and N was 0.06 (Table 2), i.e., only very little energy transfer was detected. The nucleotide band at 260 nm in the excitation spectrum of the 7AAMD/DNA complex was negligible [11, 12]. Energy transfer was absent for Hoechst as well, but not for EB. Thus, it can be stated that the chromophore of 7AAMD does not intercalate into the DNA duplex in a stacking manner between nucleotides, i.e., it is non-stacking binding. Anyway, the 7AAMD fluorescence arises mostly from hairpin-like sites and loops. It should be noted here that the efficiency of the energy migration between nucleotides in DNA and HP1 (in aqueous solution) is about zero [11, 12, 19, 23]. The absence of energy migration along a poly-nucleotide chain is very important for correct calculation of N and Qda .

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4 Estimation of Size of the 7AAMD/HP1 Complex The effective volume (V) and hydrodynamic length (diameter d) and the polarization and lifetime data of the 7AAMD/HP1 complex were determined using the Levshin– Perrin equation. Polarized emission of 7AAMD was measured with a SLM-4800 spectrofluorimeter (SLM Inc., United States). The fluorescence lifetime was determined with phase-modulation techniques (using SLM; the modulation frequency was 30 MHz). The lifetime experiments were carried out in mirror micro-cuvettes, which highly increase the fluorescence signal [19]. The effective volume of the 7AAMD/DNA complex was calculated according to the Levshin–Perrin relationship [19]: 1/P − 1/3 = (1/P0 − 1/3)(1+RTτ/ηV) where P is the measured polarization, P0 is the limit polarization = 0.5, R is the Rydberg’s constant, T is the absolute temperature in K, t is the lifetime, V is the effective volume of a particle, and h is the solvent viscosity (0.01 poise for aqueous solutions). The lifetime of 7AAMD in the complex with HP1 was 1.7 ns and the polarization was 0.39. These values coincide with our previous data obtained under similar experimental conditions [12, 16]. Thus, the V value of the 7AAMD/HP1 complex is ∼22800 Å3 . The d value of the complex is therefore ∼35 Å. In the case of free 7AAMD the V value is 4400 Å3 and d is 20.3 Å. The ratio of the volumes of the complex and single 7AAMD is 5.2. This value is smaller than the ratio between the molecular weights of the complex and free 7AAMD, which is 7.4. The observed discrepancy between the two mentioned volumes may be due to that 7AAMD is located in the cavity inside NR1 (not on the surface of HP1).

5 Prompt Binding of 7AAMD to HP1 Using stopped-flow fluorimetry, Chen et al [24]. observed two components in the 7AAMD fluorescence kinetics during its dissociation from short hairpin-like oligonucleotides in the presence of a detergent. Dissociation of 7AAMD from hairpin structures was considered to be a fast process, whereas dissociation from duplex DNA was assumed to be a slow process. The formation of the complex between HP1 and 7AAMD is a prompt process. It is much faster than the first component of 7AAMD binding to DNA. In the stoppedflow experiments, it was found that the kinetics of binding of 7AAMD to HP1 (Fig. 4) consist only of the fast single component. The characteristic time of the 7AAMD/HP1 complex formation is about 0.3 s (Table 3 ). No slower component was observed. The emission spectrum and lifetime of 7AAMD in the 7AAMD/HP1 complex were almost the same as those for 7AAMD


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Fig. 4 Stopped-flow kinetics of binding 7AAMD to HP1 (1) and substitution of 7AAMD in the 7AAMD/HP1 complex after addition of AMD (2). The concentrations were 7AAMD 2 μM, HP1 10 μM, and AMD 10 μM. The buffer was 10 mM Tris-HCl containing 0.5 mM EDTA (pH 7.6), at 26◦ C. Fast association–dissociation kinetics of binding of 7AAMD to HP1 was detected with stopped-flow fluorimeter “Applied Photophysics” (UK) with excitation at 550 nm and emission at >570 nm Table 3 Fast and slow components of 7AAMD or AMD binding to HP1 and λDNA, calculated from changes in the 7AAMD emission kinetics. The concentrations used: 7AAMD 1 μM, HP1 2.4 μM (i.e., 0.021 mg/ml), λDNA 0.014 mg/ml, and NaCl 50 mM. The buffer was 20 mM TrisHCl containing 1 mM EDTA (pH 7.5). Excitation and emission were at 570 and 610 nm. In the two upper stopped-flow experiments the concentrations were 7AAMD 2 μM, HP1 10 μM, and AMD 10 μM; excitation was at 550 nm, and emission was detected at >570 nm. The values of t1, t2, a1, and a2 were obtained by kinetic data fitting. Kinetics evaluation data were processed using Matlab 4.0 software (MathWorks, Inc.), with fitting by least-squares methods (Gauss–Newton approximation) Sample

t1 (s)

t2 (s)

A2/a1

7AAMD + HP1 7AAMD and HP1 + AMD 7AAMD + DNA 7AAMD and HP1 + DNA 7AAMD, NaCl, and HP1 + DNA

0.3 0.3 4.6 4.5 4.9

– – 40 101 147

– – 0.3 1.7 3.5

in pyridine [16]. This constitutes evidence that the binding of 7AAMD to the outer surface of HP1 is uncertain. The binding process seems to comprise rapid penetration of 7AAMD into a hydrophobic cavity inside HP1, but not the stacking intercalation of 7AAMD between the planes of nucleotides. The 7AAMD molecule might be located inside the HP1 cavity between adenine and thymine bases without stacking-like intercalation. To study the competition between 7AAMD and AMD for the binding site in HP1, excess AMD was added to the 7AAMD/HP1 complex. The fluorescence intensity decreased immediately after the addition (Fig. 5 , Table 1).

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Fig. 5 Emission spectra (at excitation wavelength 540 nm): 1 – the 7AAMD–HP1 complex, 2 – the 7AAMD–HP1 complex after addition of AMD (AMD concentration was tenfold higher than 7AAMD), 3 – 7AAMD. Cacodylate buffer, pH = 7. The concentration of 7AAMD is 1 μM

Consequently, 7AAMD was completely replaced in the complex by AMD. The replacement of 7AAMD by AMD is a very fast process. The characteristic time is about 0.3 s. The time of replacement of 7AAMD by AMD is the same as the characteristic time of the binding of 7AAMD to HP1. Thus, one can conclude that there is no energy barrier for substitution of 7AAMD by AMD in the complex with HP1 and vice versa. Similar (but not so pronounced) competition between 7AAMD and AMD was also observed in DNA.

6 Binding of 7AAMD to DNA It is well known that the binding kinetics of 7AAMD or AMD interaction with DNA are biphasic: fast (ms/s) and slow (minutes) processes take place [24–27]. Figure 6 shows the emission kinetics of 7AAMD binding to an excess of native λDNA. The experimental kinetics are described by at least two components: with characteristic times of 4.6 and 40 s (Table 3). The first component exhibits the higher amplitude (slow to fast component ratio a2:a1 is 0.3). Since λDNA is relatively short, it has almost no breaks. It can be suggested that the fast component corresponds

Fig. 6 The kinetics of 7AAMD binding to lambda DNA (2), its fitting (3), and 7AAMD without DNA (1). Excitation was at 570 nm, and emission was at 610 nm. The concentrations were 7AAMD 1 μM and λDNA 0.014 mg/ml. The buffer was 20 mM Tris-HCl (pH 7.6)


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to the binding of 7AAMD inside loops and single-strand areas, whereas the slow component corresponds to penetration of 7AAMD into the double helix. The duration of the first component in the case of λDNA was one order of magnitude longer than that for HP1. This reflects the fact that hairpin-like sites and loops in λDNA are compact rigid structures, whereas the HP1 hairpin is flexible and loosened. In the case of λDNA, the slow component corresponds to not only slow embedding of 7AAMD into a double helix, but also dilatory binding of 7AAMD with, due to thermal fluctuations, dynamically slow appearing hairpin-like sites and loops in the long DNA molecule. The rate of binding to DNA and the equilibrium fluorescence intensity of 7AAMD depend on the DNA concentration in the solution. Figure 7 shows that λDNA at concentrations of 5–10 μM of bases is unable to bind whole 1 μM 7AAMD. The 7AAMD emission achieves a plateau at a concentration of ∼22 μM of bases (i.e., 7 μg/ml; the molecular weight of λDNA is 32 × 106 ). At this concentration, λDNA binds 99% of 1 mM 7AAMD, i.e., the nucleotide/7AAMD ratio is ∼22. This means that native λDNA in aqueous solution (in buffers of low ionic strength, without salts) can form one hairpin-like site or loop every 37 Å of double helix. Fig. 7 Intensity of the 7AAMD emission depending on the λDNA concentration (mM nucleotides). Excitation was at 570 nm and emission at 610 nm. The concentration of 7AAMD was 1 mM. The buffer was 20 mM Tris-HCl containing 1 mM EDTA (pH 7.5), at 20◦ C

7 Distribution of 7AAMD from HP1 to DNA The phenoxazone chromophore of 7AAMD or AMD is located in DNA molecules in a hydrophobic environment [12, 16]. The fluorescence quantum yield of 7AAMD highly increases after 7AAMD binds to HP1, but not so highly when 7AAMD binds to DNA (Table 1). This means that photo-excited 7AAMD molecules are quenched by nucleotides in DNA stronger than in HP1. This fact allows observation of the redistribution of 7AAMD from HP1 to DNA. Indeed, a considerable decrease in the 7AAMD emission during incubation of the 7AAMD/HP1 complex with DNA was detected (Fig. 8).

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Fig. 8 Emission kinetics for redistribution of 7AAMD from HP1 to λDNA (with various λDNA concentrations). Concentrations of λDNA were 0.0 (1), 0.0035 mg/ml (2), 0.007 mg/ml (3), and 0.014 mg/ml (4). The concentration of 7AAMD was 1 μM, HP1 0.021 mg/ml. The buffer was 20 mM Tris-HCl (pH 7.6). Excitation was at 570 nm and emission at 610 nm. The curves for a one-exponential model (5) and a two-exponential model (6) are shown

Consequently, 7AAMD is transferred from HP1 to DNA after addition of DNA to the 7AAMD/HP1 complex. Rapid and slow phases in this process were observed. At high concentrations of λDNA, the redistribution kinetics were fitted by a twocomponent model with characteristic times of 4.5 and 101 s; the amplitude ratio a2/a1 was 1.7 (Table 3). The fluorescence changes caused by the slow component were larger. The first component has almost the same duration as in the case of direct binding of 7AAMD to DNA. The second component has a 2.5 times longer characteristic time compared to direct binding of 7AAMD to DNA. The process of redistribution was faster with increasing DNA concentration. The two detected components might correspond to the two different DNA sites, to which 7AAMD is transferred. The increase in ionic strength (50 mM NaCl) leads to a 1.8-fold decrease in the amplitude of the fast component, whereas the slow component does not change much. Ionic interactions control the rate of 7AAMD distribution from HP1 to DNA due to the well-known conformational transition of a loosened DNA in water without salts to a more rigid compact DNA at high ionic strength. HP1 acts as a protecting coat for 7AAMD or AMD. But this coat does not prevent the penetration of actinomycins into DNA. Perhaps hairpin oligonucleotides could be tested for medicines like molecular carriers of antibiotics.

8 Caffeine as a Potential Carrier of AMD to DNA Caffeine (1,3,7-trimethylxanthin, Fig. 9 ) clusters that spontaneously arise in its water solutions at mM concentrations [28] could be other carriers for amphiphilic heterocyclic antibiotics.


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Fig. 9 Caffeine (1,3,7-trimethylxanthin)

Caffeine and its clusters form stable complexes with many heteroaromatic molecules. At mM concentrations, AMD is strongly dimerized and also aggregated, which could reduce its ability to interact with caffeine clusters. Moreover, AMD causes heavy toxic consequences at millimolar concentrations, i.e., such concentrations are not physiologic. Unlike hairpin oligonucleotides that require a labor-consuming synthesis and could exert unknown effects on organism even in small quantities, caffeine is rather accessible, and its influence on organism has been well investigated. The safe single doze of caffeine can reach 100 mg. Caffeine possesses multiple effects at cellular level. It belongs to a class of soft psychomotor stimulators. Caffeine reduces carcinogenic action of ethidium bromide probably due to the formation of a strong complex between them. Caffeine should reduce the action of antitumor preparations by decreasing their effective concentration in blood. However, in many cases, the ability to reach the molecular targets in cancer cells rather than the concentration of medicinal substance in blood as such is important. Caffeine can easily penetrate into cells, whereas AMD cannot [29]. Action of actinomycins together with caffeine on tumor cells is much stronger [29]. Since the detection of smaller (μ molar) concentrations of AMD by spectrophotometry is impossible, the experiments were carried out with 7AAMD. One can presume that 7AAMD can form with caffeine the same complexes as AMD. Despite the presence in 7AAMD a polar basic amino group, its physical and chemical properties are very close to those of AMD. For example, 10 μM AMD immediately (for 0.3 s) displaces 1 μM 7AAMD from oligonucleotide hairpin. Note that 7AAMD itself is not used as an antibiotic only because of its high price.

9 Sorption of 7AAMD on Caffeine Clusters The fluorescence of dissolved 7AAMD is partially quenched by water. An addition of caffeine clusters to 7AAMD increases intensity of its fluorescence, which proves the transition of the antibiotic from a polar water phase into the less polar, on the surface of caffeine clusters. The constant of 7AAMD binding with the clusters reaches 106 M−1 (obtained by concentration titration). A substantial long-wave shift of excitation spectrum

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Fig. 10 Excitation spectrum of 2 μM 7AAMD (1) in a water solution, (2) in 8 mM caffeine solution, and (3) after addition DNA to the first or the second sample. Emission at 700 nm. Excitation slits of 3 nm, emission slits of 10 nm

is observed at the binding (Fig. 10), which proves the redistribution of electronic density in the antibiotic molecule as a result of its interaction with caffeine cluster. Interestingly, however, no shift of the emission spectrum occurs (Fig. 11), which unequivocally confirms the surface binding of the antibiotic. The phenoxazone chromophore of 7AAMD that strongly interacts with the surface of caffeine cluster contacts water. At photo-excitation, the dipole moment of 7AAMD sharply grows, and the relaxation warming up of the microenvironment also occurs due to a difference between the energies of the absorbed and emitted quanta. As a result, 7AAMD is desorbed from the cluster surface into the water phase, where it emits a quantum of light. Fig. 11 Emission spectrum of 2 μM 7AAMD (1) in buffer, (2) in 8 mM caffeine solution, and (3) after addition DNA to the first or the second sample. Excitation at 530 nm. Excitation slits of 10 nm, emission slits of 3 nm

These considerations are confirmed by that, upon the excitation at 560 nm, the degree of fluorescence polarization of 1 μM 7AAMD in the presence of 8 mM caffeine was the same as for free 7AAMD in water (0.28 and 0.29, respectively). The same conclusions may be inferred from the lifetime of the excited state of 7AAMD (1 ns in both cases). The emitting quantum yield is φ = τ /τ a (where τ is the


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measured lifetime of 7AAMD, and τ a its natural lifetime in absence of deactivation). This means that an increase in the antibiotic emission upon the binding with the clusters is caused by the reduction of its natural lifetime. Although the value of τ for 7AAMD is rather small, but, nevertheless, for 1 ns, the excited molecule could find time to collide with the quenchers added from outside, e.g., with dinitrophenol. The fluorescence of 7AAMD in the presence of caffeine clusters was quenched by dinitrophenol (taken in mM concentrations) at smaller degree than in their absence (Fig. 12). Fig. 12 Quenching (I0 /I) with dinitrophenol of the fluorescence of 5 μM 7AAMD (1) in the presence of 8 mM caffeine and (2) without it. λex 540, λem 660 nm. I0 and I, initial light intensity and that in the presence of quencher

The dependence of the intensity or the quantum yield of 7AAMD emission on dinitrophenol concentration can be described by the Stern–Volmer phenomenological equation: φ/φq = 1+kq Cq where φ and φ q are quantum yields of emission in the absence and in the presence of quencher, Cq is the quencher concentration, and kq is the quenching constant. The constant of 7AAMD fluorescence quenching with dinitrophenol in the presence of caffeine clusters was markedly lower than without them: Kq were on the average ∼85 and ∼280 M−1 , respectively. Therefore, the binding with caffeine leads to a steric reduction of the antibiotic availability for quencher. The dinitrophenol molecule transferred from water phase to the cluster surface deactivates by its hydroxy group and two nitro groups the excited 7AAMD molecule at the collision with it on the cluster surface. The mechanism of fluorescence quenching with nitro compounds is considered to consist in the charge transfer. The resonance energy transfer is here impossible: the emission spectrum of 7AAMD is not overlapped in any way with the light absorption band of dinitrophenol. Large dinitrophenol molecule is hardly capable of deep penetration into the densely packed caffeine cluster. If the 7AAMD phenoxazone chromophore were inside the cluster, dinitrophenol could not quench the fluorescence at all.

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10 Redistribution of 7AAMD from Caffeine Clusters to DNA 7AAMD binding with DNA results in approximately twofold growth in its lifetime and a shift of its emission spectrum to short-wavelength area. The complex formation of 7AAMD with DNA leads to a frequentative increase in the observed quantum yield (excitation at 570 nm and detection at 630 nm) since the phenoxazone chromophore of the antibiotic in a hydrophobic environment of DNA nucleotides is subjected to significantly smaller quenching than in water phase. In this case, the excitation spectrum of 7AAMD undergoes a strong long-wave shift (up to 570 nm), while the emission spectrum undergoes a shortwave shift (up to 630 nm). This is the reason why the addition of DNA solution aliquots to a solution of 7AAMD with caffeine clusters is accompanied by an additional long-wave shift of excitation spectrum (Fig. 10), the fluorescence intensity increases, and the emission spectrum is shifted to short-wave area (Fig. 11). These changes reflect the redistribution of antibiotic from a surface of caffeine clusters into the hydrophobic areas inside DNA. The final intensity of 7AAMD fluorescence in the presence of 200 μM DNA nucleotides in the buffer without caffeine is only 20% higher than in the presence of caffeine clusters (Fig. 13). Fig. 13 Dependence of fluorescence intensity of 1 μM 7AAMD on the concentration of added DNA (after recalculation on nucleotides) in Tris buffer in the (1) absence and (2) in the presence of 7 mM caffeine (λex 560, λem 630 nm)

This fact indicates that caffeine interferes with the formation of complexes of the antibiotic with DNA a little and can potentially serve as an effective carrier of an antibiotic to DNA. And the redistribution of the antibiotic to DNA in the course of dissociation of 7AAMD–caffeine complexes is caused by both the greater stability of the 7AAMD–DNA complexes and the presence in DNA of much more number of binding sites. It is possible to presume that caffeine can serve as carrier (“supplier”) to DNA of not only 7AAMD, but also AMD, since the physicochemical properties of 7AAMD and AMD are very close. The obtained data can be valid not only for model systems (solutions in water) but also for cellular systems. One of the ways of increasing the stability of the complex (i.e., for more successful passage through cell membranes) is the formation of such caffeine structures in which the antibiotic is hidden inside.


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11 Conclusion Complexes of actinomycin D (AMD) and 7-amino-actinomycin D (7AAMD) with model hairpin oligonucleotide HP1 and various types of DNA in aqueous solutions were investigated by steady-state, polarized, time-resolved, and stopped-flow fluorimetry. Prompt non-stacking binding of the actinomycins inside HP1 was observed. No energy transfer from nucleotides to 7AAMD in the complex was detected, most likely because of the absence of stacking intercalation. The data confirm the nonstacking binding of 7AAMD or AMD to HP1 and DNA in aqueous solutions. Strong competition between AMD and 7AAMD for a cavity in HP1 was observed. The decrease in the 7AAMD emission after addition of DNA to the 7AAMD/HP1 complex indicates that actinomycins can be redistributed from HP1 to DNA. The biological significance of this finding is that hairpin oligonucleotides can serve as molecular carriers, transferring actinomycins to DNA. Addition of 7AAMD to the caffeine clusters leads to its sorption on the cluster surface. Photo-excitation of 7AAMD leads to its desorption from the surface into the aqueous phase and emission of a quantum. Fluorescence of 7AAMD in the presence of caffeine clusters is quenched by dinitrophenol more weakly than without clusters due to decreased steric availability of the antibiotic to the quencher. Addition of 7AAMD–caffeine complexes to DNA leads to a long-wavelength shift in the excitation spectrum and an increase in the fluorescence intensity along with a shift of the fluorescence spectrum to the short-wavelength area. This fact reflects redistribution of the antibiotic from the caffeine surface to the hydrophobic areas inside DNA. Acknowledgments N.V. is grateful to Dr. R. Wadkins (USA), Dr. A. Kovalev (Germany), Dr. I. Savintsev and Ph.D. student M. Bitekhtina (Russia) for kind assistance and to CRDF and NATO for financial support.

References 1. T. Kaji, H. Takamatsu, H. Noguchi, H. Tahara, Y. Adachi, H. Kajiya, K. Kawakami, S. Nomoto, and E. Satou, Yolk sac tumour of the ovary in a conjoined twin. Eur. J. Pediatr. Surg. 7, 311–312 (1999). 2. K. Shen, J. Lang, and H. Huang, Treatment of childhood genital malignancies, Zhonghua Fu Chan Ke Za Zhi 36, 360–363 (2001). 3. N. Ducrey, M. Nenadov-Beck, and B. Spahn, Update of orbital rhabdomyosarcoma therapy in children, J. Fr. Ophtalmol. 25, 298–302 (2002). 4. T. Sokolov, A. Stoianova, I. Mumdjiev, and A. Mihova, Treatment of Ewing’s sarcoma with 2 different protocols, Ann. Med. Interne 152, 497–501 (2001). 5. V. V. Frolkis, Aging and Increase of Life Expectancy, Nauka (Leningrad, USSR, 1988). 6. W. Muller and D. M. Crothers, Interactions of heteroaromatic compounds with nucleic acids: the influence of heteroatoms and polarizability on the base specificity of intercalating ligands, Eur. J. Biochem. 54, 267–277 (1975). 7. V. Gurski, Structure of DNA-actinomycin complexes, Mol. Biol. (Moscow) 3, 749–756 (1969). 8. M. A. Krivtsova, E. B. Moroshkina, H. Hamman, E. N. Glibin, and E. V. Frisman, Study of interaction of DNA with small-molecular ligands of various structure, Mol. Biol. (Moscow) 15, 613–621 (1981).

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9. H. Robinson, Y. -G. Gao, X. -L. Yang, R. Sanishvili, A. Joachimiak, and A. H. -J. Wang, Crystallographic analysis of a novel complex of actinomycin D bound to the DNA decamer CGATCGATCG, Biochemistry 40, 5587–5592 (2001). 10. J. G. Snyder, Binding of actinomycin D to DNA: evidence for a nonclassical high-affinity binding mode that does not require GpC sites, Proc. Natl. Acad. Sci. U S A 86, 3968–3972 (1989). 11. I. V. Savintsev and N. L. Vekshin, Formation of complexes of actinomycins with DNA in solutions and films, Appl. Biochem. Microbiol. (Moscow) 41, 1–8 (2004). 12. I. V. Savintsev and N. L. Vekshin, Binding sites for 7-amino-actinomycin D and actinomycin D in DNA and model nucleotide systems, Mol. Biol. (Moscow) 36, 725–730 (2002). 13. R. M. Wadkins and T. M. Jovin, Actinomycin D and 7- aminoactinomycin D binding to singlestranded DNA, Biochemistry 30, 9469–9478 (1991). 14. R. M. Wadkins, D. Vladu, and C. Tunng, Actinomycin D binds to metastable hairpins in single-stranded DNA, Biochemistry 37, 11915–11923 (1998). 15. R. M. Wadkins, C. Tunng, P. M. Vallone, and A. S. Benight, The role of the loop in binding of an actinomycin D analog to hairpins formed by single-stranded DNA, Arch. Biochem. Biophys. 384, 199–203 (2000). 16. N. Vekshin, I. Savintsev, A. Kovalev, R. Yelemessov, and R. Wadkins, Solvatochromism of the excitation and emission spectra of 7-aminoactinomycin D: implications for drug recognition of DNA secondary structures, J. Phys. Chem. B 105, 8461–8467 (2001). 17. A. E. Kovalev, A. A. Yakovenko, and N. L. Vekshin, Investigation of 7-amino-actinomycin D and DNA by fluorescence correlated microscopy, Biophysics (Moscow) 49, 1030–1037 (2004). 18. F. Sha and F. M. Chen, Actinomycin D binds strongly to d(CGAGACG) and d(CGTCGTCG), Biophys. J. 79, 2095–2104 (2000). 19. N. L. Vekshin, Photonics of Biopolymers (Springer, Berlin, 2002). 20. N. L. Vekshin, Energy Transfer in Macromolecules (SPIE, Bellingham, 1997). 21. N. L. Vekshin, Migration of energy in DNA, estimated by fluorescence of intercalating dye, J. Appl. Spectrosc. (Minsk) 65, 794–798 (1998). 22. M. Saito, M. Kobayashi, S. -I. Iwabuchini, Y. Morita, Y. Takamura, and E. Tamiya, DNA condensation monitoring after interaction with Hoechst 33258 by atomic force microscopy and fluorescence spectroscopy, J. Biochem. 136, 813–823 (2004). 23. N. L. Vekshin, Screening hypochromism in molecular aggregates and biopolymers, J. Biol. Phys. 25, 339–354 (1999). 24. F. M. Chen, C. M. Jones, and Q. L. Johnson, Dissociation kinetics of actinomycin D from oligonucleotides with hairpin motifs, Biochemistry 32, 5554–5559 (1993). 25. R. Bittman and L. Blau, Stopped-flow kinetic studies of actinomycin binding to DNAs, Biochemistry 14, 2138–2145 (1975). 26. S. C. Brown and R. H. Shafer, Kinetic studies of actinomycin D binding to mono-, oligo-, and polynucleotides, Biochemistry 26, 277–282 (1977). 27. K. R. Fox and M. J. Waring, Kinetic evidence for redistribution of actinomycin molecules between potential DNA-binding sites, Eur. J. Biochem. 145, 579–586 (1984). 28. M. A. Bitekhtina and N. L. Vekshin, Caffeine clusters as transmitters of actinomycin antibiotics to DNA in solution, Russ. J. Bioorganic Chem. 34 (2), 234–238 (2008). 29. N. L. Vekshin, Biophysics of DNA-actinomycin nano-complexes (Pushchino: Photon-vek, 2009).

Fluorescence Spectroscopy in Optoelectronics, Photomedicine, and Investigation of Biomolecular Systems Danuta Wróbel, Alina Dudkowiak, and Jacek Goc

1 Introduction Fluorescence spectroscopy represents an universal method and very useful tool for the examination of excited states of molecules and deactivations of their energy excess by emitting photons. In this contribution we describe some examples and show how fluorescence spectroscopy can give insights into the interactions of light with organic matter to follow processes important in several fields. One of them is investigation of light energy conversion to electric energy in organic dye-sensitized photovoltaic devices and optoelectronics. In medicine, fluorescence spectroscopy can be very useful in diagnosis of cancerous cells. Also dynamic and light-induced inter- and intramolecular deactivation processes in molecular systems can be followed by this technique. This chapter shows that fluorescence spectroscopy is suitable in many new branches where not only radiative processes but also their dynamics can be followed. Also important is the search for mechanisms responsible for photophysical interactions in the systems. The systems under investigation can find future practical application in photoelectronics, photovoltaics, photomedicine, and biosystems with the use of organic materials. In particular in this chapter we present some experiments illustrating the possibilities of steady-state fluorescence spectroscopy in the investigation of photocurrent generation in an organic dye-sensitized cell, intermolecular electron transfer (ET) in a mixture composed of porphyrinic dyes and a chinone acceptor, intramolecular ET in donor–acceptor supermolecular covalent porphyrin–fullerene hybrids, and creation of an excimer. Also studies of effectiveness of dye incorporation into biological cells, properties, interaction, and location of dyes in cell structures, and the dye’s ability to trigger photochemical reactions and emit fluorescence in biological systems are presented. We are also going to review the advanced techniques D. Wróbel (B) Institute of Physics, Faculty of Technical Physics, Poznan University of Technology, Nieszawska 13A, 60-965, Poznan, Poland e-mail: [email protected] C.D. Geddes (ed.), Reviews in Fluorescence 2008, Reviews in Fluorescence 2008, DOI 10.1007/978-1-4419-1260-2_10, C Springer Science+Business Media, LLC 2010

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of time-resolved fluorescence quenching and application of ultrafast fluorescence spectroscopy in the study of dynamic light-induced inter- and intramolecular deactivation processes in molecular systems. Application of the time-correlated single photon counting technique for multi-exponential fluorescence decay in model systems of photosynthetic pigments is also presented.

2 Fluorescence Spectroscopy as a Tool in Organic Photovoltaics The molecular structures of selected organic materials discussed in this chapter are presented in Fig. 1.

2.1 Fluorescence and Photovoltaics In this section we focus our attention on organic dye fluorescence and its capabilities in a study of solar energy to electric energy conversion. The year 1991 in which Brian O’Regan and Michael Grätzel demonstrated photovoltaic devices for electricity generation based on a photoactive organic dye which sensitized a semiconductor, began a new era in solar photovoltaics based on organic materials [1]. The idea of O’Regan and Grätzel initiated an avalanche of work on organic materials that could be new generation materials for photovoltaic systems. It is very well known that fluorescence behavior of dyes is strongly connected with their molecular structure [2]. For about 20 years numerous papers have provided evidence of a close correlation between the dye molecular structure and its photoactivity in photocurrent generation when embedded in a photodevice. In photovoltaics, the singlet excited states of dyes or other photoactive molecular materials (e.g. organic macrocyclic dyes, supra- and supermolecular systems based on fullerenes or polymers with dyes) play an essential role in initiation of charge transfer from a dye to sensitize a semiconductor layer [3]. The simplest dyesensitized photodevice (DSPV), which can serve as a light-to-electric energy cell for photoconversion, is a sandwich-like photoelectrochemical cell (PEC – Fig. 2) [4]. The dye system (dye, dye–fullerene, dye–polymer, or others) after dissolution in a proper medium (organic solvent, nematic liquid crystal, polymer, etc) is introduced between two electrodes (metal and semiconductor) distanced with a 20– 60 μm spacer. After illumination with white or colored light, a photoresponse can be observed in close or short circuit modes. The mechanism, which is responsible for photosignal generation, involves electron injection from the excited molecular system (primary electron donor) to the semiconductor electrode (electron acceptor). A metallic electrode (e.g. Au, Ag) serves as a secondary donor and provides an electron in order to make a dye molecule return to its ground state [4–6]. Very good coincidence of a photovoltaic spectrum (dependence of photovoltaic signal in PEC versus excitation wavelength) and dye absorption spectrum is

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A

Me:H2, Mg, Zn, Pb

Me: Mg, Zn, R1-16: none or F (or Cl) R2,3,6,7,10,11,14,15: O(CH2)3CH3

B

C

Ar-3,5 – di(tert-butylo)phenyl

Fig. 1 Molecular structures of selected organic materials used and discussed in Sect. 2; (A) porphyrins (left) and phthalocyanines (right), (B) fluorescein (left) and phenolphthalein (right), and anthraquinone (bottom), (C) zinc porphyrin (left) and covalent porphyrin–fullerene (right), adapted from [10, 11, 16, 33, 42]

observed, and it evidently confirms responsibility of the dye for photovoltage generation [7, 8]. To study a correlation between the dye molecular structure and its photoactivity in photocurrent generation in PEC, the following families of porphyrins and phthalocyanines have been selected: metal-free and metal-complexed dyes, dyes substituted with aromatic groups or long alkyl/alkyloxy chains as well as fluorinated dyes. Porphyrins are characterized by strong absorption in the range of about 420–450 nm (Soret band) and much less intense Q bands in the long-wavelength region. Phthalocyanines have an intense “red” absorption band and

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Fig. 2 Scheme of a photoelectrochemical cell (PEC) and of the mechanism of photocurrent generation, adapted from [7]

e– S1 hν Fermi level

e–

S0 DYE IN SOLUTION

SEMICONDUCTOR

METAL

a much weaker Soret band [9]. Firstly, in this part of the paper we briefly describe the photocurrent dependence on the dye molecular structure. Attachment of substituents to the main molecular core in porphyrins and phthalocyanine leads to changes in their spectroscopic properties and in consequence, their photoelectric features. In the experiments tetraphenyl porphyrins (P’s) and phthalocyanines (Pc’s) complexed with the following ions: magnesium (Mg), zinc (Zn), and lead (Pb) were used [4]. The photoelectric examinations have shown a strong influence of the type of metal ion in the molecules on the values of the photocurrent generated in PEC. The Zn and Mg dyes give the highest photocurrents [4, 7]. Their counterparts, the dye complexed with the Pb ion, show very weak or no photoresponse [7]. Our electron spin resonance (ESR) investigation [10] has shown that a π -electron in a conjugated macroring porphyrin system is responsible for photoelectroactivity of the dyes and the metal ion markedly alters the distribution of the electron cloud in P and Pc dyes. Since the metal ion is able to change electron distribution thus it can influence the photoeffects; the higher the reactivity of the metal the stronger the π -electron delocalization. The presence of extra aromatic rings attached to the porphyrin’s ring or to the indole unit in phthalocyanines can lead to changes in the photoresponse due to a higher number and delocalization of π-electrons. However, when long alkyl or alkyloxy chains are bonded to the main molecular core, the effectiveness of the dyes in photoresponse is markedly diminished since most of the energy absorbed is lost in nonradiative transitions due to the flexibility of long chains and thus the energy is lost in thermal processes instead of in charge-separation processes [11]. A drastic rise in the photocurrent is observed in phthalocyanine when it is substituted with fluorine or chlorine atoms (FZnPc, ClZnPc) [12]. Fluorine and chlorine atoms attached externally to the main molecular macrorings strongly perturb the electron density distribution. According to the literature, fluorine has unique photophysical properties owing to its high electronegativity and therefore, it is expected to have electron withdrawing character but when it is linked to macrocycling systems it is able to convert into electro-donating species by the mesomeric resonance coupling. Conjugation of the fluorine (or chlorine) with the π -electron system in aromatic dyes can lead to increased photocurrent (Fig. 3) [12].

Fluorescence Spectroscopy in Optoelectronics 0 3 Photocurrent [nA/cm2]

Fig. 3 Photocurrent signals for selected dyes; ZnPc (1), FZnPc (2), ZnPcAlkyl (3). Arrows indicate “light on,” adapted from [11, 42]

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The charge separation process which directs the light-response of systems strongly competes with fluorescence and other processes, for example nonradiative thermal relaxation or charge recombination. Thus, to gain deep insight into the problem of performance of the photovoltaics based on organic materials, fluorescence measurements are extremely useful. In particular, not only the spectral fluorescence range but also fluorescence quantum yields and lifetime values are important. We focus our attention on selected dyes characterized by large photoresponses: magnesium and zinc porphyrins and phthalocyanines (good photoconverters), and those giving weak or no photosignals in PEC: lead dye (poor photoconverters). The lifetimes of porphyrins and phthalocyanines are of an order of nanoseconds. This means that ET should take place before a dye molecule decays spontaneously from the first excited state to the ground state. On the basis of fluorescence and absorption data the fluorescence natural lifetimes (τ N ) and fluorescence quantum yields (F ) can be evaluated [13, 14]. The data are collected in Table 1. Among the dyes investigated, fluorine zinc phthalocyanine (FZnPc) (Aldrich) shows unique properties: Stock’s absorption-fluorescence shift is the largest among the dyes under study; this dye is characterized by a long natural lifetime (τ N ) and high fluorescence quantum yield (F ). Also magnesium and zinc P’s and Pc’s exhibit long τ N but much lower F in comparison to FZnPc [11]. In contrast to zinc and magnesium dyes, their counterpart PbP shows rather weak fluorescence [10]. Moreover, thermal deactivation parameters in alkyl or alkyloxy porphyrins and phthalocyanines (evaluated on the basis of photoacoustic spectroscopy (PAS) [15]) are much higher with regard to those in other dyes, due to thermal deactivation processes. The fluorescence quantum yields of magnesium and zinc dyes are of

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Table 1 Spectral parameters of selected organic materials, based on [7, 10, 11, 33, 41, 43]

Sample

Fluorescence quantum yield, F

Natural lifetime, τ N (ns)

Average lifetime, τ 0 (ns)

Average lifetime in the presence of AQ, τ F (ns)

Thermal deactivation parameter, (a.u.)

H2 P MgP ZnP PbP MgPc ZnPc FZnPc ZnPcAlkyl PF

0.02 0.08 0.07 0.02 0.21 0.11 – 0.10 >0.01

16.3 31.8 25.1 6.2 11.2 7.7 16.5 14.0 –

0.33 2.54 1.76 0.12 – – – – –

0.30 1.11 1.07 0.08 – – – – –

– 1.00 1.20 – 1.23 3.18 3.52 7.12 –

an order of 0.1–0.2 and their thermal deactivation is low. Thus, it can be supposed that most of the energy absorbed could be efficiently utilized in a charge separation process. Thus, even fluorescence competes with the charge separation process as a sufficiently long fluorescence lifetime of the first excited singlet state is required for photocurrent generation since fluorescence decay has to be long enough for charge separation to occur when the first excited state is involved in this process. This supposition is supported by the results for fluorinated zinc phthalocyanine (FZnPc), which is characterized by a long lifetime and the highest fluorescence quantum yield among the dyes under study. A very nice example which confirms how photoelectric properties could be correlated with fluorescence behavior and with dye molecular structures is fluorescein and phenolphthalein (Sigma) [16]. Fluorescein and phenolphthalein differ in molecular structure; there is an O-bridge linked to two benzene rings in fluorescein [2]. This chemical modification has a great influence on the fluorescence features of the dyes: fluorescein and phenolphthalein in aqueous polyvinyl alcohol (PVA) (pH 8.5) absorb in the wide range of 430–530 nm and 430–630 nm, respectively (with a maximum at 496 and 558 nm, respectively). The molar absorption coefficient is twice as high for fluorescein ( = 0.46 × 104 M−1 cm−1 ) when one compares with that for phenolphthalein ( = 0.21 × 104 M−1 cm−1 ). Drastic changes are also observed in fluorescence; fluorescein is characterized by very intensive emission with a maximum at 520 nm and a very high fluorescence quantum yield of about 92% [2], whereas no fluorescence signal is provided from phenolphthalein [16]. The reason for this difference is the presence of the O-bridge, which makes the fluorescein molecule rigid, which leads to high fluorescence. Not only radiative fluorescence competes with the charge separation process. The fast thermal relaxation is known to be essential for photocurrent generation in PEC. PAS [15] allows determination of how much of the absorbed excitation energy is thermally deactivated in nonradiative processes. Thus, we also monitored phenolphthalein and fluorescein with PAS [16] however, the kinetics of thermal


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relaxation in phenolphthalein differs from that in fluorescein. Only for fluorescein, the decrease in the PAS signal in light modulation frequency is in accordance with the Rosencwaig theory [15]. In phenolphthalein a contribution of a triplet state to the thermal relaxation cannot be excluded. Since the fluorescence in fluorescein differs markedly from that in phenolphthalein, one can expect phenolphthalein dye to effectively convert light energy to electric energy. Taking into regard the relation between the absorbance (the molar absorption coefficient of fluorescein is twice as high as that of phenolphthalein) and the photovoltaic response observed in PEC with fluorescein or phenolphthalein (much higher photosignal in phenolphthalein than that in fluorescein) and the fluorescence results (quenched fluorescence in phenolphthalein when compared to high emission in fluorescein), it is evident that the ability of phenolphthalein for charge separation dominates over that in fluorescein. The photoresponses in PEC filled with the dyes are confirmed by the values of the photocurrent; phenolphthalein is characterized by a twice higher photocurrent value than fluorescein [16]. Taking into account the classical results of Birks [2] as well as those of Bowers and Porty [17] concerning the fluorescence and triplet quantum yields of fluorescein in aqueous solvent (0.97) we can suppose that the thermal deactivation and charge separation can be estimated to at least 0.03 in fluorescein and the photovoltaics in this dye have no chance of being observed. Moreover, taking into account the energy conservation law, fluorescence and photoacoustic evidences as well as the photoelectric behavior of phenolphthalein with regard to fluorescein, the capability of phenolphthalein for charge separation upon illumination cannot be excluded.

2.2 Fluorescence Quenching – Donor–Acceptor Pair in Mixed Bimolecular Systems Steady-state fluorescence spectroscopy can also be used as a perfect tool in the investigation of intermolecular ET. This section is devoted to the investigation of photoinduced electron transport in a mixture composed of porphyrin and anthrachinone (AQ) in dimethylsulfoxide (DMSO) by quenching fluorescence and steady-state ESR. As shown in [10] this molecular system could be a simple model of ET from an excited dye to a semiconductor electrode as an acceptor. In this experiment metal-free or metallic tetraphenylporphyrin dyes: H2 P, MgP, ZnP, and PbP (Porphyrin System) and AQ (Aldrich) were used. The porphyrin dyes are characterized by a strong Soret band in the range 440–470 nm and by much less intensive Q bands in the range 500–800 nm [9]. MgP and ZnP have large molar absorption coefficients and good effectiveness in light energy to electric energy conversion [4]. In contrast to the above-mentioned dyes, PbP is not as efficient in photoconversion. Fluorescence measurements supported by the absorption study allowed evaluation of the natural lifetimes (6.2–31.8 ns depending on the dye), fluorescence quantum yields (0.2–0.8), and average lifetimes in the absence (0.12–2.54 ns) and the presence (0.08–1.11 ns) of AQ. The results showed a decline of the investigated

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porphyrin dye’s emission upon successive addition of AQ as the external quencher. The fluorescence quenching data and analysis of the Stern–Volmer equation [10, 18, 19]: F0 /F = 1 + KQ [Q] (where F0 and F are the dye fluorescence intensities in the absence and presence of the quencher Q, and KQ is the Stern–Volmer quenching constant which can be expressed as KQ = τ 0 kq [20]) together with fluorescence lifetime, τ 0 of the dye in the absence of the quencher and biomolecular quenching rate, kq proved the linear character of the Stern–Volmer plots confirming homogeneous dynamic quenching of porphyrin dyes by AQ and permitted evaluation of the Stern– Volmer quenching constants (1.5 × 104 M−1 to 1.7 × 104 M−1 depending on the dyes) [10, 18]. The bimolecular quenching rate kq for metallic-free and metal porphyrins (MgP, ZnP, PbP) ranged from 0.7 × 1013 M−1 s−1 to 1.3 × 1014 M−1 s−1 . The lowest kq values were found for MgTPP and ZnTPP in contrast to PbTPP whose quenching was very fast in the presence of AQ. The details of the fluorescence and ESR experiments can be found in [10]. The fluorescence quenching in the presence of AQ was supposed to be induced by ET from porphyrin moieties to AQ. On the basis of the fluorescence lifetimes and the equation kET = 1/τ F −1/τ 0 [21] (where τ F and τ 0 are the fluorescence lifetimes of the porphyrin dyes in the presence and absence of AQ, respectively) the rate constant of ET can be evaluated as 0.50 × 109 s−1 , 0.36 × 109 s−1 and 4.17 × 109 s−1 for MgPc, ZnPc, and PbPc, respectively. The light-induced ESR signals confirmed the thesis as to the origin of the unpaired electrons [10]. No signal was observed for AQ and the porphyrins in the absence of the quencher either in the dark or in the light. MgP and ZnP in the light were most sensitive to the presence of AQ from among the dyes investigated, showing a significant increase in the number of unpaired electrons in addition to being very good electron donors. In contrast, a modest change in the ESR signal was observed for PbP, since its unpaired electron concentration was the lowest. The evaluated g-values (2.0043 and 2.0045 for MgP and ZnP, respectively) and the ESR linewidth (B = 0.35 mT) provided evidence allowing identification of the sources of the electrons [10, 21, 22]. We showed that the fluorescence quenching spectroscopy supported by the ESR examinations permits identification of the origin of the unpaired electron in the porphyrin–chinone systems. The ESR parameters of the dyes are collected in Table 2.

Table 2 Fluorescence and ESR parameters for porphyrin–anthracene mixture and covalent porphyrin–fullerene system, based on [10, 36] Stern–Volmer quenching constant, Sample KQ ∗ 104 (M−1 )

Bimolecular quenching rate, kq ∗ 1013 (M−1 s−1 )

Electron transport rate, kET ∗ 109 (s−1 )

g-factor

B (mT)

H2 P MgP ZnP PbP PF

7.0 0.7 0.9 13.0 –

0.30 0.30 0.30 0.30 –

2.0046 2.0043 2.0045 2.0044 2.0027

0.28 0.35 0.35 0.32 0.55

2.3 1.7 1.5 1.5 –


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At this point it is worth mentioning again that MgP and ZnP are the best photoconverters of light energy to electric energy from among the dyes investigated, relative to PbP showing very poor photoresponse. The experimental fluorescence and ESR results are shown in Fig. 4. The kq result for PbP which shows faster fluorescence quenching (kq = 1.3 × 1014 M−1 s−1 ) when compared to that for MgP (kq = 0.7 × 1013 M−1 s−1 ) or ZnP (kq = 0.9 × 1013 M−1 s−1 ) and the largest changes in the ESR signals observed for Mg and Zn dyes in the presence of the quencher upon illumination seem to be in contradiction. However, analysis of the fluorescence results in combination with the ESR results, permits identification of the mechanisms responsible for the effects observed. The reason for the incoherence between the fluorescence and ESR observations could be the differences in the order of the rate values in the time range of fluorescence (ns) and ESR (μs). It is well known that not only ET but also thermal relaxations strongly decrease lifetimes [2, 10, 23] when spin-orbit processes are involved. The second very important reason could be the charge recombination process, which always accompanies and competes with electron separation and can proceed at different rates in PbP and MgP or ZnP. The low kq for MgP and ZnP and their highest ESR signals when compared to those for PbP, strongly suggest a slower charge recombination process occurring in the electron donor–acceptor chain in MgP and ZnP when compared to PbTPP. The values of the lifetimes, fluorescence quantum yields, and ET rate constants could support differences in charge recombination for MgP, ZnP, and PbP. The values of kq and kET of MgP and ZnP are as follows: kq = (0.7–0.9) × 1013 M−1 s−1 and kET = (0.36–0.50) × 109 s−1 , when compared to the values for PbP; kq = 1.3 × 1014 M−1 s−1 and kET = 4.1 × 109 s−1 . In this section we have shown that the fluorescence quenching experiments accompanied by the ESR examinations can explain why some dyes could be perfect

4

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40 2 20 3

0 0

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100

0 –6.0

–5.5

–5.0

–4.5

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Log C (AQ)

Fig. 4 Fluorescence [Stern–Volmer curves (A)] and ESR data (B) for a porphyrin–anthraquinone mixture, adapted from [10]

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agents in light energy to electric energy conversion (MgP and ZnP), while others (PbP) are useless in organic photovoltaics.

2.3 Fluorescence in Supermolecular Porphyrin–Fullerene Systems Supermolecular systems built up of chromophores [24] covalently linked to fullerene [25] or polymers [26] are perfect electron donor–acceptor systems owing to the strongly conjugated π-systems. The supermolecular system can serve as a model of light energy conversion in photosynthetic reaction centers [27–29] or in organic photovoltaics [30–32]. P’s or Pc’s covalently bonded to fullerene are good examples of systems showing intramolecular interaction and ET upon photoexcitation. In this section we present the results of steady-state fluorescence spectroscopy when applied to porphyrin (P) and porphyrin covalently linked to fullerene C60 (PF) dissolved in DMSO or chloroform (obtained kindly from Prof. Diderich and Dr. Bonifazi – Zürich). In our spectroscopic examinations (absorption and steady-state fluorescence) monomeric porphyrin was used as a reference sample. The first stage of the study of the system was absorption measurement which showed the strong contribution of the ground state to the second excited state transition in the range of the Soret bands (400–450 nm) and much less intensive Q bands (500–600 nm) connected with transition from the singlet state to the first excited singlet state of the porphyrin moieties [33]. The bathochromic shift (of about 10 nm) in the absorption spectra of the supermolecular PF system with respect to that in the reference sample indicates very strong influence of the fullerene on the electron redistribution in porphyrin moieties. The strong interaction between porphyrin moieties and fullerene was also confirmed by our fluorescence examinations. The fluorescence spectra of the porphyrin studied revealed bands at about 590 and 640 nm in DMSO and 587 and 633 nm in chloroform, respectively, and are assigned to the emission of monomeric porphyrins (Fig. 5) [33, 34]. After porphyrin linkage to fullerene (PF) the porphyrin fluorescence shape depends very strongly on the excitation wavelength and the type of solvent or to be more exact on the solvent polarity (DMSO or chloroform). As shown in [33] porphyrin emission in PF is markedly quenched by fullerene. In PF porphyrin fluorescence intensity declines more than ten times in contrast to that of the porphyrin. This result indicates the occurrence of processes competitive to radiative emission. On the one hand, they could be nonradiative intersystem crossing transitions and internal conversion [33]. On the other hand, the strong fluorescence quenching can result in intramolecular ET from the porphyrin species to fullerene. In this reaction porphyrin serves as an electron donor and fullerene is an electron acceptor [33, 35]. To verify the thesis of intramolecular interaction leading to intramolecular ET from porphyrin to fullerene units, the ESR study was undertaken. The experiments were performed in the dark and in the light (white light of 40 mW cm−2 ). The experimental method is described in detail in [36]. The ESR parameters are collected in

Fluorescence Spectroscopy in Optoelectronics A

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Fig. 5 Absorption (A, B) and fluorescence spectra (C, D) of porphyrin (P; left) and a porphyrin– fullerene system (PF; right) in DMSO and chloroform, adapted from [33]

Table 2. In the dark either no or very tiny signals of the porphyrin were observed. This result evidently shows that the dye molecules in their ground state do not generate unpaired electrons. Moreover, about 20-times higher ESR signals in the porphyrin–fullerene sample were observed when compared to that in the porphyrin. The g-factor (2.0027) and linewidth B (approximately 0.55 mT) were evaluated. The ESR parameters correspond perfectly to those of porphyrins and phthalocyanines [22, 37, 38]. The appearance of the ESR photosignal confirmed light-induced generation of unpaired spin states can be explained by (1) low deactivation rate of the triplet states, (2) existence of unpaired electrons in broken π -bonds in the porphyrin macrorings, (3) creation of a polaron state of porphyrins owing to deactivation of porphyrin triplet states. Thus, the g-factor values of about 2.0027 can be attributed to the unpaired electrons in porphyrin triplet states or on porphyrin cation radicals [39]. The fullerene also exhibits weak fluorescence, which indicates a contribution of its excited state in radiative processes. The process of excitation energy transfer (EET) in the porphyrin–fullerene supermolecular system under study was observed besides ET between porphyrin and fullerene units. Such a supposition can be confirmed by the fact of the porphyrin moieties emission (observed as a sharp peak at 535 nm placed against the background of 550–750 nm of porphyrin fluorescence) when the system was excited in the absorption range of fullerene (λexc =

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250 nm). Confirmation of the occurrence of EET in PF was also found for the systems dissolved in DMSO and also in chloroform and excited at different excitation wavelengths. The fluorescence and emission excitation spectra provide information on an excimer formation in porphyrin–fullerene supermolecular systems. In the covalent porphyrin–fullerene, the fluorescence methods were also used for discussion of excimer creation thanks to influence of fullerene on chromophore properties. Two experimental approaches: fluorescence and emission excitation measurements were applied. The fluorescence measurements carried out for the porphyrin–fullerene samples revealed new bands (or humps) that appeared in the range of 530–595 nm as a result of excimer formation. The locations and intensities of the new bands were affected by both the type of samples and the solvents (maximum at 535 and 590 nm in chloroform and DMSO, respectively). Confirmation as to excimer creation can be discussed as follows: (1) these bands (or humps) were not seen in standard P fluorescence but were observed in PF fluorescence; they indicate the presence of the excimer in the supermolecular porphyrin–fullerene systems under study; (2) variation in excitation emission spectra shapes depends on observation regions (not shown); this indicates coexistence of various spectral porphyrin moieties; (3) in the observation range beyond 600 nm emission originates from porphyrin; in contrast, when the observations were performed in the range of 540–590 nm, an intense band at about 520–540 nm appeared and was assigned to the emission of the excimer. The presence of the extra bands in the energy range below the energy of the porphyrin second excited state and a strong influence of the solvent polarity (highly polar DMSO and chloroform of low polarity) suggest the formation of the excimer with the second excited singlet state as a result of strong interaction between the ground and excited states of the system [33]. In this section we presented some evidence which supports the successful use of fluorescence spectroscopy in following strong interactions in supermolecular systems like porphyrin–fullerene dyads. Strong interaction can lead to charge redistribution and also to creation of the excimer. Such systems are able to form a charge-separated state, P•+ −F•− [30] and the excimer [40], which could be a precursor of such a state.

2.4 Enhanced Fluorescence – Dyes in Colloids of Metallic Particles Fluorescence spectroscopy has become one of the important tools for assays in biochemical and biomedical research, as well as in the biological sciences. For the last decade much attention has been paid to a very promising metal-enhanced fluorescence (MEF) method. This effect is related to the resonance interaction between the light-excited pigments and the surface plasmons, activated in certain conditions in the metallic surface. The specific interaction of colloidal metals and the dyes resulting in their unique spectral, electronic, and chemical properties is determined by


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the small size and high surface-volume ratio of nanoscopic systems. Moreover, the small metallic, silver and gold particles shine in bright colors in reflected light. Surface Plasmon Resonance (SPR) is the effect of energy resonance arising from the collective oscillation of migrated free electrons on metal subwavelength size particle surfaces. Plasmon resonance takes place when the wavevector of the incident light is equal to the wavevector of the surface plasmons [41]. In the early 1970 s, Drexhage et al. found that the emissive lifetime of a fluorophore localized at a certain distance from the metal film depends on this very distance [42]. Since then, it has been found that fluorescence can be quenched when the fluorophore is in close proximity to a metal particle, but enhanced when the fluorophore is localized at a certain distance from it. Such a fluorescence enhancement by the metal particle is known as metal-enhanced fluorescence. The metal-enhanced effect is a result of surface plasmons induced by the incident light or by the excited fluorophores. The MEF fluorescence occurs because of a coupling of the fluorophore with the plasmon resonance excited at the metal surface. MEF is caused by increasing the excitation and emission rate of the fluorophore in a localized electromagnetic field. The effect of enhancement depends on the particle shape and size [43, 44]. The dye–metal particle systems interact with incident radiation in two modes: by scattering or absorption. The scattered component is responsible for the fluorescence enhancement for the large particles, while the absorption component that causes fluorescence quenching is dominant for the small particles. A MEF emission scheme and energy diagram is presented in Fig. 6. B

A ABS

Fl S1

Dye

ABS

Fl m Silver particle

Γ

Γm

k

S0

Fig. 6 Metal-enhanced fluorescence emission scheme (A) and energy diagram (B), adopted from [45]. ABS – Absorbance, Dye – Fluorescein [56], Fl – dye prompt fluorescence, Flm – additional emission from excited plasmons, – unmodified radiative decay rate, m – metal-modified radiative decay rate, k – nonradiative rates

Nonradiative energy transfer occurs from the excited fluorophores located at a certain distance to noncontinuous metallic films. The fluorescence emission is partially plasmon coupled and the system radiates [45]. Quantum yields m of the enhanced fluorescence signals of fluorophores in close proximity to metallic nanostructures could be described as: m = ( + m ) / ( + m + k) ,

(1)

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where is the unmodified radiative decay rate, m is the metal-modified radiative decay rate, and k is the nonradiative rate. Free-space quantum yield, 0 can be described as: 0 = / ( + k) .

(2)

Surface plasmon-coupled emission (SPCE) is similar to the phenomenon of SPR. In SPCE fluorophores in the excited state are coupled with metallic structures, which results in surface plasmons radiation into the substrate [46]. In SPCE the excited state fluorophore undergoes near-field interactions with a thin metal film. This interaction creates surface plasmons which radiate into the substrate at a well-defined angle. The effectiveness of the SPCE depends on the fluorophore distance and orientation relative to the metal surface [46]. The phenomenon occurs for fluorophores within 200 nm of a thin metal film making this a near-field phenomenon [47]. Many types of surfaces have been tested for MEF, including silver island colloid nanotriangles, nanorods, and even fractal-like silvered surfaces [45]. The MEF phenomenon is efficient when the fluorophore is approximately 10–20 nm from the metal island [48]. Enhancements have typically been observed for various organic fluorophores, such as fluoresceins, rhodamines, and cyanines [49–51], and also for inorganic fluorescent quantum dots and nanocrystals [52, 53]. The magnitude and wavelength of the plasmon peak depend on the average surface concentration of nanoparticles, as well as their sizes, shapes, and spatial distribution [54, 55]. Figure 7 shows the exemplary fluorescence enhancement spectra of fluorescein in a mixture with colloidal silver particles [56]. The measurements were made for fluorescein water solutions and its mixtures with silver particles, in the concentration range from 10−6 to 10−3 M. A significant growth of the fluorescence quantum yield in the presence of colloidal particles was only observed in the range of concentrations from 2·10−4 to 8·10−4 M. The SPR effect depends not only on the size of the metallic particles interacting with the dye but also on the distance between the dye molecule and the metal. The distance determines the nature of the phenomenon

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Fig. 7 Fluorescence enhancement spectra of fluorescein in mixture with colloidal silver particles, at dye concentration: (A) 4·10−4 M, (B) 5·10−4 M, adopted from [56]. Curves: (1) fluorescein (without silver), (2) fluorescein mixture with colloidal silver particles


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(fluorescence enhancement or quenching). The distances between the fluorescein molecules and the silver particles were within the range 50 to 1,000 nm. However, the above factors are not the only ones influencing the SPR effect, as the dye nature is also important. It has been proven by the corresponding tests for other dyes with large fluorescence quantum yield: rhodamine B and rhodamine 6G, carried out within the same concentration range. For these two dyes no significant changes in the fluorescence were observed after addition of colloidal silver to the solution [57, 58]. The purpose of the investigation was to determine the usefulness of fluorescein for the construction of sensors and indicators of metal presence in the environment. The possibility of controlling the fluorophore–metal coupling, excited state deactivation pathways and spatial distribution of the emission permits employment of the MEF in devices for fluorescence sensing. Recently, the use of fluorescently labeled metal particles has become attractive due to near-field interaction of the fluorophore with metal nanoparticles. The wavelength of plasmon resonance is closely relevant to the metal core including metal species, core size, and core shape as well as the dielectric properties of the coating layer. Fluorescence enhancement by metal particles is applied to increase the sensitivity of biological assays and increase the detectability and photostability of fluorophores [59]. A great challenge in immunoassay is to increase the sensitivity of DNA microarrays by MEF, using surface-bound metal nanoparticles [60].

3 Dye Fluorescence for Medical Photodynamic Study This section summarizes the fluorescence spectroscopy study of dyes in model systems whose results would permit the choice of sensitizers with the optimum photophysical properties fitted for particular medical purposes [61, 62–75]. The amount of energy absorbed by a given dye-sensitizer can be deactivated in radiative (fluorescence, delayed fluorescence, or phosphorescence) and nonradiative processes (internal conversion (IC), intersystem crossing (ISC), energy transfer, etc). By means of optical spectroscopy methods supported by photothermal techniques it is possible to estimate the efficiency of some of those processes and to draw information on the amount of energy used by the dye-sensitizer to initiate photochemical reaction and/or generate reactive oxygen species [63, 65, 70–74]. The dyes with efficiently occupied triplet states are usually very effective in photochemistry. Hence, the dye should be characterized, at least, by high photostability under prolonged illumination and high probability of ISC transition, to be well suited for photodynamic therapy (PDT). It seems that it is much simpler to find the optimum dyes for photodynamic diagnosis (PDD). First at all, they have to exhibit high yield of fluorescence emission to help establish the size and location of pathologically changed tissues. For medical applications the preliminary selection of photosensitizers and photomarkers can be made on the basis of study performed in vitro for dyes in model systems. The efficiency of two-step procedures on different groups of dyes

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was analyzed [63, 67, 70]. At first, the selection was made by comparing some spectral properties and parameters of dyes in organic solvents [63, 65, 69, 71–74]. As the dye properties depend strongly on their close surroundings and molecular intra- and interaction, for stilbazolium merocyanines (Mero’s) (obtained from Prof. I. Gruda) [70] and some tetrapyrrolic dyes [64, 66], the next step for promising dye-candidate selection was based on analysis of their interactions with macromolecules and/or with specially prepared mononuclear cells isolated from human blood.

3.1 Spectral Properties of Dyes and their Interactions in Model Systems The search for dyes most suitable for medical applications was in our study limited to two groups of dyes: polymethine dyes (Mero) [63, 70, 76] and cyclic tetrapyrrolic dyes (porphyrins, chlorines) [64–66, 69, 71–74]. Below we shall present as an example, the results concerning some Mero dyes (Fig. 8) and bacteriochlorophyll (BChl) c as well as its free-metal derivative bacteriopheophytin (BPhe) c.

X2 X1

N

( CH

O

CH ) n

X3

Mero B B* T*

X1 (CH2)11– OH (CH2)11– OH (CH2)11– OH

X2 H H OCH3

X3 H H H

W*

(CH2)15– CH3

OCH3

H

H I U*

(CH2)6– OH (CH2)6– OH (CH2)11– OH

H NO2 NO2

NO2 NO2 NO2

Fig. 8 Molecular structure and notation of stilbazolium merocyanine (Mero) dyes, ∗ – described salt form of dye

The type of side groups attached (Xi ) influences the spectral properties of Mero dyes. The dyes with H or OCH3 groups (Fig. 8) are soluble in ethanol and exhibit the absorption, fluorescence excitation, and emission spectra of very similar shapes (Figs. 9a and 10a) as well as similar Stokes shifts varying from 105 to129 (Table 3). The dyes comprising NO2 groups (Fig. 8) are soluble in methanol and exhibit the absorption and fluorescence spectra of different shapes (Figs. 9a and 10a), much different values of the full width at half maximum (FWHM) of the fluorescence and absorption bands and smaller Stokes shift (about 70–77 nm) (Table 3). It is clear that these dyes occur in several spectral forms characterized by different fluorescence yields. These species could be created by different interactions with the solvent and/or by mutual interaction of the dye molecules [63, 67, 68, 73]. The relative fluorescence yields of the dyes (Table 3) were calculated according to the methods described in [14] using coumarine as a reference dye [77]. These yield values, calculated for different conditions, for example in N2 , O2 , or air atmosphere, are necessary to obtain the information about the oxygen quenching of the dye singlet states as well as about the yields of (singlet-triplet) ISC transition.


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Fig. 10 Normalized fluorescence emission spectra of some Mero dyes in ethanol (A) and PVA film (B); λ exc – B: 400 nm, T∗ : 415 nm, U∗ : 426 nm (459 nm in PVA film), adopted from [44, 54]

The yield of fluorescence (F ) of the dyes soluble in methanol is much lower (almost zero) than that of the dyes with H or OCH3 groups (Table 3). The results obtained for Mero with the acceptor (NO2 ) groups could suggest the presence of various nonfluorescent, solvated or aggregated forms of dyes in the solution [63, 73]. The dyes with NO2 groups attached exhibit low yield of fluorescence therefore, from this point of view, they could be useful rather in photodynamic therapy than for diagnostic purposes. Using the fluorescence yields determined for Mero and the data obtained from photothermal measurement [laser-induced optoacoustic spectroscopy (LIOAS)] [34, 78–80] analysis, the T ET products can be calculated (Table 3) from the equation: T ET = (1 − α) Ehv − F Es ,

(1)

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Table 3 Spectral parameters of Mero dyes in alcohol solvents (B, B∗ , T∗ , W∗ in ethanol; H, I, U∗ in methanol) in contact with air atmosphere; A – absorption, F – fluorescence, FWHM – full width at half maximum, λmax – wavelength at maximum, λF-A – Stokes shift, F T , and ET – yield of fluorescence, triplet formation, and triplet state energy, respectively, based on [44, 54] A

F

Mero

FWHM λmax (nm) (nm)

FWHM λmax (nm) (nm)

F

λF-A (nm)

T ET (kJ/mol)

B B∗ T∗ W∗ H I U∗

400 400 414 414 432,459 428 429

504 503 543 543 536 498 500

0.16 0.13 0.30 0.28 ≈0 ≈0 0.01

106 105 129 129 77 70 71

116.4 107.7 97.2 43.5 65.5 160.0 8.7

59.4 57.0 64.2 62.9 92.1 63.5 75.7

60.3 60.3 76.4 75.6 86.2 79.7 51.0

where: T , ES , ET are the yield of triplet formation, singlet and triplet state energy of the dye, respectively, Ehν is the molar energy of the incident photons at the excitation wavelength used in photothermal (LIOAS) measurements, α (=k1 ) describes the part of energy exchanged promptly into heat, i.e. in a time shorter than 0.5 μs; the value was obtained, respectively, by Marti et al. [79] and deconvolution [80] methods of LIOAS signals. As follows from Table 3, some of the Mero dyes investigated exhibit high value of T ET product, which suggests that they should be effective in photodynamic reactions. The photochemical parameters (T ET ) obtained by means of the fluorescence and LIOAS analysis are difficult to correlate with the molecular structure of the Mero dyes [63, 73]. But, it seems that the complementary analysis of their optical spectra and photothermal signals of the dyes in organic solvents, give simple criteria for preliminary selection of the dyes from a huge group and help to find the best sensitizers for medical photodynamic treatment or diagnosis. By using polymer (PVA) as a simple model of biological membrane it was possible to predict the dye–cell interaction [63, 73, 76]. A particularly strong effect of the PVA presence in solution and in film on the absorption and fluorescence spectra (Figs. 9b and 10b) when compared to those taken in alcohol solutions has been observed only for Mero U∗ (Figs. 9a and 10a). The absorption maxima of the other Mero dyes in PVA solution were shifted in respect to those in PVA films (to the blue) but in PVA film (Fig. 10a) the positions of the main absorption bands are similar to those observed in alcohols (Fig. 9a). In the case of Mero H in a fluid solution of PVA, the absorption maxima exhibit some shift but the shapes of the spectra in methanol, fluid PVA, and PVA film in every case show at least the presence of two dye species irrespective of the surroundings (Figs. 9 and 10). The results show that properties of Mero dyes and their interactions with macromolecules depend strongly on the type of the side groups attached [63, 73]. As follows from a comparison of the spectral properties of the dyes in fluid and rigid PVA and in alcohol solutions, the interactions of different Mero dyes with


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macromolecular chains are very different and depend on the medium fluidity and the polymer presence. The strongest interaction with the polymer was observed for Mero U∗ . It is expected that this dye could interact with various components of cells and cell membranes [63]. The other Mero dyes that, as follows from absorption and fluorescence spectra (Figs. 9 and 10), do not enter into strong interactions with PVA polymer (except Mero U∗ ) could be promising candidates for staining blood cells. Such dyes should probably be easier retained inside the cells, because they would not be able to form efficient complexes with some components of the cell membrane (e.g. lipids, macromolecules). Mero U∗ because of its strong interaction with the polymer in solution as well as in film, seems to be unsuitable for cell staining [63]. On the basis of the optical spectroscopy measurements for the whole set of Mero dyes the effect of (1) the presence of salt (no difference observed for Mero B and B∗ ), (2) the attachment of NO2 groups in position X2 and X3 (from a comparison of Mero U∗ and I), (3) the length of the Mero chain (e.g. from the spectra of Mero T∗ and W∗ ) on the dye–polymer interactions has been excluded [63, 73]. The information obtained by optical spectroscopy methods supported by photothermal measurements should help in choosing proper candidates for applications in medicine. On the basis of the results presented in Figs. 9 and 10, Table 3, Mero B, B∗ , T∗ , and I, because of their spectral and photochemical properties (high T ET ) have been applied as promising sensitizers to stain blood cells (see Sect. 3.3). To estimate the value of the T parameter (Eq. 3.1) important for the dyesensitizer selection process, it is necessary to know from literature [81] or evaluate from independent experiments (e.g. phosphorescence spectra) the energy of the dye triplet state (ET ). For chlorin (BChl c and BPhe c) in monomeric form, the yield of fluorescence (F ) was established in alcohol solutions saturated with O2 , air, and N2 at various temperatures (not shown) [65, 69]. The F values (Table 4) were calculated using BChl c in acetone as a reference [82]. The F values obtained under oxygen and air atmosphere were similar within the calculation’s accuracy. These values are, as expected, the highest in nitrogen compared with those in air and in oxygen atmosphere, but they are independent of temperature (in the limit of the error). This may suggest that in the temperature range used, ethanol viscosity is stable and does not influence either the fluorescence or/and the ISC yields [69]. As follows from Table 4 the α-values of the dyes investigated depend also on the oxygen presence. The α parameter describes the prompt heat release, including vibrational relaxation Sn →S1 (IC), nonradiative deactivation S1 →S0 (IC), and S1 →T1 (ISC) transition [34, 69]. Using the F values and the preexponential factors (ki ) obtained from the deconvolution of LIOAS [80], the T and efficiencies of singlet oxygen generation ( ) were calculated (Table 4), from Eqs. (1) and (2) [34]:

T ET − Ehv k2 , = T S = E

(2)

where: E , S represent the electronic energy of singlet oxygen and the fraction of the dye triplet states quenched by oxygen, respectively.

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The calculated values of T for the pigments investigated show that BPhe c has higher yield of triplet state formation than BChl c. The effect of quenching of the pigment triplet states (Table 4) accompanied by a decrease in F as well as a rise in energy deactivated by fast thermal processes (α) has been observed for these pigments [65, 69]. The relatively high values of α as well as of the fluorescence yield for BChl c correspond to a lower yield of triplet state generation (Table 4). The BPhe c pigment exchanges less excitation energy into heat in a shorter time than BChl c (Table 4), which is highly fluorescent [65, 69]. Thus, it seems that BChl c could serve as a fluorescence probe for diagnostic purposes, but not as a sensitizer in PDT. It is known that the fluorescence yield of pigments in biological cells is about one order of magnitude lower than in most of the solvents [66, 81]. Because of the high fluorescence yield of BChl c it seems that after the pigment incorporation into the cells, the BChl c emission should be still reasonably measurable. The advantage of BPhe c is the high efficiency of triplet state formation; therefore this pigment could induce photochemical reactions during photodynamic treatment [65, 69]. Table 4 Photophysical parameters and yields obtained for BChl c and BPhe c in ethanol under air atmosphere, description of parameters as in Table 3.1; α – a fraction of excitation energy exchange into heat promptly (in shorter time than time resolution of apparatus), – yield of singlet molecular oxygen production, S – the fraction of triplet states quenched by oxygen, based on [46, 50] Dye

Atm.

F

α

T

S

BChl c

O2 air N2 O2 air N2

0.20 0.22 0.28 0.14 0.15 0.24

0.72 0.63 0.58 0.65 0.59 0.52

0.58

0.52

0.90

0.76

0.70

0.92

BPhe c

The results are expected to help establish which pigments could serve as potential sensitizers in further studies, in view of their potential applications in medicine. It should be realized that the fluorescence and triplet state generation yields in solvents could be different from those in the cells, therefore to verify this conclusion, the pigments incorporated into human lymphocyte cells have been studied (see Sect. 3.3).

3.2 Incorporation of Dyes into Cells To investigate the properties of dyes introduced into normal and pathological human cells, mononuclear cells were isolated from heparinized venous blood from healthy donors. The purity and number of cells (monocytes, granulocytes, and lymphocytes) were established using flow cytometry [66, 70]. It was estimated that the content of lymphocytes was about 75% of cells, henceforth the cells were referred to as lymphocytes. The lymphocytes obtained from healthy donors were the so-called


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resting (R) (normal) cells. One part of them was stimulated with phytohemagglutinin (PHA). The chemically modified (stimulated) (S) cells have one common feature, i.e. vigorous proliferation, and they can be used as a model for leukemia cells. In the dark, the dyes dissolved in alcohol were incubated into cells. The control samples were uncolored cells with addition of alcohol. During the incubation, the concentration of dyes in the sample volume was 10−4 M (for spectral investigations) and 10−4 –10−5 M (for estimation of cell apoptosis) [70]. The photographs of stained cells were taken using a fluorescence microscope. Some exemplary photographs of cancerous (JUCAT) cells stained by incubation with Mero B are shown in Fig. 11a. The pictures show that these dye molecules are gathered predominantly in the cell membrane [70]. A.

B. lymphocyt

C. granulocyt

Fig. 11 Photographs of (A) cancerous cells stained by Mero B taken under a fluorescence microscope (adopted from [8]) as well as of (B) lymphocyte and (C) granulocyte stained by BChl c observed under a confocal microscope, adopted from [54]

For all spectral and biochemical studies the colored and uncolored cells were resuspended in solutions of physiological salt. For detection of apoptosis the annexin-V (AV) conjugated fluorescein was used. For biochemical analysis by flow cytometry, the cells were incubated with AV-fluorescein and with propidium iodide (PI). PI is commonly used for the differentiation of apoptotic from necrotic cells.

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Fluorescence of the stained cells on a flow cytometer, using a 515-nm bandpass filter for fluorescein (FITC) emission detection and a filter > 600 nm for PI emission detection, was observed [70]. Table 5 shows some examples of results obtained for lymphocytes incubated with Mero sensitizers. The effect of staining by Mero B and Mero T∗ on the cell survival is shown in Table 5. As follows from Table 5, the staining of the resting cells as well as alcohol addition were, at the dye concentrations and illuminations used in the spectral investigations, only weakly cytotoxic. It was shown earlier [83] that the same dye was incorporated to a different degree of efficiency in various types of cells (lymphocytes, granulocytes) and that different Mero dyes exhibit different incorporation yields in similar types of cells. A similar conclusion follows from the pictures of stained mononuclear cells taken by the confocal microscopy method (Fig. 11b and c) [73]. Table 5 Effect of Mero B and T∗ on the structure of the resting lymphocyte membrane [measured by annexin binding (AV)] and survival (PI test – showed the part of necrotic cells) of the cells in light nonexposed samples, based on [51] Resting cells

Living cells (%)

AV – binding cells (%)

PI – test (%)

without alcohol with alcohol With alcohol and Mero B with alcohol and Mero T∗

99.70 99.45 97.15 96.11

0.10 0.26 0.74 1.26

715

S

680

760

τ3 (ps)

A3 (%)

39.5±2.6 22 42.8±1.9 25 55.7±2.7 75 65.3±3.4 40 46.5±3.4 18 54.2±2.6 20 71.2±5.9 13 30.3±2.5 48

χ2 2.69 1.18 2.79 1.17 5.81 1.38 5.36 1.91 4.23 1.08 3.89 0.95 2.00 1.25 5.99 1.88

Fluorescence Spectroscopy in Optoelectronics A

770

1000

500 Delayed Luminescence [counts]

Fig. 16 Time-resolved delayed luminescence spectra of Chl b in isotropic PVA at various temperatures: (A) 290 K; (B) 100 K; (C) 8 K (λexc = 425 nm), adopted from [144]. Time windows curve: (1) 0.5–5.5 μs; (2) 5.5–10.5 μs; (3) 10.5–15.5 μs; (4) 20.5–25.5 μs

267

690

0 B 1 2 x2 3 4

2000

770

1000 680 0 C

770

2000

1000 680 0 600

700 Wavelength [nm]

800

The green sulfur bacterium Prosthecochloris aestuarii contains giant lightharvesting complexes called chlorosomes. The main pigment of chlorosome is bacteriochlorophyll (BChl) c with a small admixture of BChl a. From the comparison of theoretical predictions with experiments such as spectral hole burning [128], fluorescence spectra [129], or other spectral results [120, 121] it follows that very strong intermolecular interactions of BChl c molecules impose their arrangement in chains located on the surface of the rod elements [120, 121]. The aim of this investigation is to explain the role of different pigment forms in excitation energy transfer and fluorescence emission. Fluorescence lifetime (τ ) was measured for several wavelengths of exciting and fluorescence light using the method of frequency domains [14]. The phase modulation lifetime measurements (frequency domain) were made for unstretched films under the magic angle conditions using the same front-face adapter. The excitation was provided by a mode-locked Ti-sapphire laser (Tsunami from Spectra Physics) (Fig. 14). The repetition rate of 80 MHz and power of about 1 W was divided down to 4 MHz. The Ti-sapphire laser output (780–900 nm) was frequency-doubled using an Inrad model 5-050 doubler/tripler [90]. Experimental data in the frequency domain lifetime measurements are usually [14] presented as increasing values of the frequency-dependent phase angle and

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decreasing values of the modulation. Examples of such curves obtained for whole bacteria samples for various excitations and 750 nm observation wavelengths are presented in Fig. 17. As follows from this example the fitting was very good. From the investigation of bacteria [108, 111] and model systems containing BChl c aggregates [113, 120–132] it is known that in organisms and in model systems there exist not only monomers of pigment and large oligomers but also several other forms (small aggregates – tetramers, BPhe c, etc.). The excitation wavelengths from the Soret band were used: 420 nm (BPhe c monomer absorption), 438 nm (monomers and small aggregates of BChl c absorption), and 450 nm (predominantly BChl c oligomer absorption). Fluorescence lifetime calculations were performed for the emission at 680 nm (emission of monomers and small aggregates of BChl c and BPhe c) and at 750 nm (predominantly BChl c oligomer fluorescence) [90]. The absorption and fluorescence bands of various forms of pigments strongly overlap. The way to improve the resolution of different lifetimes of the fluorescence emitted by such biological samples is global analysis of the lifetime results [14]. Global analysis is based on the assumption that the decay time of a given component is independent of fluorescence and excitation wavelengths. In our studies a simplifying assumption was made that only three emitting species are present in the sample. It is known that even oligomers occur at least in two different forms [133, 134]. Also small aggregates could be not uniform. Previous [108] lifetime results

Phase Angle [deg]/ Modulation [%]

100 80 60 40 20

Fig. 17 Frequency response of BChl c fluorescence intensity decay in PVA film observed at λem = 750 nm. The solid lines represent the best global analysis of three-exponential fit to the data at λexc = 450 nm (circles), λexc = 438 nm (small squares), and λexc = 420 nm (squares), adopted from [105]

Deviations Phase [deg] Modulation [%]

0 1 0 –1 1 0 –1 10

100 Frequency [Mhz]


269

Table 7 Fluorescence lifetime (experimental and global analysis data) for mixture of oligomers and other forms of BChl c in PVA, based on [105]. τ i – decay time of i-component, α i – amplitude, fi – fractional contribution Experimental data λexc λem [nm] [nm] 420

680

438 450 420 438 450

750

τ i [ns] α i

fi

2.969 0.101 4.394 0.163 0.256 0.013 0.216 0.176 0.163

0.005 0.995 0.002 0.998 0.061 0.939 1.000

0.135 0.865 0.051 0.949 0.567 0.433 1.000

0.163

17.7

Global analysis [ns]

χ2

τ i [ns] α i

fi

0.488

7.03

0.378

7.76

0.151

14.0

0.216 0.176

16.6 10.8

3.987 0.136 3.987 0.136 3.987 0.136 0.183

0.106 0.894 0.068 0.932 0.018 0.982 1.000

0.004 0.996 0.002 0.998 0.001 0.999 1.000

[ns] χ 2 0.545 26.4 0.397 0.204 0.183 27.0

obtained on a similar set of samples have shown that the decay times can be divided into three groups: the longest times are characteristic of monomers (from 5.6 to 7.4 ns), the intermediate times (1.1–2.7 ns) probably characterize small aggregates of BChl c, and the shortest times (0.001–0.37 ns) describe oligomers. We suppose for our analysis that the decay of fluorescence is a superposition of three or fewer exponential components. The simplest lifetime results are obtained for an artificial mixture of oligomers and other forms of BChl c in model systems in PVA. The decay of the fluorescence observed in the region of oligomeric emission (at 750 nm) can be sufficiently well described by the mono-exponential component, corresponding at various excitation wavelengths to τ values slightly differing from 0.2 ns (Table 7). Analysis of the polarization absorption spectra (not shown) implies that there must be at least two oligomeric forms, but they have to have similar lifetimes. Global analysis gives the value of this decay equal to 0.183 ns, but the accuracy of the fit is rather low. Previously [108] the largest decay component of this emission was found to have a lifetime from 0.001 ns till 0.01 ns for various wavelengths of light excitations. In our study a strong component with a short decay time is observed at 680 nm, while the global analysis revealed the strong component with τ = 0.136 ns together with a much weaker component with a longer lifetime (about 4 ns, Table 7). The light polarization does not critically change the contributions from various decay components for the unstretched film. As follows from our results, in both regions of observation the strongest component has a short lifetime ranging from 0.136 to 0.183 ns. The intensity of emission from monomers and small aggregates is comparable with that of the emission from oligomers (Fig. 17). The longer lifetime component is of course more easily seen in steady-state emission spectra, but f i are low for 680 nm observation. This suggests strong excitation energy transfer from monomers to oligomers responsible for shortening of the monomeric lifetimes [111].

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Lifetime measurements of bacteria whole cells and bacteria fragments in PVA were also carried out. The fluorescence of bacteria and bacteria fragments has to be approximated by three different lifetime components. They are different for whole bacteria and for their fragments and for observations at 680 and 750 nm. It clearly shows that in contrast to the ordered model systems, the pigments located in the organisms give slightly different lifetimes for each type of pigment aggregates because of their different environment [90]. Measurement of fluorescence lifetimes using time-resolved fluorescence quenching techniques were very convenient for investigation of aggregation properties of fluorophores in model systems composed of two or more fluorophores in various forms of aggregation [122, 135]. Mixtures of chlorophyll a (Chl a) and chlorophyll b (Chl b) in anisotropic PVA films at different concentration ratios of Chl a to Chl b were made [122]. In PVA, the pigment aggregates are much more stable than in fluid solutions and can be oriented to enable their extensive investigation by polarized light spectroscopy. Fluorescence lifetimes of Chl a to Chl b mixtures were measured using single photon counting apparatus [122] and analyzed on multi-exponential components fit. Fluorescence decays of various aggregates are complex and have been best approximated by two or three exponents. Analysis of the decay curves has shown that at high Chl a and low Chl b concentrations the following three forms of Chl a aggregates occurred: “dry” Chl a (aggregated with polymer) with a fluorescence maximum at 676 nm (F676), hydrated dimer (Chl a 2H2 O)2 or (Chl a H2 O)2 (F700– 705 nm), and oligomers of hydrated dimers (F750–770 nm). Chl b in the same matrix also occurs in at least three forms: “dry” (F653 nm), “wet”-hydrated aggregates (F662 nm), and aggregated forms with the fluorescence at 710–720 nm and/or 730–740 nm regions. In all cases water plays an important role in chlorophyll aggregation [136–138]. The efficiency of excitation energy transfer from Chl b to various aggregates of Chl a is different. The yield of the energy transfer from Chl b to Chl a oligomers is high. The results are discussed on the basis of previous data concerning the monolayers of Chl a and literature data concerning aggregation of chlorophylls in various model systems. As follows from the analysis of fluorescence decay, the shortest lifetimes (0.14–0.28 ns) are for Chl a oligomers and aggregates of Chl b (0.10–0.16 ns). A strong component of the lifetime 2.8 ns is related to hydrated dimers of Chl a. The longest fluorescence lifetimes belong to monomer and “wet” forms of both pigments. In the above experiment, the TCSPC technique enabled identification of the types of pigment aggregates. This section presented a brief review of the advanced techniques of time-resolved fluorescence quenching and application of ultrafast fluorescence spectroscopy to the study of dynamic light-induced inter- and intramolecular deactivation processes in molecular systems. Some similarities between the aggregated forms or chlorophylls and pigment forms observed in organisms are indicated. In polymer films, aggregated forms of pigments are stable and oriented; therefore, their properties and the conditions of their formation can be established. It is possible to find several common properties of various types of chlorophyll aggregates occurring in the model systems used even when exact positions of the spectral bands and values of


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the lifetimes of emission vary with the type of model and conditions of sample preparation. Therefore, these common features can be extrapolated to in vivo conditions. TCSPC is a powerful technique, very useful for establishing fluorescence lifetimes, which permit identification of various molecular individuals.

5 Summary The studies reviewed in this chapter very well illustrate how steady-state fluorescence spectroscopy can be used to design organic materials with the most optimal properties needed for specified applications. We have shown that fluorescence spectroscopy represents a helpful tool in determination of some molecular processes occurring in both simple and complex organic materials for their applications in novel sciences, technology, medicine, and bioscience. The fluorescence behavior of several dye families has been used to show and discuss the molecular phenomena essential in photovoltaics, medical diagnosis, and investigation of pigment/dye properties in vivo and in model systems. It has been shown that an understanding of some physical mechanisms on the molecular level with fluorescence spectroscopy is a prerequisite for an interpretation of the lightinduced processes like: dye intra- and interactions, excimer formation, electron and energy transfer, covalent modification changes in electron distribution, photodegradation processes, and plasmon resonance of dye–metal particle systems. Knowledge of the features of the dye molecules excited state, fluorescence decay kinetics, and kinetics of the excited state’s deactivation in radiative processes can help in understanding how a particular material property determines parameters important for the construction of novel photovoltaic devices and a new generation of photosensitizers. Investigation of fast energy deactivation kinetics can also help to understand complex molecular processes that occur in antenna systems and reaction centers of photosynthetic organisms. Acknowledgments The authors are grateful for research grant support: Polish Ministry of Education and Higher School, project 2008–2011 and the Poznan University of Technology. D.W. greatly appreciates the assistance of MSc Eng. Kornelia Lewandowska and Andrzej Wróbel for their help in preparation of the figures.

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Multicolor Imaging with Fluorescent Proteins in Mice In Vivo Imaging with Fluorescent Proteins Robert M. Hoffman

Abstract The high extinction coefficients, quantum yields, and unique spectral properties of fluorescent proteins are optimal for imaging in live animals in real time. The important aspects of cancer in living animals, including tumor cell mobility, invasion, metastasis, and angiogenesis, can be imaged. Fluorescent proteins enable whole-body imaging of tumors on internal organs. Proteins which are spectrally distinct have allowed the color coding of cancer cells growing in vivo with distinction of different cell types, including host from tumor, with single-cell resolution. Individual cells can be multicolored to study nuclear–cytoplasmic dynamics in the living animal.

1 Introduction 1.1 GFP and Other Fluorescent Proteins as Imaging Agents GFP and related fluorescent proteins are a homologous family, having emission spectra from 442 to 645 nm [1]. This family of proteins range in size from 25 to 30 kDa. Fluorescent proteins form internal chromophores that do not require cofactors or substrates in order to fluoresce. These fluorescent proteins have very high extinction coefficients ranging up to approximately ε = 95,000. In addition, they have very high quantum yields ranging up to 0.8 [2]. These properties make fluorescent proteins exceedingly bright. GFP’s large two-photon absorption is important for in vivo applications [2]. Another important feature is the spectral distinction of many members of the family of fluorescent proteins. Therefore, a set of multicolor fluorescent proteins can be utilized simultaneously for multifunctional in vivo R.M. Hoffman (B) AntiCancer, Inc., San Diego, CA 92111 and Department of Surgery, University of California, San Diego, CA 92103, USA e-mail: [email protected]


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imaging. Fluorescent proteins require no substrate, are nontoxic, stable at 37◦ C, and are relatively resistant to photo-bleaching [3]. These properties make fluorescent proteins optimal for cellular in vivo imaging.

1.2 First Use of Fluorescent Proteins in Animals to Visualize Tumor Cells Fluorescent proteins were first used in animals by Chishima et al. [4] to visualize cancer cells. Fluorescent proteins can be used to image, including whole-body imaging, essentially any type of cancer process including primary tumor growth, tumor cell motility, tumor cell invasion, intravasation, extravasation, metastatic seeding, metastatic colonization, angiogenesis, cell deformation, drug sensitivity, effects of molecular alterations, and tumor–host interaction. Many colors of fluorescent proteins are now known and can be used to color code cancer cells of a specific genotype or phenotype. Upon co-implantation in a single animal, the interaction between the variants can be visualized. For example, a highly metastatic variant can be labeled with GFP, and its low metastatic variant can be labeled with red fluorescent protein (RFP). The host and the tumor can be differentially labeled with fluorescent proteins. The host mouse can be a transgenic organism expressing GFP in all of its cells or specifically in cells of interest such as endothelial cells. The GFP-expressing mice can be transplanted with cells expressing RFP. Interaction of the tumor cells with the host cells can then be imaged. Cells themselves can be double labeled with GFP in the nucleus and RFP in the cytoplasm. With such double-labeled cells, nuclear–cytoplasmic dynamics can be imaged. In addition to cancer, many other processes can be visualized in vivo with fluorescent proteins including gene expression [5], infection [6], and behavior of stem cells [7, 8].

1.3 First Use of Fluorescent Proteins for Whole-Body Imaging GFP was first used for whole-body imaging by Yang et al., who visualized primary and metastatic tumor growth in real time [5, 9]. The intrinsic brightness of fluorescent proteins enables cells to be visualized inside intact animals by whole-body imaging. GFP-expressing primary and metastatic tumor growth could be visualized by whole-body imaging in numerous organs [9]. An RFP-expressing human pancreatic tumor cell line was introduced as tissue fragments into the pancreas of nude mice by surgical orthotopic implantation (SOI). As the tumors were growing, whole-body optical imaging was used to track, in real time, the growth of the primary tumor and the formation of metastatic lesions that developed in the spleen, bowel, portal lymph nodes, omentum, and liver. The images were used for quantification of tumor growth in each of these organs [10]. Tumor area measured by GFP imaging correlated with tumor volume measured in

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the opened animal, which validates the use of whole-body imaging to quantitate tumor growth and metastasis. Whole-body imaging with this model was used to compare standard and experimental agents for pancreatic cancer [11, 12]. Peyruchaud and colleagues [13] established a GFP-expressing bone metastasis subclone of MDA-MB-231 (B02/GFP.2) by repeated in vivo passages in bone by using the heart injection model. When injected into the tail vein of mice, the selected cells grew preferentially in bone. Whole-body fluorescence imaging of live mice showed that bone metastases could be detected about 1 week before radiologically distinctive osteolytic lesions developed. Furthermore, when the tumor-bearing mice were treated with a bisphosphonate, the progression of established osteolytic lesions and the expansion of breast cancer cells within bone were inhibited. Using wholebody GFP imaging, Peyruchaud et al. [14] showed that the angiogenesis inhibitor angiostatin greatly inhibited tumor growth in bone by inhibiting osteoclast activity. Using a different approach, human ovarian tumor cells (SKOV3.ip1) were made to express GFP by infecting with a replication-deficient adenoviral (Ad) vector encoding GFP [15]. The infected cells showed high GFP fluorescence, and when implanted into mice, intraperitoneal tumors as small as 0.2 mm in diameter could be detected by whole-body imaging within 24 h [16]. In another study [17], however, GFP-expressing tumors could not be detected by whole-body imaging until 7 days of s.c. tumor cell inoculation. These results strongly contrast with the results described above. This discrepancy shows the need to use bright cells and appropriate instrumentation and techniques for whole-body fluorescence imaging. 1.3.1 Whole-Body Imaging of Gene Expression Transgene expression in intact animals can be visualized by whole-body imaging. GFP expressed in the cells of brain, liver, pancreas, prostate, and bone was visualized by whole-body imaging. Both nude and normal mice were labeled by directly injecting adenoviral GFP. Within 5–8 h of adenoviral GFP injection, the fluorescence of the expressed GFP in brain and liver became visible, and whole-body images were recorded at video rates. The GFP fluorescence continued to increase for at least 12 h and remained detectable in liver for up to 4 months. The method requires only that the expressed gene or promoter be fused or operatively linked to GFP [5]. 1.3.2 Whole-Body Imaging of Angiogenesis Angiogenesis can be whole-body imaged in GFP-expressing tumors with fluorescent proteins. The nonluminous induced capillaries generated from the host are clearly visible against the very bright tumor fluorescence when examined by wholebody imaging. Whole-body imaging of tumor angiogenesis was demonstrated by injecting GFP-expressing Lewis lung carcinoma cells into the s.c. site of the footpad of nude mice. The footpad is relatively transparent, with comparatively few resident blood vessels, allowing quantitative imaging of tumor angiogenesis in the intact

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animal. Capillary density increased linearly over a 10-day period as determined by whole-body imaging. Similarly, the GFP-expressing human breast tumor MDAMB-435 was orthotopically transplanted into the mouse fat pad, where wholebody optical imaging showed that blood vessel density increased linearly over a 20-week period. The GFP-expressing angiogenesis mouse models can be used for real-time in vivo evaluation of agents inhibiting or promoting tumor angiogenesis in physiological microenvironments [18].

1.3.3 Whole-Body Imaging of Bacterial Infection The luminance of GFP-expressing bacteria enabled whole-body imaging in intact infected animals. This simple, nonintrusive technique can show in great detail the spatial–temporal behavior of the infectious process. The bacteria, expressing the GFP, are sufficiently bright as to be clearly visible from outside the infected animal and recorded with simple equipment. Introduced bacteria were observed in several mouse organs including the peritoneal cavity, the stomach, the small intestine, and the colon. Instantaneous real-time images of the infectious process were acquired by using a color charge-coupled device video camera by simply illuminating mice at 490 nm. The whole-body fluorescence imaging described here is fast and requires no extraneous agents. The progress of Escherichia coli GFP through the gastrointestinal tract of the mouse after gavage was followed in real time by whole-body imaging. Bacteria, seen first in the stomach, migrated into the small intestine and subsequently into the colon, an observation confirmed by intravital direct imaging. An i.p. infection was established by i.p. injection of E. coli GFP. The development of infection over 6 h and its regression after kanamycin treatment were visualized by whole-body imaging. This imaging technology affords a powerful approach to visualize the infection process and determine the tissue specificity of infection and the spatial migration of the infectious agents [6]. Zhao et al. [19] showed that genetically modified bacteria could selectively kill tumors in mice. A genetically modified bacterial strain, Salmonella typhimurium A1, auxotrophic for Leu and Arg, was labeled with GFP. The GFP-expressing bacteria grew in the cytoplasm of PC-3 human prostate cancer cells and caused nuclear destruction. These effects were visualized in cells labeled with GFP in the nucleus and RFP in the cytoplasm. In vivo, the bacteria caused tumor inhibition and regression of xenografts visualized by whole-body imaging with the PC-3 prostate cancer cells labeled with RFP grafted into nude mice. The GFP bacteria continued to proliferate in the PC-3 tumor, which stopped growing but did not continuously grow in normal tissue [19]. In another approach, Yu et al. [20] have shown that GFP-expressing bacteria injected intravenously into live animals entered and replicated in solid tumors and metastases. The tumor-specific amplification process was visualized in real time using GFP fluorescence, which revealed the locations of the tumors and metastases. Localization of tumors was visualized by the GFP expressed by the emitting


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microorganisms in immunocompetent and in immunocompromised rodents with syngeneic and allogeneic tumors.

1.3.4 External Imaging Through Skin Flaps Opening a reversible skin flap in the light path markedly reduces signal attenuation, increasing detection sensitivity manyfold. The observable depth of tissue is thereby greatly increased. For example, GFP-expressing pancreatic tumors and their angiogenic microvessels were externally imaged by means of a peritoneal-wall skin flap [21].

1.3.5 Whole-Body Imaging of Graft–Versus-Host Disease Panoskaltsis-Mortari et al. [22] developed a mouse system to track the migration and homing of cells in a setting of bone marrow transplantation (BMT)-induced graft-versus-host disease (GVHD) after systemic infusion using GFP transgenic cells. Whole-body images taken at various time points after cell infusion revealed the early migration of allogeneic cells to peripheral lymphoid organs, with later infiltration of GVHD target organs. Localization of GFP-expressing cells could be seen through the skin of shaved mice, and internal organs were easily discernible. Infusion of different cell populations revealed distinct homing patterns. Whole-body imaging provided a simple way to identify the critical time points for expansion of the transplanted cells in various organs.

1.4 Autofluorescence Is Not a Problem with Fluorescent Protein-Based In Vivo Imaging The beauty of fluorescent proteins is their brightness, availability in multiple colors with distant spectra, and simplicity of use. The intrinsic fluorescence is so bright that very simple equipment can be used for many in vivo studies of tumor growth and metastasis, drug sensitivity, and angiogenesis. The strong signal from the fluorescent proteins readily distinguishes it from autofluorescence of the skin or other organs. For screening large changes in fluorescence and other macro imaging studies, equipment as simple as LED flashlights with an appropriate excitation filter and a simple emission filter is sufficient [23, 24]. The size of the whole-body imaged tumor was comparable to open images. Even more striking was that the intensity of the whole-body image was 70% of the open image. Although some information is lost with whole-body imaging due to light scattering, a remarkable amount of information was obtained, even with such simple instrumentation. More highly sophisticated equipment including highly sensitive color CCD cameras as well as dual-photon lasers can be used for ultra high-resolution in vivo imaging of fluorescent protein expression [25]. Software that can automatically

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identify areas of fluorescence in the animals and quantify the fluorescent area and intensity has been developed. The use of tunable filters allows the isolation of any individual spectrum in any fluorescent pixel. This technique eliminates autofluorescence as well as enables high-resolution spectral distinction when multiple fluorescent proteins are being used or when the fluorescence signal emanates from deep in the animal. Spectral resolution enables, for example, high-resolution whole-body visualization of tumor blood vessels [26].

1.5 Use of Whole-Body Imaging with Fluorescent Proteins to Measure Drug Response in Real Time A novel, p.o.-administered cytosine analogue, CS-682, was shown to inhibit metastatic growth in the MIA-PaCa-2-RFP orthotopic mouse model of pancreatic cancer. This model has two important features that make it an ideal system to study the effects of CS-682. First, it is a highly malignant orthotopic model that, left untreated, spontaneously gives rise to extensive dissemination to the lymphatics, peritoneum, and solid organs, as well as ascites. Second, its high level of selective tumor RFP fluorescence facilitates in vivo visualization and quantification of tumor growth and metastasis development. The model, therefore, allows real-time evaluation of the drug on pancreatic cancer without the need for laparotomy, substrates, contrast agents, or other invasive procedures [12]. Tumor RFP fluorescence facilitated real-time, sequential imaging and quantification of primary and metastatic growth and dissemination in vivo. Mice were treated with various p.o. doses of CS-682 on a five times per week schedule until death. At a dose of 40 mg/kg, CS-682 prolonged survival compared with untreated animals (median survival 35 days versus 17 days; P = 0.0008). At nontoxic doses, CS-682 effectively suppressed the rate of primary tumor growth. CS-682 also decreased the development of malignant ascites and the formation of metastases, which were reduced significantly in number in the diaphragm, lymph nodes, liver, and kidney. Selective RFP tumor fluorescence enabled noninvasive real-time comparison between groups during treatment and facilitated identification of micrometastases in solid organs at autopsy. Thus, we have demonstrated that CS-682 is an efficacious antimetastatic agent that significantly prolongs survival in an orthotopic model of pancreatic cancer [12]. A potential therapeutic agent for breast cancer based on galectin-3, an NH(2)terminally truncated form of galectin-3 (galectin-3C), was found to be efficacious in reducing tumor growth and inhibiting metastases. For efficacy studies, nude mice bearing orthotopically implanted tumors derived from breast cancer cell line MDAMB-435 expressing GFP were treated with galectin-3C or a vehicle control i.m. twice daily for 90 days. Mean tumor volumes were statistically significantly less in mice treated with galectin-3C compared with control mice, and fewer numbers of mice exhibited lymph node metastases in the treated group compared with the control group as visualized with GFP imaging [27].


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1.6 Imaging the Relationship of Tumor Cells and Blood Vessels Following injection of tumor cells stably expressing GFP into the tail vein of mice, it was possible to visualize single tumor cells in blood vessels [28]. Huang [29], Li [30], and their respective coworkers visualized GFP tumor cell–vessel interaction by using skin window chambers in rodents and observed angiogenic effects very early in tumor colony formation. When as few as 60–80 tumor cells were present, increased vasodilation and vessel morphology were observed. When as few as 100 cells were present, neovascularization was induced. Moore and colleagues have also visualized vessels in a GFP-expressing rodent cell line [31]. Al-Mehdi et al. [32] and Wong et al. [33] observed the steps in early hematogenous metastasis of tumor cells expressing GFP in subpleural microvessels in intact, perfused mouse and rat lungs. Metastatic tumor cells attached to the endothelia of pulmonary pre-capillary arterioles and capillaries. Extravasation of tumor cells was rare. Early tumor colony formation was observed entirely within the blood vessels. Rat tongue carcinoma cell lines expressing GFP have been used to investigate the formation of micrometastasis. The cells were injected into the portal vein and then tracked by using intravital video microscopy [34]. The two cell types—LM-GFP metastatic and E2-GFP nonmetastatic tongue carcinoma cells—were immediately arrested in the sinusoidal vessels near terminal portal venules. The E2-GFP cells disappeared from the liver sinusoid within 3 days, whereas a substantial number of LM-GFP cells remained in the liver—possibly because these cells formed stable attachments to the sinusoidal wall. Upon examination of the process with a confocal laser scanning microscope, only LM-GFP cells were shown to grow in the liver. Mook et al. [35] noted that initial arrest of colon cancer cells in sinusoids of the liver was due to size restriction after injection of the CC531S-GFP rat tumor cell line. Sturm et al. [36] injected GFP-expressing mouse colon 26 tumor cells into the spleen of immunocompetent BALB/c mice. Some of the tumor cells were trapped in the presinusoidal vasculature as well as in the sinusoids. Some of the tumor cells were attached to the vessel wall and some were seen to extravasate. Wang et al. [37] visualized the trafficking of metastatic cells targeting the liver via the portal vein using GFP-expressing cancer cells. Within 72 h of transplantation on the ascending colon in nude mice, metastasis was visualized ex vivo on a single-cell basis around the portal vein by GFP imaging. Brown et al. [25] showed that multiphoton laser scanning microscopy could provide high–resolution, three-dimensional images of angiogenesis-related gene expression and that this technique could be used to investigate deeper regions of GFP-expressing tumors in dorsal skin-fold chambers. To monitor the activity of the vascular endothelial growth factor (VEGF) promoter, Fukumura et al. [38, 39] made transgenic mice that express GFP under the control of the VEGF promoter. Multiphoton laser scanning microscopy showed that the tumor was able to induce the activity of the VEGF promoter and subsequent vessel formation.

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Wong and colleagues [40] showed that death of transformed, metastatic, rat embryo cells, which were expressing GFP, occurred via apoptosis in the lungs 24–48 h after injection into the circulation. Bcl-2 overexpression conferred apoptosis resistance for 24–48 h after injection. This inhibition of apoptosis led to a greater number of macroscopic metastases. Chang and coworkers [41] used CD31 and CD105 to identify endothelial cells and GFP labeling of tumor cells. These studies showed that colon carcinoma xenografts had mosaic vessels with focal regions where no CD31/CD105 immunoreactivity was detected and tumor cells appeared to contact the vessel lumen. Wyckoff et al. [42] used metastatic (MTLn3) and nonmetastatic (MTC) cell lines derived from the rat mammary adenocarcinoma 13762 NF, expressing GFP to measure tumor cell density in the blood, individual tumor cells in the lungs, and lung metastases. Metastatic cells showed greater orientation toward blood vessels, whereas nonmetastatic cells fragmented when interacting with vessels. A major difference in intravasation between metastatic and nonmetastatic cells was thus visualized by GFP in the primary tumor. Ahmed et al. [43] used transgenic mice expressing GFP driven by the mouse mammary virus promoter (MMTV). This model was used for imaging of transgenic mammary tumors and metastases at the single-cell level of resolution. Glinskii et al. [44] reported here that GFP-expressing human prostate carcinoma growing orthotopically efficiently deliver viable metastatic cells to the host circulation. This is in contrast to the ectopic tumors of the same lineage, which do not deliver live cells into the circulation. Coinjection of an equivalent mixture of isolated and cultured circulating GFP-expressing clones and parental RFP-expressing human prostate carcinoma cells revealed that the selected GFP-labeled viable circulating cells have an increased metastatic propensity relative to the RFP-labeled parental cells. The identification and isolation of highly malignant viable circulating human prostate carcinoma cells from orthotopic but not ectopic models suggests that the tumor microenvironment plays a very important role in enabling metastasis to occur. Berezovskaya et al. [45] have now demonstrated that the metastatic human prostate carcinoma cells selected for survival in the circulation have increased resistance to anoikis, which is apoptosis induced by cell detachment. Increased expression of the apoptosis inhibitory protein XIAP contributes to this anoikis resistance of the circulating metastatic human prostate carcinoma cells and thereby to their ability to form distant metastasis.

1.7 Visualizing Cellular and Nuclear Deformation and Dynamics in Small Blood Vessels Yamamoto et al. [46] have genetically engineered dual-color fluorescent cells with one color in the nucleus and the other in the cytoplasm that enable real-time nuclear– cytoplasmic dynamics to be visualized in living cells in vivo as well as in vitro.


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Nuclear GFP expression enabled visualization of nuclear dynamics, whereas simultaneous cytoplasmic RFP expression enabled visualization of nuclear–cytoplasmic ratios as well as simultaneous cell and nuclear shape changes. Common carotid artery injection of dual-color cells and a reversible skin flap enabled the external visualization of the dual-color cells in capillaries in the mouse brain, where extreme elongation of the cell body as well as the nucleus occurred in order for the tumor cells to enter the capillaries.

1.8 Imaging Tumor Cell Deformation and Migration in Blood Vessels of Live Mice in Real Time Tumor cells expressing GFP in the nucleus and RFP in the cytoplasm were injected into the heart of nude mice. A skin flap on the abdomen was made and spread on a flat stand. Highly elongated cancer cells in capillaries in the skin flap in living mice were visualized. The cells in the capillaries elongated to fit the width of these vessels. The average length of the major axis of the cancer cells in the capillaries increased to 3.97 times their normal length. The nuclei increased their length 1.64 times in the capillaries. Cancer cells that were arrested in capillaries over 8 μm in diameter could migrate up to 48.3 μm/h. The data suggest that the minimum diameter of capillaries where cancer cells are able to migrate is approximately 8 μm. Dual-color cells could be visualized in larger vessels as well with an occasional cell extravasating [47]. Extravasated cancer cells were also visualized to become intimately involved with blood vessels. Fluorescent protein imaging can visualize the cellular and nuclear dynamics of tumor cells within and associated with blood vessels in real time [47].

1.9 Color Coding of Cancer Cells Determines Clonality of Metastasis Yamamoto et al. [48] described the use of GFP-labeled or RFP-labeled HT-1080 human fibrosarcoma cells to determine clonality by simple fluorescence visualization of metastatic colonies after mixed implantation of the red and green fluorescent cells in SCID mice. Resulting pure red or pure green colonies were scored as clonal, whereas mixed yellow colonies were scored as nonclonal. In a spontaneous metastasis model originating from footpad injection in severe combined immunodeficient mice, 95% of the resulting lung colonies were either pure green or pure red, indicating monoclonal origin, whereas 5% were of mixed color, indicating polyclonal origin. In an experimental lung metastasis model established by tail vein injection in SCID mice, clonality of lung metastasis was dependent on cell number. The lung metastases were also visualized by external fluorescence imaging in live animals through skin-flap windows over the chest wall. Lung metastases were observed on the lung surface of all mice. SCID mice well tolerated multiple surgical procedures for direct-view imaging via skin-flap windows. Real-time metastatic

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growth of the two different colored clones in the same lung was externally imaged with resolution and quantification of green, red, or yellow colonies in live animals. The color coding enabled determination of whether the colonies grew clonally or were seeded as a mixture with one cell type eventually dominating, or whether the colonies grew as a mixture. The simultaneous real-time dual-color imaging of metastatic colonies gives rise to the possibility of color-coded imaging of clones of cancer cells carrying various forms of genes of interest [49].

1.10 Imageable Tumor–Host Models A transgenic GFP nude mouse with ubiquitous GFP expression has been developed. The GFP nude mouse was obtained by crossing nontransgenic nude mice with the transgenic C57/B6 mouse in which the β-actin promoter drives GFP expression in essentially all tissues [50]. In the adult mice, all the organs brightly expressed GFP, including the heart, lungs, spleen, pancreas, esophagus, stomach, and duodenum. The skinned skeleton highly expressed GFP. Pancreatic islets showed GFP fluorescence. The spleen cells were also GFP positive. RFP-expressing human cancer cell lines, including PC-3-RFP prostate cancer, HCT-116-RFP colon cancer, MDA-MB-435-RFP breast cancer, and HT1080-RFP fibrosarcoma, were transplanted into the transgenic GFP nude mice. All of these human tumors grew extensively in the transgenic GFP nude mice [51]. These models show the details of the tumor–stroma interaction, especially tumor-induced angiogenesis and tumorinfiltrating lymphocytes. The GFP-expressing tumor vasculature, both nascent and mature, of the GFP host mouse could be readily distinguished interacting with the RFP-expressing tumor cells. GFP-expressing dendritic cells were observed contacting RFP-expressing tumor cells with their dendrites. GFP-expressing macrophages were observed engulfing RFP-expressing cancer cells. GFP lymphocytes were seen surrounding cells of the RFP tumor, which eventually regressed [52]. With fluorescent protein imaging we are now able to visualize the cell–cell interactions of tumor and stroma in the living state. Li et al. [7] and Amoh et al. [8] have shown that the neural-stem cell marker nestin is expressed in hair follicle stem cells and the blood vessel network interconnecting hair follicles in the skin of transgenic mice with nestin-regulatoryelement-driven GFP (ND-GFP). The hair follicles were shown to give rise to the nestin-expressing blood vessels in the skin. Tumor angiogenesis was visualized by dual-color fluorescence imaging in ND-GFP transgenic mice after transplantation of the murine melanoma cell line B16F10 expressing RFP. ND-GFP was highly expressed in proliferating endothelial cells and nascent blood vessels in the growing tumor. Results of immunohistochemical staining showed that the blood vessel-specific antigen CD31 was expressed in ND-GFP-expressing nascent blood vessels. Progressive angiogenesis was readily visualized during tumor growth by GFP expression. RFP tumor cells were visualized inside ND-GFP blood vessels. Doxorubicin inhibited the nascent tumor angiogenesis as well as tumor


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growth in the ND-GFP mice transplanted with B16F10 GFP [53]. These new dual-color imageable models of tumor angiogenesis enable new insights into this process. The ND-GFP gene was crossed into nude mice on the C57/B6 background to obtain ND-GFP nude mice. ND-GFP was expressed in the brain, spinal cord, pancreas, stomach, esophagus, heart, lungs, blood vessels of glomeruli, blood vessels of skeletal muscle, testes, hair follicles, and blood vessel network in the skin of NDGFP nude mice. Human lung cancer, pancreatic cancer, and colon cancer cell lines as well as a murine melanoma cell line and a breast cancer tumor cell line expressing RFP were implanted orthotopically and an RFP-expressing human fibrosarcoma was implanted subcutaneously in the ND-GFP nude mice. These tumors grew extensively in the ND-GFP mice. ND-GFP was highly expressed in proliferating endothelial cells and nascent blood vessels in the growing tumors, visualized by dual-color fluorescence imaging. The ND-GFP transgenic nude mouse model enables the visualization of nascent angiogenesis in human and mouse tumor progression. These results suggest that this model is useful for the imaging of the angiogenesis of human as well as rodent tumors and the visualization of the efficacy of angiogenetic inhibitors [54]. Duda et al. [55] noted at the time of transplantation that tumor fragments contain “passenger” cells: endothelial cells and other stromal cells from the original host. They investigated the fate of GFP-labeled endothelial and nonendothelial stromal cells after transplantation in syngeneic mice. Angiogenic stroma associated with tumor or adipose tissue persisted when transplanted, remained functional, and governed the initial neovascularization of grafted tissue fragments for more than 4 weeks after implantation. The passenger endothelial cells survived longer than other stromal cells, which are replaced by host-activated fibroblasts after 3 weeks. The transplantability of tumor stroma suggests that the angiogenic potential of a tumor xenograft depends on the presence of passenger endothelial cells and other stromal cells within the xenograft.

1.11 Advantages of GFP Imaging Over Luciferase and Other Optical Imaging Techniques The GFP approach has several important advantages over other optical approaches to imaging. In comparison with the luciferase reporter, GFP has a much stronger signal and therefore can be used to image unrestrained animals. The fluorescence intensity of GFP is very strong since the quantum yield is about 0.8 [56–60]. The protein sequence of GFP has also been “humanized,” which enables it to be highly expressed in mammalian cells [61]. In addition, GFP fluorescence is fairly unaffected by the external environment since the chromophore is protected by the three-dimensional structure of the protein [62]. The excitation wavelength is quite long at 490 nm [57–60], which does not quench the fluorescence and therefore longterm measurements can be made. In vivo, GFP fluorescence is mainly limited by

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light scattering, which, as noted above, can be overcome by spectral resolution [26], skin flaps [21], and endoscopes [63–65] such that single cells can be imaged externally. Longer wavelength fluorescence proteins, such as RFP, can also be used to reduce scatter. An improved method of whole-body GFP imaging made use of a laser excitation source and band-pass filters matched specifically to GFP and constitutive tissue fluorescence emission bands. Processing of the primary GFP fluorescence images acquired by the CCD camera subtracted background tissue autofluorescence. This approach achieved 100% sensitivity and specificity for in vivo detection of a population of BxPc3 pancreatic tumor cells, in which 10% were transfected with GFP, after subcutaneous grafting or orthotopic implantation in the pancreas of nude mice [66]. The luciferase reporter technique requires that animals are anesthetized and restrained so that sufficient photons to construct a pseudo-image can be collected. Furthermore, this process must be carried out in a virtually light-free environment and animals must be injected with the luciferin substrate, which has to reach every tumor cell in order to be useful. The clearance of the luciferin also results in an unstable luciferase signal [67]. These limitations preclude studies that would be perturbed by anesthesia, restraint, or substrate injection and also make high-throughput screening unfeasible. Expression of firefly luciferase (Luc) can be used to visualize tumor growth and regression in response to various therapies in mice. However, detection of Luc-labeled cells in vivo was limited to at least 1,000 human tumor cells [68, 69]. Real images are not produced. Instead, pseudocolor images indicating photon counts are seen. Thus, cellular or subcellular imaging has not been achieved with luciferase as has been done with fluorescent proteins. The high-intensity signal produced by GFP allows unrestrained animals to be imaged without any perturbation or substrate—irradiation with nondamaging blue light is the only step needed. Images can be captured with fairly simple apparatus and there is no need for total darkness. A fusion reporter vector harboring a Renilla luciferase reporter gene, a reporter gene encoding a monomeric RFP, and a mutant herpes simplex virus type thymidine kinase was tested in vivo. A highly sensitive cooled charge-coupled device camera compatible with both luciferase and fluorescence imaging compared these two signals from the fused reporter gene using a lentivirus vector in 293T cells implanted in nude mice. The signal from RFP was approximately 1,000 times stronger than that from luciferase [70]. Since the two proteins were transcribed from the same promoter, these data suggest similar numbers of molecules of each in the cells, further emphasizing the relative brightness of fluorescent proteins. Near-infrared probes activated by the action of proteases [71–73] can also be used for optical imaging of tumors. This approach requires substrate injection and the tumor must contain a specific protease that cleaves the substrate. Tumors on normal tissues such as the liver that also contain these proteases cannot be visualized because background signals are too high.


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1.12 New Features and Models with Fluorescent Proteins A mutant of Anemonia sulcata chromoprotein asCP has been generated. The mutant protein is capable of irreversible photoconversion from the nonfluorescent to a stable bright-red fluorescent form (“kindling”). This “kindling fluorescent protein” (KFP1) can be used for precise in vivo photolabeling to track the movements of cells, organelles, and proteins [74]. A dual-color monomeric protein, photoswitchable cyan fluorescent protein (PSCFP), has been described. PS-CFP is capable of efficient photoconversion from cyan to green, changing both its excitation and emission spectra in response to 405-nm light irradiation. Complete photoactivation of PS-CFP results in a 1,500fold increase in the green-to-cyan fluorescence ratio. PS-CFP was used as a photoswitchable tag to study trafficking of the human dopamine transporter in living cells [75]. Structural data have shown that fragmentation of the chromophore is an intrinsic, autocatalytic step toward the formation of the mature chromophore of the photoactivatable Anemonia sulcata GFP [76]. Recently, a GFP has been cloned from a coral that has the property of sensitive and reversible wavelength-dependent on–off switching. This switchable GFP has been shown to be useful for studying protein trafficking between nucleus and cytoplasm [77]. Future studies could use the switchable GFPs in vivo. As mentioned above, a transgenic GFP nude mouse with ubiquitous GFP expression has been developed. In the adult mice, all the organs brightly express GFP. RFP-expressing human cancer cell lines grew extensively in the transgenic GFP nude mice [51]. This dual-color model shows the details of the tumor–stroma interaction, especially tumor-induced angiogenesis and tumor-infiltrating lymphocytes. Recently, Vintersten et al. [78] reported the development of an RFP-expressing transgenic mouse with expression in many tissues. A variant of DSRed, DSRedT3, was used to develop the ES cells to make this transgenic mouse. The availability of GFP and RFP transgenic mice gives rise to the possibility of GFP–RFP chimeric mice to color code host cells with specific genes of interest that could control specific aspects of tumor growth and progression. Their effects could then be imaged upon transplantation of specific tumor types which would fluoresce a third color. A GFP transgenic rat has recently been developed using the CAG promoter. GFP expression in brain, lung, liver, and islet tissues was restricted to early developmental stages, but it was continuously strong in the exocrine pancreas, kidney, and cardiac and skeletal muscles [79]. An albumin enhancer/promoter-driven Alb-DsRed2 transgenic rat has now been developed that expresses DsRed2 in hepatolyses. To study the transdifferentiation of bone marrow cells into albumin-producing cells, bone marrow cells from the Alb-DsRed2 Tg rat were injected into rats having acute and chronic liver damage. DsRed2-positive cells were generated in the recipient liver after bone marrow injection, thereby visualizing the differentiation of bone marrow cells into hepatocytes [80].

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1.13 Advantages of Fluorescent Protein Imaging Over Quantum Dots Multiphoton microscopy techniques and transgenic mice expressing GFP were combined with quantum dots. These fluorescent semiconductor nanocrystals were customized to specifically image tumor vessels. The recruitment of quantum dotlabeled bone marrow-derived precursor cells to the tumor vasculature was also visualized [81]. Unlike fluorescent protein-expressing cells, the quantum dots could not be passed on to progeny cells, thereby precluding time course tracking of the initiation and development of metastasis or other long-time-dependent studies. Fluorescent proteins are also now available in multiple colors [82] and do not have the toxicity associated with quantum dots [83–86].

1.14 Growth of Fluorescent Protein-Expressing Cells in Syngeneic Models: Lack of Apparent Immunological Response A syngeneic murine colon carcinoma cell line (colon 26 cells) was transfected with enhanced green fluorescent protein (EGFP). The transfected cells maintained the highly malignant attributes of the wild-type cells. Following injection into the portal circulation of Balb/c mice, liver metastases occur in the same time span in both parental and GFP cells, indicating no immune reaction against the GFP-expressing cells [36, 87]. Using a syngeneic mouse model harboring primary, genetically modified, mycdriven lymphomas expressing GFP, whole-body imaging showed that disruption of apoptosis downstream of p53 by Bcl-2 confers a highly aggressive phenotype, metastatic throughout the mouse body [88]. The myc-driven, GFP-expressing lymphomas with p53 or INK4a/ARF mutations or overexpressed Bcl-2 respond poorly to cyclophosphamide therapy in vivo compared to wild-type cells as shown by whole-body imaging [89]. No immune reactions against the GFP cells were observed in this syngeneic system either. B16F0-GFP mouse melanoma cells were injected into the tail vein or the portal vein of 6-week-old C57BLy6. Whole-body optical images showed metastatic lesions in the brain, liver, and bone of B16F0-GFP, again suggesting no immune reaction against the GFP-expressing cells [9]. A syngeneic model of the Lewis lung carcinoma in which the carcinoma cells are labeled GFP was established. The tumor cells were transplanted on the dorsal side of the ear of C57-B16 mice in order to give the tumor cells access to the lymphatic system. This model of the Lewis lung carcinoma extensively metastasized to numerous lymph nodes throughout the body of the animal as well as visceral organs as visualized by fluorescence microscopy using the bright GFP signal. Twenty-one different metastatic sites, including lymph nodes throughout the body, were identified among the cohort of transplanted animals. The data demonstrate a predilection of the Lewis lung carcinoma for lymphatic pathways of metastasis throughout the


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animal body. This syngeneic model again demonstrates no immune reaction against the GFP-expressing tumor cells [90]. These studies strongly indicate that GFP-expressing cells are not more immunogenic than parental cells and therefore GFP is not particularly antigenic.

2 Conclusions Tumor cells stably expressing GFP and other fluorescent proteins in vivo are a powerful new tool for cancer research. Stability of expression has been studied by Naumov et al. [91], who noted that all the CHO-K1-GFP cells used in their study were stably fluorescent (measured by flow cytometry) even after 24 days of growing in medium where they were deprived of selective pressure. This finding implies that GFP can be stably expressed in cells in vivo. This feature has been proved true for all cells studied so far and is exemplified by the generation of extensive GFP-expressing metastases. The use of GFP-expressing tumor cells in fresh tissue or live animals [4, 9, 21, 28, 91] has provided new insights into the real-time growth and metastatic behavior of cancer. Several independent studies, which include an extensive comparison between metastases of GFP-transduced carcinoma and the parental cell lines [36, 92, 93], have shown that GFP or RFP transduction and expression do not affect metastatic behavior including syngeneic models [36]. These studies indicate that GFP expression does not demonstrably affect the basic properties of the cells. GFP can be transfected into any cell type of interest and used as a cytoplasmic marker to show the general outlines of cells in vivo and fine morphological details such as long slender pseudopodial projections [91]. Cells which express GFP in the nucleus and RFP in the cytoplasm offer even more cellular details in vivo [48, 51]. The development of tumor cells that stably express GFP at high levels has enabled investigation of tumor and metastatic growth in a completely noninvasive manner, by using whole-body imaging [9]. For the first time, tumor growth and metastatic studies, including drug evaluations, can be done and quantified in real time in unperturbed individual animals. The potential of this technology is very high. A further advantage of GFP-expressing cells is the increased contrast between brightly fluorescent tumor tissue and blood vessels within it. The ability to visualize and quantify blood vessel development in metastases in vivo will greatly facilitate studies of angiogenesis and the testing of effects of antiangiogenic agents on metastatic development [18, 21, 91]. A transgenic GFP nude mouse with ubiquitous GFP expression has been developed. In the adult mice, all the organs brightly express GFP. RFP-expressing human cancer cell lines grew extensively in the transgenic GFP nude mouse [51]. This dual-color model shows the details of the tumor–stroma interaction, especially tumor-induced angiogenesis and tumor-infiltrating lymphocytes. Red fluorescent protein transgenic rodents have not been developed, opening the possibility of multicolor normal chimeras as well as tumor–host chimeras [80].

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The GFP approach has several important advantages over other optical approaches to imaging. In comparison with the luciferase reporter, GFP has a much stronger signal and therefore can be used to image unrestrained animals. Fluorescent proteins are also available in multiple colors in addition to green, yellow, blue, and red. Shaner et al. [82] have developed mutants with new colors (see below). Multiple colors enable multiple events to be simultaneously imaged. Fluorescent protein imaging not only enables whole-body imaging but also, due to its cellular resolution and multiple colors, gives rise to a new field of in vivo cell biology.

2.1 Example of the Use of Fluorescent Proteins to Discover Properties of Stem Cells Nestin-driven (ND) GFP stem cells isolated from the hair follicle bulge area that are negative for the keratinocyte marker keratin 15 can differentiate into neurons, glia, keratinocytes, smooth muscle cells, and melanocytes in vitro. These pluripotent ND-GFP stem cells are positive for the stem cell marker CD34, as well as keratin 15-negative, suggesting their relatively undifferentiated state. The apparent primitive state of the ND-GFP stem cells is compatible with their pluripotency. Furthermore, cells derived from ND-GFP stem cells can differentiate into neurons after transplantation to the subcutis of nude mice. These results suggest that hair follicle bulge area ND-GFP stem cells may provide an accessible, autologous source of undifferentiated multipotent stem cells for therapeutic application [94].

2.2 Future Directions The applications of in vivo cellular imaging with fluorescent proteins should markedly expand with the development of proteins with new colors. Shaner et al. [82] have taken the Discosoma RFP and converted it through multiple amino acid substitutions into a monomer. With further genetic modification, this group has created from the Discosoma RFP monomer a series of modified proteins with multiple new colors from yellow-orange to red-orange. These new colored proteins include mBanana, mOrange, dTomato, tdTomato, mTangerine, mStrawberry, and mCherry with increasingly longer emission maxima. It is expected that many additional colored proteins will be isolated from various organisms and modified to produce even more colors. The availability of a large number of different colored proteins will enable simultaneous imaging of multiple cellular events in vivo, surpassing what can be visualized both in vivo and in vitro today. These switchable GFPs that can either be kindled and/or be extinguished or change colors have been shown to be useful in studying protein trafficking between nucleus and cytoplasm [77]. Future studies could use the switchable GFP in vivo.


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The availability of GFP and RFP transgenic rodents [78, 80] gives rise to the possibility of GFP–RFP chimeras. Such chimeras could be used to color code host cells that effect tumor growth and progression. Their effects could then be imaged upon transplantation of specific tumor types which would fluoresce a third color. Ilyin and coworkers visualized glioma cells in rats by inserting a fiber-optic endoscope through a preimplanted guide cannula. Tumor monitoring was coupled to confocal microscopy so that visualization of the fluorescent signals from the C-6 glioma-GFP cells that had been preimplanted in the brain was very sensitive [63]. Funovics et al. [95] described the design and construction of a miniaturized multichannel near-infrared endoscopic imaging system developed for high-resolution imaging of mice. This endoscope was used to visualize tumor cells transplanted orthotopically in mice. The developed device should be useful for in vivo imaging using fluorescent proteins [64]. Stanziale et al. [65] showed that the herpes simplex virus NV1066 expressing GFP can be used to assess oncolytic therapy in a minimally invasive, laparoscopic system in mouse models of gastric cancer. After intraperitoneal administration of NV1066-GFP, macroscopic tumor foci begin to express GFP visualized by direct laparoscopy with the appropriate fluorescent filters. Noncancerous organs were not infected and did not express GFP. Thus, GFP expression in intraperitoneal tumors can be visualized laparoscopically, allowing detection and localization of viral gene therapy. A fluorescence system for studying protein dynamics in mitosis has been established in a human cell line expressing histone H3 and truncated importin α as fusions to cyan fluorescent protein (CFP) and RFP, respectively, to visualize the chromosomes and the nuclear envelope in living cells, respectively [96]. This is a further development to apply to the new field of in vivo cell biology that can visualize nuclear and chromatin changes during cancer progression.

2.3 Multiphoton Imaging Brown et al. [25] showed that multiphoton laser scanning microscopy (MPLSM) [97] could provide high–resolution, three-dimensional images of angiogenesis gene expression and that this technique could be used to investigate deeper regions of GFP-expressing tumors in dorsal skin-fold chambers. MPLSM offers significant advantages such as improved signal-to-background ratio, longer sample lifetimes as well as greater imaging depths. Condeelis and coworkers have used this method for intravital imaging of tumor cell behavior in both orthotopic tumor transplant models and transgenic mice that express GFP. This method has opened the primary tumor to imaging at the cellular level of resolution at increased depth [92, 98]. To monitor the activity of the vascular endothelial growth factor (VEGF) promoter, Fukumura et al. [38, 39] made transgenic mice that express GFP under control of the VEGF promoter. Multiphoton laser scanning microscopy showed that the tumor was able to induce the activity of the VEGF promoter and subsequent vessel formation.

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2.4 Potential Human Use of Fluorescent Proteins One obvious use of fluorescent proteins in humans would be to enable the tracking of the primary tumor prior to surgery or chemotherapy treatment and then to monitor for fluorescence after treatment to identify possible recurrence or metastasis. While currently it is possible to express fluorescent proteins in mouse models of human tumors using retroviral or liposomal delivery of the gene, many of these gene expression techniques have yet to be refined where their use in other than preclinical trials is ethically approved. However, in principle, the idea of being able to essentially label a primary tumor in a patient and then follow that tumor throughout the treatment regimen would undoubtedly revolutionize cancer treatment. In a proof-of-principle experiment, Hasegawa et al. [99] have shown that injection of the GFP gene cloned into a retroviral vector was expressed in intraperitoneally growing human stomach cancer in a nude mouse model. Eventually, widespread tumor growth was visualized by GFP expression, including liver metastasis. No normal tissues were found to be transduced by the GFP-expressing retrovirus. Thus, reporter gene transduction of the primary tumor enabled detection of its subsequent metastasis. Future human studies could thus use intra-operative transfection with highly efficient vectors coding for fluorescent proteins that would enable visualization of tumor recurrence after surgery with the use of fluorescence endoscopy and other optical methods. Genetic labeling of the tumor cells with GFP will be inherited by the progeny of the tumor cells. This would be useful for detecting the proliferation of tumor cells that would not be killed by standard therapy or gene therapy associated with the GFP labeling. To ensure tumor-specific expression, GFP linked to a tumor-specific promoter could be introduced systemically and only the tumor cells would turn on the expression of the GFP plasmid. To determine whether the carcinoembryonic antigen (CEA) promoter could control the expression of the GFP gene, human MKN45-GFP stomach cancer cells were injected into the peritoneal cavity of BALB/c nude mice [100]. A CEA-EGFP plasmid was then introduced into the peritoneal cavity using liposomes. GFP-fluorescent tumor nodules were subsequently detected by fluorescence stereomicroscopy. In another approach, GFP was conjugated to the transferrin promoter in order to target disseminated tumors in vivo [101]. When GFP gene conjugates were systemically administered through the tail vein to nude mice that had been subcutaneously inoculated with tumor, GFP expression was only detected in the tumor. Varda-Bloom and colleagues [102] developed a tissue-specific gene therapy directed at angiogenic blood vessels of tumor metastasis using an adenovirus-based vector containing the murine preproendothelin-1 (PPE-1) promoter driving GFP. High expression and specific activity of PPE-1 were achieved by systemic administration of the adenoviral vector to mice bearing Lewis lung carcinoma tumors. GFP expression was detected in the new vasculature of primary tumors and lung metastasis. The highest area of expression was in the angiogenic endothelial cells of the metastasis.


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Cancer surgery requires the complete and precise identification of malignant tissue margins including the smallest disseminated lesions. Internal GFP fluorescence can intensely illuminate even single cells, but requires GFP sequence transcription within the cell. Introducing and selectively activating the GFP gene in malignant tissue in vivo is made possible by the development of OBP-401, a telomerase-dependent, replication competent adenovirus expressing GFP. This potentially powerful adjunct to surgical navigation was demonstrated in 2 nude mouse models that represent difficult surgical challenges—the resection of widely disseminated cancer. HCT-116, a model of intraperitoneal disseminated human colon cancer, was labeled by virus injection into the peritoneal cavity. A549, a model of pleural dissemination of human lung cancer, was labeled by virus administered into the pleural cavity. Only the malignant tissue fluoresced brightly in both models. In the intraperitoneal model of disseminated cancer, fluorescence-guided surgery enabled resection of all tumor nodules labeled with GFP by OBP-401. This suggests that adenoviral-GFP labeling tumors in patients can enable fluorescence-guided surgical navigation [103, 104]. The above approach to labeling tumors in a mouse with a tumor-specific adenovirus can be applied to human beings after additional safety issues have been addressed. Such adenoviruses could also use more red-shifted proteins which would have advantages for imaging deeper into tissues [105, 106]. It could even be possible to deliver genes to tumors by the above method that express color coded proteins identifying the position of cancer cells in the cell cycle [107]. This would be highly useful for following the effects of chemotherapy in the patient’s tumor. It could even be possible to deliver genes coding for multiple colors of fluorescent proteins with appropriate recombinant sites between them allowing for a multitude of lineages within the tumor to be individually labeled with different colors [108] which could facilitate the identification of cancer stem cells within the patient. Because of its great utility, GFP was chosen for the Nobel Prize in Chemistry in 2008 [109–111]. It is expected that this great potential in fluorescent proteins will find great utility to directly treat human disease. Endoscopy and laparoscopy offer real possibilities of clinical application of fluorescent protein imaging. Detection of single cancer cells will become possible in patients. Futuristic applications could include implantation of microlight sources and detectors which could signal external detectors of the presence of a single new fluorescent cancer cell growing in the patient.

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Genetically Encoded Fluorescent and Bioluminescent Probes for Illuminating Cellular Signaling Pathways Yoshio Umezawa

The intracellular signaling can be monitored in vivo in living cells by genetically encoded intracellular fluorescent and bioluminescent probes or indicators, which include second messengers, protein phosphorylation, protein–protein interactions, and protein localizations in organelles. These probes are of general use not only for fundamental biological studies but also for assay and screening of possible pharmaceutical or toxic chemicals that inhibit or facilitate cellular signaling pathways.

1 Introduction Intercellular signaling substances (first messengers) such as neurotransmitters, cytokines, and hormones functioning, respectively, in nerve, immune, and endocrine systems bind either to ion-channel, kinase, or G protein-coupled membrane receptor proteins and trigger the respective downstream intracellular signaling. The intracellular signaling can be monitored in vivo in living cells [1] by genetically encoded intracellular fluorescent and bioluminescent probes or indicators. The probes include second messengers such as nitric acid (NO), inositol 1,4,5-trisphosphate (IP3 ), cyclic guanosine 3 ,5 -monophosphate (cCMP), and phsphatidylinositol-3, 4,5-trisphosphate [2–7], protein phosphorylation [8–10], protein–protein interactions [11–14], and protein localizations in organelles [15–19]. These probes are of general use not only for fundamental biological studies but also for assay and screening of possible pharmaceutical or toxic chemicals that inhibit or facilitate cellular signaling pathways.

Y. Umezawa (B) Department of Chemistry, School of Science, The University of Tokyo Hongo, Bunkyo-ku, Tokyo 113-0033, Japan e-mail: [email protected]


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2 Second Messengers 2.1 Nitric Oxide (NO) Nitric oxide (NO) is a small uncharged free radical that is involved in diverse physiological and pathophysiological mechanisms. NO is generated by three isoforms of NO synthase (NOS), endothelial, neuronal, and inducible ones. When generated in vascular endothelial cells, NO plays a key role in vascular tone regulation, in particular. Here we describe an amplifier-coupled fluorescent indicator for NO to visualize physiological nanomolar dynamics of NO in living cells (detection limit of 0.1 nM) (Fig. 1). This genetically encoded high-sensitive indicator revealed that approximately 1 nM of NO, which is enough to relax blood vessels, is generated in vascular endothelial cells even in the absence of shear stress. Nanomolar range of basal endothelial NO thus revealed appears to be fundamental to vascular homeostasis [2].

2.2 Inositol 1,4,5-Trisphosphate (IP3 ) Inositol 1,4,5-trisphosphate (IP3 ) is a key second messenger in many cell types and also in distinct subcellular regions of single living cells; however, little is examined about the subcellular dynamics of IP3 in a variety of cell types. We have developed fluorescent indicators to locate IP3 dynamics in single living cells based on an intramolecular fluorescence resonance energy transfer (FRET) (Fig. 2). Our indicator has visualized IP3 dynamics in the cytoplasm of cultured cells and even in single thin dendrites of hippocampal neurons, which has been unseen previously. We have further localized the present indicator in the nucleus and pinpointed nuclear IP3 dynamics. The observation with our nuclear IP3 indicator has solved a question on nuclear propagation of IP3 from the cytoplasm and has drawn a conclusion that the nuclear IP3 dynamics synchronously occurs with cytosolic IP3 dynamics evoked by agonist stimulations. The present approach contributes to understand when, where, and how IP3 is generated and removed in a variety of living cells [3].

2.3 Cyclic GMP cGMP acts as a signaling molecule that regulates various physiological processes such as the relaxation of muscle cells and neuronal activity. Imaging the dynamics of cGMP in single living cells is of importance to reveal the molecular mechanisms underlying cellular and organ functions. We developed a fluorescent indicator for cGMP, named CGY, and succeeded in achieving concentration changes in single living cells (Fig. 3). CGY is composed of PKG I, which specifically recognizes cGMP, and CFP and YFP connected to its flanking ends. FRET between the two fluorescent proteins was found to increase upon cGMP-induced conformational change

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Fig. 1 An amplifier-coupled fluorescent indicator for visualizing NO in single living cells. (a) Schematic representations of domain structures of sGCα, sGCβ, CGY, sGCα-CGY, and sGCβCGY. The amino acid sequence of FLAG tag and linker (Ln) is shown at the bottom. The heterodimer of sGCα-CGY and sGCβ-CGY was named “NOA-1.” (b) Principle of the present NO indicator, NOA-1. sGCα-CGY and sGCβ-CGY are spontaneously associated to form a matured heterodimer, that is, NOA-1. NOA-1 binds with NO and generates cGMP at the rate of 3000– 6000 molecules/min. Thus generated cGMP binds to the CGY domain in NOA-1 make NOA-1 emit a FRET signal. Approximately 99.9% of cGMP molecules thus generated diffuse and bind to NO-free NOA-1. As a result, even a single NO molecule can trigger a large number of NOA-1 to emit FRET signals. Even if sGCα-CGY and sGCβ-CGY exist as monomers, the monomers also emit FRET signals upon binding with generated cGMP

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Fig. 2 Fluorescent indicators for IP3 in single living cells. (a) Principle of fretino for visualizing IP3 . CFP and YFP are different colored mutants of green fluorescent protein from Aequorea victoria with mammalian codons and additional mutation. Upon binding of IP3 with the hIP3 R1224–579 within fretino, a conformational change of fretino takes place, which changes the efficiency of FRET from CFP to YFP. (b) Schematic representations of domain structures of the present fretinos. Shown at the top of each bar are the restriction sites. hIP3 R1224–579 and hIP3 R11–604 are derived from a IP3 -binding domain of human IP3 R1 and selectively binds with IP3 . R504Q shows a mutation, in which the arginine 504 is replaced with a glutamine to partly reduce the affinity of the mutant hIP3 R1224–579 with IP3 . K508A represents a mutation, in which the lysine 508 is replaced with an alanine to inhibit the binding of IP3 with the mutant hIP3 R1224–579 . NLS stands for a nuclear localization sequence, the amino acid sequence of which is shown at the bottom


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Fig. 3 cGMP indicator based on fluorescence resonance energy transfer (FRET). (a) Schematic diagram showing how to detect cGMP on the basis of FRET between CFP and YFP. The cylindrical β-can structure and the central chromophore of GFPs are drawn according to the crystal structure of Aequorea victoria GFP. The precise overall structure of PKG Iα1–47 and relative orientations of the GFPs are not known. However, small-angle X-ray scattering data have revealed a marked conformational change in PKG Iα upon cGMP binding. (b) Pseudocolor images of CFP/YFP emission ratios expressed as pseudocolor images before (time 0) and at several time points after the addition of 1 mM 8-Br cGMP, obtained from the CH)-KI cells expressing the indicator [6]

in the receptor. We demonstrated that the thus developed fluorescent indicators (CGYs) reversibly responded to cGMP produced in nitric oxide (NO)-stimulated cells. Using CGYs, we found novel cGMP dynamics including NO-induced cGMP oscillation in single living cells. The present genetically encoded fluorescent indicators open a way not only for understanding the complex dynamics of cGMP signaling in single cells and organisms but also for discovering pharmaceuticals such as isoform-specific inhibitors for phosphodiesterases [6].

2.4 Phosphatidylinositol-3,4,5-trisphosphate (PIP3 ) Phosphatidylinositol-3,4,5-trisphosphate (PIP3 ) regulates diverse cellular functions, including cell proliferation and apoptosis, and has roles in the progression of

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diabetes and cancer. However, little is known about its production. We have developed fluorescent indicators for PIP3 based on fluorescence resonance energy transfer (FRET) [7]. These novel PIP3 indicators are composed of two distinctly colored mutants of GFP and a PIP3 -binding domain. The PIP3 level was observed by dualemission ratio imaging, thereby allowing stable observation without the problem of artifacts described above. Furthermore, these indicators were fused with localization sequences to direct them to the plasma membrane or endomembranes, allowing

Fig. 4 Fluorescent indicators for PIP3 in single living cell. (a) Principle of fllip for visualizing PIP3 . CFP and YFP are different colored mutants of green fluorescence protein from Aequorea victoria with mammalian codons and additional mutation. Upon binding of PIP3 with the PHD within fllip, a filp-flop-type conformational change of fllip takes place, which changes the efficiency of FRET from CFP to YFP. (b) Schematic representations of domain structures of the present fllips. Shown at the top of each bar are the restriction sites. PHD is derived from human GRP1 (261– 382) and selectively binds with PIP3 . PHD-R284C is a mutant PHD, in which the arginine 284 is replaced with a cysteine not to bind with PIP3 . Ln1, Ln2 and Ln3: linkers, the amino acid sequences of which are shown in the bottom. MLS1 and MLS2: membrane localization sequences to the plasma membrane and the endomembranes, respectively, the amino acid sequences of which are also shown at the bottom [7]


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localized analysis of PIP3 concentrations. Using these fluorescent indicators, we analyzed the spatio-temporal regulation of the PIP3 production in single living cells. To examine PIP3 dynamics, we used a pleckstrin homology (PH) domain from GRP1 [20], which selectively binds PIP3 , fused between cyan (CFP) and yellow (YFP) fluorescent protein variants, through rigid α-helical linkers 12 consisting of repeated EAAAR sequences. Within one of the rigid linkers, a single di-glycine motif was introduced as a hinge. We then tethered the chimeric indicator protein to the membrane by fusing it with a membrane localization sequence (MLS) through the rigid α-helical linker. Thus, after PI(3)K activation, the PH domain binds to PIP3 and a significant conformational change of the indicator protein occurs through the flexible di-glycine motif introduced into the rigid α-helical linker. This “flip-floptype” conformational change of the indicator protein causes intramolecular FRET from CFP to YFP, allowing detection of PIP3 dynamics at the membrane (Fig. 4). We named this indicator “fllip” (fluorescent indicator for a lipid second messenger that can be tailor-made). The developed flip allows a spatio-temporal examination of PIP3 production in single living cells. After ligand stimulation, PIP3 levels increased to a larger extent at the endomembranes (that is, the endoplasmic reticulum and the Golgi) than at the plasma membrane (Fig. 5). This increase was found to originate from in situ production at the endomembranes, a process stimulated directly by receptor tyrosine kinases endocytosed from the plasma membrane to the endomembranes. The demonstration of PIP3 production through receptor endocytosis addresses a long-standing question about how signaling pathways downstream of PIP3 are activated at intracellular compartments remote from the plasma membrane.

Fig. 5 Response of fllip-em to PIP3 at the endomembranes after PDGF stimulation. (a) Pseudocolor images of the CFP:YFP emission ratio before (0 s) and 120, 300, or 600 s after addition of 50 ng ml−1 PDGF. Experiments were performed at 25◦ C on CHO-PDGFR cells expressing fllip-em. (b) Time course of fllip-em and fllip-pm emission ratio after stimulation with PDGF [7]

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3 Fluorescent Indicators for Imaging Protein Phosphorylation Protein phosphorylation by intracellular kinases plays one of the most pivotal roles in signaling pathways within cells. The kinase proteins catalyze transfer of the phosphate of ATP and phosphorylation of the hydroxy groups of serines, threonines, and/or tyrosines on the substrate proteins. Upon this phosphorylation, the substrate proteins are subject to conformational changes caused by the negative charges of the phosphates, which subsequently trigger their enzymatic activation and interaction with their respective target proteins. To reveal the biological issues related to the kinase proteins, electrophoresis, immunocytochemistry, and in vitro kinase assay have been used. However, these conventional methods do not provide sufficient information about the spatial and temporal dynamics of signal transduction based on protein phosphorylation and dephosphorylation in living cells. To overcome the limitations of investigating kinase signaling, we developed genetically encoded fluorescent indicators for visualizing the protein phosphorylation in living cells [8]. The principle of the present method is schematically shown in Fig. 6. A substrate domain for a kinase protein of interest is fused with a phosphorylation recognition domain via a flexible linker sequence. The tandem fusion unit consisting of the substrate domain, linker sequence, and phosphorylation recognition domain is sandwiched with two fluorescent proteins of different colors, cyan fluorescent protein (CFP) and yellow fluorescent protein (YFP), which serve as the donor and acceptor fluorophores for fluorescence resonance energy transfer (FRET). As a result of the phosphorylation of the substrate domain and subsequent binding of the phosphorylated substrate domain with the adjacent phosphorylation recognition domain, FRET is induced between the two fluorescent units, which causes phosphorylation-dependent changes in the fluorescence emission ratios of the donor and acceptor fluorophores. Upon activation of the phosphatases, the phosphorylated substrate domain is dephosphorylated and the FRET signal is decreased. We named this indicator “phocus” (a fluorescent indicator for protein phosphorylation that can be custom made). At the phosphorylation recognition domain within phocus, not only endogenous domains such as src homology 2 (SH2) domains, phosphotyrosine binding (PTB) domains, and WW domains but also single chain antibodies (scFvs) immunized with the phosphorylated substrate sequences of interest are available. Thus, the present method has more general applicability for kinase signaling in living cells compared with previously reported fluorescent indicators based on uncontrollable conformational changes in the substrate peptides themselves upon phosphorylation. By using suitable substrate and phosphorylation recognition domains, we have developed a large number of phocuses for several key protein kinases including a receptor tyrosine kinase, insulin receptor(1), a serine/threonine protein kinase, Akt/PKB(2), and a non-receptor tyrosine kinase, c-Src(3) (Table 1). In addition, these phocuses have been further tailored to visualize local activity of the respective protein kinases in living cells by fusing appropriate localization sequences/domains with each phocus (Table 1) [7, 8].


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Fig. 6 Fluorescent indicators for protein phosphorylation in living cells. (a) Principle of phocus for visualizing protein phosphorylation. CFP and YFP are different colored mutants of green fluorescent protein (GFP) derived from Aequorea victoria with mammalian codons and additional mutations. (b) The CFP/YFP emission ratio is pseudocolor images of the CFP/YFP emission ratios before (time 0) and at 40, 80, 300, and 600s after the addition of 100 nM insulin, obtained from the CHO-IR cells expressing phocus in which a nuclear export signal peptide attached next to YFP [8]

4 Protein–Protein Interactions 4.1 Split-GFP is Spliced upon Protein–Protein Interactions To monitor protein–protein interactions (PPIs), a new method with general applicability was developed based on protein splicing (Fig. 7). In this process, an intervening protein sequence is excised and the flanking protein fragments are

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Table 1 Detailed information on composition of each fluorescent indicator for protein phosphorylation and on each location to be directed in single living cells

Kinase

Indicator

Localization domain

Insulin receptor phocus-2 pp (tyrosine kinase)

PH-PTB domain from IRS-1

Akt/PKB (serine/ theronine kinase) c-Src (tyrosine kinase)

Aktus eNOS-Aktus Bad-Aktus Srcus

– eNOS1–35 Tom201–33 –

PTB-Srcus

Shc56–206

Localization Cytosol, nucleus and membrane ruffles including insulin receptor Cytosol Golgi Mitochondria Cytosol Cytosol and at the EGFR

Substrate domain

Phosphorylation recognition doman

ETGTEEYM-KMDLG

p85α330–429

RGRSRSAP RGRSRSAP RGRSRSAP EEEIYGEFF

14-3-3η82–235 14-3-3η82–235 14-3-3η82–235 c-Src148–248

EEEIYGEFF

c-Src148–248

Fig. 7 (a) Principle of the split-GFP system. (b) Fluorescent spectra in the presence or absence of interacting proteins. (c) Fluorescent images of Escherichia coli carrying interacting proteins (brighter spots) [12]


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spliced together. In this splicing system, the flanking pieces are the N- and C-terminal fragments of green fluorescent protein (GFP). The intervening sequence is dnaE derived from Synechocystis. An interaction between protein A and B brings the two parts of dnaE close enough to fold together properly and initiate the splicing and linking of the two EGFP halves with a peptide bond. Reconstitution of EGFP is monitored by its fluorescence. For a proof of the principle, calmodulin and its target peptide, known as M13, were chosen. The fluorescence increased upon expression of the two proteins and its intensity was proportional to the number of interacting protein pairs. The protein interactions were thereby observed in bacterial colonies on LB agar plates. This system is also applicable for monitoring protein interactions in mammalian cells. Unlike an earlier protein interaction assay, the split-GFP system involves the reconstitution of GFP and does not require that the PPIs occur near the cell nucleus and reporter genes or that an enzyme substrate be present. This will make the method generally more useful and enable screening of the interactions in the cytosol or at the inner membrane level [11–13].

4.2 Locating a Protein–Protein Interaction by Split Renilla luciferase Complementation For spatial and quantitative kinetic analysis of protein–protein interactions (PPIs) in living mammalian cells, a method was developed, in which PPI-induced complementation of split Renilla luciferase triggers spontaneous emission of luminescence, with a cell membrane permeable substrate, coelenterazine [14]. This split Renilla luciferase complementation readout was shown to work for locating a PPI between tyrosine-phosphorylated peptide (Y941) of IRS-1 and SH2 domain of PI3K among insulin signaling pathways in living Chinese hamster ovary cells overexpressing human insulin receptors (CHO-HIR). It was thereby found that the insulin-stimulated interaction occurred near to the plasma membrane in the cytosol. The principle of the present split Renilla luciferase complementation strategy is shown in Fig. 8. To monitor the interaction between two proteins A and B, N-terminal half of the split Renilla luciferase is fused to protein A and protein B to C-terminal half of the split Renilla luciferase. Interaction between protein A and protein B and the consequent juxtapositioning of the split Renilla luciferase simultaneously leads to formation of the complement Renilla luciferase, thereby spontaneously emitting bioluminescence with its cell membrane permeable substrate, coelenterazine in situ in living mammalian cells. Spontaneous emission of luminescence by a particular PPI-induced complementation of the split Renilla luciferase together with its membrane permeable substrate, coelenterazine, allows noninvasive imaging of the sites and time of its occurrence in living cells. The PPI between Y941 and SH2n in the CHO-HIR cells expressing sRL91 was thereby imaged with and without insulin stimulation as shown in Fig. 9. Upon 100 nM insulin stimulation, luminescence emitted by

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Fig. 8 A schematic diagram of the complementation strategy based on split Renilla luciferase. The split Renilla luciferase complementation method for locating PPIs relies on the spontaneous emission of luminescence upon PPI-induced complementation of the split Renilla luciferase, with a cell membrane permeable substrate, coelenterazine, in situ in living mammalian cells [14]

Fig. 9 Spatial analysis of the interaction between Y941 and SH2n in living cells. These luminescence microscopic images of CHO-HIR cells were taken with a CCD camera for 300 s (a) and 60 s (b) as exposure times, respectively, in PBS supplemented with a 20% coelenterazine substrate buffer. These images were acquired with the cells expressing sRL91 ((a), left and center), in the absence (left) or presence (center) of 100 nM insulin, and with the cells expressing only full-length Renilla luciferase (hRL124C/A) (b). A phase-contrast transmission image of the cells expressing sRL91 was shown in Fig. 5 ((a), right), which was obtained in PBS for 20 ms as an exposure time. The luminescence intensity is represented with a color scale [14]

complement Renilla luciferase increased only near to the plasma membrane, whereas such bright contrast was not observed in the absence of insulin. This indicates that with insulin stimulation, the interaction between Y941 and SH2n occurred near to the plasma membrane in the cytosol. The CHO-HIR cells expressing fulllength Renilla luciferase (hRL124C/A) emitted luminescence uniformly throughout the cells with its substrate, coelenterazine (Fig. 9b), which precludes the possibility that the result in Fig. 9a is due to an excessive accumulation of coelenterazine in and right below the plasma membrane [14].


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5 Protein Localization in Organelles 5.1 Mitochondria-Targeting Protein One of the most distinct features of eukaryotic cells, in particular mammalian cells, is different compartmentalization of each protein. The protein localization is tightly bound to its function such that preferential localization of a protein is often an essential step toward determining its function. Therefore, functional assays aimed at characterizing the cellular localization of proteins provide a backbone of functional annotations from which new biological significance can be formulated. A technique for the identification of the proteins localized in the organelles largely relies on the preparative isolation of respective compartments through cell fractionation and two-dimensional electrophoresis, combined with mass spectrometry. This is a useful technique for systematic identification of proteins. However, it totally depends on the yield and purity of the intracellular organelle, and therefore, the technique cannot be used for the organelle that is hardly isolated. The control of intricate networks within eukaryotic cells relies on differential compartmentalization of proteins. A method was developed that allows rapid identification of novel proteins compartmentalized in mitochondria by screening large-scale cDNA libraries. The principle is based on reconstitution of splitenhanced green fluorescent protein (EGFP) by protein splicing of DnaE derived from Synechocystis. The cDNA libraries are expressed in mammalian cells following infection with retrovirus. If a test protein contains a functional mitochondrialtargeting signal (MTS), it translocates into the mitochondrial matrix, where EGFP formation occurs by protein splicing (Fig. 10). The cells harboring this reconstituted EGFP are screened rapidly by fluorescence-activated cell sorting (Fig. 11).

Fig. 10 Scheme showing how EGFP is formed by protein splicing of DnaEs when a test protein is localized in the mitochondrial matrix. Three-dimensional structures colored blue and red represent DnaEn and DnaEc, while light and dark gray represent amino- and carboxyl-terminal halves of EGFP, respectively. Orange strand means MTS. OM and IM are mitochondrial outer and inner membranes, respectively [15]

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Fig. 11 Flow cytometry profiles and fluorescent images of representative cloned cells. Left panels: Fluorescence intensity of cloned cells (blue curve) and uninfected BNL1MEmito cells (red curve) were analyzed by flow cytometry. The total cell counts analyzed were 105 cells. Right panels: Each cloned cell was cultured on the glass slide and confocal image of the live cells harboring reconstituted EGFP was taken. The cells were stained with TMRE to show the mitochondrial localization of individual cells. Stacked confocal images show that reconstitution of EGFP occurred in the mitochondria. Bar, 10 μm. [15]

From the screened cells, the cDNAs are isolated and identified. The analysis of 258 cDNAs revealed various MTSs. New transcripts corresponding to mitochondrial proteins were identified. This method provides a broadly applicable means for mapping proteins distributed within intracellular organelle in different tissues or disease states [15].

5.2 ER-Targeting Protein To address difficulties encountered with this GFP- or epitope-tagged approach, we have developed a method with general applicability for high-throughput identification of the genes from large-scale cDNA libraries that encode proteins with the ER signal sequences [16]. The principle is based on reconstitution of split-EGFP fragments by protein splicing (Fig. 12). The protein splicing is a post-translational event involving precise excision of an internal sequence termed intein, and ligation of the flanking sequences, termed N- and C-exteins, by a peptide bond. While most of inteins are composed of single polypeptides, a pair of functional and naturally split intein-coding sequence has been found from the split dnaE genes in the genome


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Fig. 12 Scheme of basic principle and strategy for identifying ER-targeting proteins. (A) Principle for detecting translocation of a test protein into ER using protein splicing of split-EGFP. EGFPc is connected with DnaEc and ER-targeting signal (ERTS), which is predominantly localized in the ER. A test protein is connected with the EGFPn and DnaEn, which is expressed in the cytosol. When the test protein translocates into the ER, the DnaEn interacts with DnaEc and protein splicing results. The EGFPn and EGFPc are linked together by a peptide bond, and the reconstituted EGFP recovers its fluorescence. (B) Domain structures of DnaE-tagged EGFP before and after protein splicing showing sequences of the boundaries between DnaEn and EGFPn, between DnaEc and EGFPc, and between EGFPn and EGFPc

of Synechocystis sp. PCC6803. With this split fragments of the DnaE intein, we have previously developed a new split-EGFP reporter for identifying mitochondrial proteins. The fluorescence of the split-EGFP reporter can be recovered by protein

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splicing when the splicing protein of DnaE is assembled by protein transports into mitochondria. This basic concept was extended for designing a new indicator for identifying proteins transported into the ER. A tandem fusion protein containing an ER-targeting signal (ERTS) and the C-terminal fragments of DnaE and EGFP localizes in the lumens of the ER (Fig. 12a). cDNA libraries are genetically fused to the sequences encoding the N-terminal fragments of DnaE and EGFP. If test proteins expressed from the cDNAs contain an ERTS, the fusion products translocate into the ER, in which the N- and C-terminal halves of DnaEs are brought close enough to fold correctly, thereby initiating protein splicing of the split EGFP to recover its EGFP fluorescence (Fig. 12b). The fluorescent cells are collected by FACS, and from each clone, cDNA is retrieved and its sequence is analyzed. Using this method, we were able to identify 109 non-redundant genes that encode proteins transported to the ER.

5.3 Nucleocytoplasmic Trafficking of Functional Proteins Nucleocytoplasmic trafficking of functional proteins plays a key role in regulating gene expressions in response to extracellular signals. We developed a genetically encoded bioluminescent indicator for monitoring the nuclear trafficking of target proteins in vitro and in vivo [17]. The principle is based on reconstitution of split fragments of Renilla reniformis (Rluc) by protein splicing with a DnaE intein. A target cytosolic protein fused to the amino-terminal half of Rluc is expressed in mammalian cells. If the protein translocates into the nucleus, the Rluc moiety meets

Fig. 13 When AR is bound to 5α-dihydrotestosterone (DHT), it translocates into the nucleus and brings the N- and C-terminal halves of DnaEs close enough to fold correctly, thereby initiating protein splicing to link the concomitant Rluc halves with a peptide bond. The C-terminal half of split Rluc was located beforehand in the nucleus by a fused nuclear localization signal. The cells containing this reconstituted Rluc allow one to monitor nuclear translocation of AR with its luminescence by coelenterazine as the substrate [17]


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the carboxy-terminal half of Rluc, which is localized in the nucleus with a fused nuclear localization signal (NLS), and full-length Rluc is reconstituted by protein splicing (Fig. 13). The bioluminescence is thereby emitted with coelenterazine as the substrate. The principle of the approach is an extension of the earlier developed method to identifying mitochondrial proteins [15]. The method of cell-based screening with the genetically encoded indicator provided a quantitative measure of the extent of nuclear translocation of AR upon stimulating with various chemicals. Currently, high-throughput screening tools for protein translocation into the nucleus have mostly depended on GFP-tagged approach or its variant-tagged approach in combination with the fluorescence microscopy and computer-driven imaging system. The systems offers only semiquantitative information, because it is difficult in each cell to accurately sort out and distinguish the fluorescence of GFP-tagged proteins localized only in the nucleus from that left in the cytosol. In addition, precision of the observed fluorescence intensities from the nucleus obtained with the statistical analysis is not high, because the number of the cells examined under a fluorescence microscope is limited. On the contrary, the present method allowed one to determine the subcellular localization of AR by the luminescence signals generated only when the AR is localized in the nucleus. AR remaining in the cytosol did not induce reconstitution of split Rluc and therefore no background luminescence was observed. PCB and procymidone have been suspected of having neurotoxic and antiandrogenic effects, respectively, and possibly adversely influence the hormonal activities of living animal’s brains. We showed the usefulness of the split Rluc reporter for monitoring AR translocation into the nucleus in living mice by implanting the aforementioned COS-7 cells in the depth of 3 mm in the mouse brain and

Fig. 14 Effects of inhibitors on AR translocation into the nucleus in the mouse brain. PCB and procymidone were found to have an ability to pass through the blood–brain barrier, to reach the brain and to inhibit the AR signal transduction in the organ [17]

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measured emitted bioluminescence there with a cooled CCD camera; we thereby investigated distribution of the chemicals in the brain of living mice. As expected, 2 h after intraperitoneal injection of PCB or procymidone, both chemicals were found to completely inhibit the DHT-stimulated translocation of AR, where coelenterazine was injected intracerebrally (Fig. 14). From the results, it was concluded that PCB and procymidone have an ability to pass through the blood–brain barrier in 2 h, to reach the brain, and inhibit the AR signal transduction in the organs [17]. Similar genetically encoded bioluminescent probes were developed for illuminating protein nuclear transport induced by phosphorylation or by proteolysis [18]. A genetically encoded stress indicator was also reported for noninvasively imaging endogenous corticosterone in living mice [19]. Acknowledgments This work was supported by Japan Science and Technology Agency (JST) and Japan Society for the Promotion of Science (JSPS).

References 1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11. 12. 13. 14. 15. 16. 17. 18. 19. 20.

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Fluorescent Protein FRET Applications Protein Engineering, Intracellular Sensing, and Interaction Screening Annalee W. Nguyen, Xia You, Abeer M. Jabaiah, and Patrick S. Daugherty

1 Introduction Fluorescent proteins provide the opportunity to glimpse inside a living cell and obtain useful biochemical information pertaining to a variety of biological systems. These probes are unique when compared with synthetic fluorophores since they are genetically encoded and can be expressed and retained within the living cell. And, importantly, fluorescent protein fusions have enabled investigation of intracellular transport and localization of various proteins [1–3]. While fluorescent proteins are already ubiquitous in biological research, they are set to become key tools for intracellular sensing, reversible protein interaction screening, and protein engineering.

1.1 FRET Theory Förster resonance energy transfer (FRET) is a distance-dependent phenomenon between two compatible fluorophores, termed the donor and the acceptor. The spectral response to changes in distance between the donor and the acceptor has been dubbed as “spectroscopic ruler,” [4] allowing molecular distances to be measured simply by obtaining a fluorescence spectrum. Under non-favorable conditions for FRET (FRET-off state), excitation of the donor fluorescent molecule results in normal donor emission. If, however, FRET occurs (FRET-on state), the excitation energy of the donor is transferred through dipole–dipole coupling to the acceptor, resulting in emission typical of the acceptor molecule. The difference in fluorescence spectrum between the FRET-on and FRET-off states is defined by the intrinsic properties of the donor and acceptor and the detection sensitivity.

P.S. Daugherty (B) Department of Chemical Engineering, University of California, Santa Barbara, CA 93106-5080, USA e-mail: [email protected]


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Theodor Förster developed a theory describing resonance energy transfer in 1948 [5], relating the photophysical properties of the donor and acceptor to the FRET efficiency and defining the conditions required for FRET. Förster theory predicts that the efficiency of energy transfer is given by E=

R60 R60 + r6

The variable r is the distance between the donor and the acceptor fluorophore and R0 is the “Förster distance.” Factors such as overlap between donor emission and acceptor excitation spectra, donor quantum yield, acceptor extinction coefficient, relative orientation of the dipole moments of the donor and acceptor, and the refractive index of the medium between the fluorophores are combined into the Förster distance [6]. The physical meaning of R0 is the distance at which FRET is 50% efficient. In addition to efficiency, detection of FRET is strongly influenced by the extent of donor emission bleed-through into the acceptor channel, direct excitation of the acceptor at donor excitation wavelengths, and overall brightness of the FRET signal. These additional factors are not considered by Förster’s equation.

1.2 Practical Implications of FRET Theory Förster’s theory indicates that for fluorescent protein FRET pairs, where the fluorophores are separated by roughly 3 nm due to the surrounding protein cage, FRET is at a maximum at around 3 nm and drops off to a minimum at about 10 nm [7]. However, since Förster’s theory was developed for point dipoles in a homogeneous medium, it may provide a less accurate description of the large fluorophore held within the protein cage of the fluorescent protein. The value of R0 for different fluorescent protein pairs has been calculated by multiple groups and ranges from 4.5 to 5.5 nm [7–9]. Though R0 values can be calculated for fluorescent protein FRET pairs with generally accepted assumptions about relative orientation and refractive index, they rarely correlate with the usefulness of a given pair and do not capture the performance of the pair in a particular application.

1.3 Common Applications of FRET The most common uses of fluorescent protein FRET pairs are in the detection of protease activity (Fig. 1a) and protein–protein interactions (Fig. 1b) and for small molecule detection. In the later application, the fluorescent protein donor and acceptor are fused to the termini of a protein that exhibits ligand-dependent conformation changes (Fig. 1c). For example, fusion of CFP and YFP to the termini of bacterial periplasmic binding proteins has generated maltose [10] and glucose [11] indicators.

Fluorescent Protein FRET Applications

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Fig. 1 Common uses of fluorescent protein FRET in molecular biology including (a) protease probes, (b) protein–protein interaction probes, and (c) small molecule indicators

Periplasmic maltose and glucose binding proteins undergo a reversible conformational change that shifts the relative distance between the N- and C-termini upon ligand binding. This conformational change can be observed as a FRET signal change. This approach is potentially generalizable since periplasmic binding proteins exhibit a wide variety of binding specificities [12]. Similarly, the calcium binding protein calmodulin has been used to develop calcium detection probes termed “chameleons” [13].

1.4 The RRC A straightforward approach to describe the FRET state of a system is to calculate the resonance energy transfer ratio change (RRC). The RRC provides a convenient means to measure the dynamic range of a FRET pair. For example, the maximum dynamic range of a small molecule indicator usually differs greatly from that of a protease probe, and this difference is reflected in the RRC. The FRET ratio is defined as the fluorescence intensity at the acceptor’s emission maximum divided by the intensity at the donor’s emission maximum. The RRC is the FRET ratio in the FRET-on state divided by the FRET ratio in the FRET-off state (Fig. 2).

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Fig. 2 Relevent intensity measurements for the measurement of the RRC. The RRC is given by the product RRC=(IC /ID )/(IB /IA )= (IA × IC )/(IB × ID )

2 Fluorescent Protein FRET Pairs The first fluorescent protein pairs demonstrated to be capable of FRET include BFP– GFP [14] and CFP–YFP [14] and later GFP–DsRed [15]. While these pairs exhibit spectral overlaps appropriate for energy transfer, each has unique advantages and disadvantages. Both BFP and CFP possess particularly low quantum yields [7]. Consequently, detection of blue or cyan fluorescence in the FRET-off state is difficult. The use of YFP is also problematic since it is pH and chloride sensitive, as well as susceptible to photobleaching [16]. The detection sensitivity of both BFP–GFP and CFP–YFP is also limited by increased cellular autofluorescence upon excitation with light sources compatible with CFP or BFP [17, 18]. Though DsRed does not exhibit the above problems, the obligate tetrameric form of the protein is incompatible with most fusion protein experiments [19]. In addition, the slow maturation time and intramolecular FRET that occurs in the DsRed tetramer complicates GFP–DsRed FRET measurements [19]. Improvements in FRET detection have been obtained using variants of the original fluorescent proteins. Brighter forms of both CFP and YFP, known as Cerulean [20] and Venus [21], respectively, have been developed and improve the signal-tonoise observed using CFP–YFP pairs [20]. Mutations in Venus also confer improved chloride sensitivity, maturation rate, and pH sensitivity [21]. Variants of the CFP– YFP pair have been used successfully to investigate protein–protein interactions [22, 23], detect interactions of small molecules with binding proteins [10, 11, 13], and even screen for protease inhibitors [24]. The development of a green–red pair has proven challenging. Commonly used Aequorea victoria-based GFPs (e.g., avGFP (F64L, S65T) [25]) could be useful as FRET donors, since they possess relatively sharp excitation and emission peaks, a high quantum yield, and an excitation wavelength maximum that yields reduced autofluorescence. However, the DsRed tetramer is unsuitable as a FRET acceptor because it often induces intolerable levels of aggregation. Monomerization was


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achieved through extensive directed evolution, resulting in the monomeric red fluorescent protein mRFP1 [26]. Though mRFP1 has proven useful in a variety of labeling applications, the GFP–mRFP1 FRET pair does not yield sufficient overlap for FRET. Other monomeric RFPs have been developed recently [27, 28] and may lead to the development of suitable FRET acceptors. Fluorescent proteins derived from stony corals have been shown to undergo efficient FRET. Cyan and orange fluorescent proteins MiCy and mKO, respectively [8], were isolated independently and possess spectral properties appropriate for energy transfer. In fact, MiCy–mKO has a high ratio change (14-fold) and exhibits some advantages over CFP–YFP, such as reduced cross-excitation and pH sensitivity [8]. Yet, MiCy is an obligate dimer, and thus is less useful in some experiments involving fusion proteins. Directed evolution of MiCy to a monomeric form might make this pair more useful if the high RRC can be maintained.

3 Enhancing FRET Through Directed Evolution The general applicability of the FRET pairs discussed above is limited by their oligomeric state or relatively small signal change between the FRET-on and FREToff states (Table 1). Pairs are typically chosen in a mix-and-match style, without consideration for the interactive nature of FRET between the donor and acceptor proteins. Ratiometric signal changes for monomeric pairs chosen in this way do not exceed fivefold. Table 1 RRC values of fluorescent protein FRET pairs in protease detection assays Donor FP

Linker length (aa)

Acceptor FP

RRC

References

BFP CFP Cerulean CFP CFPC7 MiCy∗ CyPet

25 20 20 20 6 14 20

GFP YFP YFP Venus Venus mKO YPet

5 3 2 5 10 14 20

[14] [29] [29] [29] [30] [8] [29]

∗

MiCy is an obligate dimer.

3.1 Linker Length Optimization Directed evolution has been applied to optimize FRET between coupled donor and acceptor pairs. In one study, the length of a linker region that encoded a caspase3 cleavage site between CFP and Venus was varied to generate a small library of deletion mutants [30]. Colony-based screening of the resulting variants resulted in the isolation of a CFP deletion mutant in which seven unstructured residues were removed and the protease susceptible linker between CFPC7 and Venus was significantly shortened. These changes nearly doubled the dynamic range relative to a

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CFP–Venus fusion with an 18 amino acid linker [30]. This surprising result would have been difficult to predict using rational design and exemplifies the importance of screening for FRET itself rather than simply the properties of the individual fluorescent proteins.

3.2 Directed Evolution of the CFP–YFP Pair A fluorescent protein pair possessing a substantially enhanced dynamic range was discovered by mutagenesis of a CFP–YFP fusion and screening of large libraries using fluorescence-activated cell sorting (FACS) [29]. CFP and YFP were mutated individually and screened for improved FRET and brighter whole-cell fluorescence when either the CFP–YFP fusion (FRET-on) or the CFP alone (FRET-off) was expressed. Flow cytometric screening of Escherichia coli cells expressing the constructs enabled screening of more than 108 variants for improved brightness and RRC. Appropriate gating of the cell population resulted in gradual enrichment of brighter clones (Fig. 3). The resulting variants, CyPet and YPet (cyan and yellow

Fig. 3 An example of FACS of fluorescent protein mutants. A pool of CFP mutants (a) is compared to the population after one (b), two (c), and three (d) rounds of sorting. Figure reprinted from Nguyen et al.29


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fluorescent proteins for energy transfer), exhibited a RRC of 20-fold when measured after trypsin cleavage in vitro [29]. Measurement of the signal change in living human embryonic kidney cells demonstrated significantly improved signal change of a CyPet–YPet caspase-3 substrate upon apoptosis induction (Fig. 4). The CyPet–YPet pair enables generation of a wide variety of high FRET efficiency protease probes. Various linkers have been demonstrated to yield RRC values similar to those obtained using the caspase-3 substrate (Table 2). The improved efficiency of CyPet–YPet also extends the range of substrate lengths that can be used as linkers between the fluorescent proteins. Linker lengths of up to 40 amino acids have been tested between CyPet and YPet and enabled detection of FRET (AWN unpublished data). Recent work in our laboratory with CyPet and YPet has shown that the improvement in FRET efficiency translates to protein interactions and periplasmic binding protein-based small molecule indicators. As a model system for protein interactions, two known interaction partners, an SH3 domain (Mona) and Mona binding peptide (P2), were fused to YPet and CyPet, respectively. Coincubation of the YPet-Mona

Fig. 4 Apoptosis detection using fluorescent protein FRET. Upon apoptosis induction (b, d) cells expressing the CFP–YFP (a, b) or CyPet–YPet (c, d) caspase-3 probe experience a shift in fluorescence signature. Figure printed from Nguyen et al.29

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A.W. Nguyen et al. Table 2 Linker sequences tested between CyPet and YPet with trypsin cleavage Substrate

Sequence

RRC

Gly-Ser Caspase-3∗ Enteropeptidase∗ TEV-protease∗ HIV-protease Amyloid-β

GGSGSGTGGSGSGGSGSGSGGS GGSGSGTGDEVDVPRGSGGS GGSGSGTGDDDDKGSGSGGS GGSGSGTENLYFQSGGSGSGGS GGSGSGTVSFNFPQITLGSGGS GGSGSGTKLVFFAEDGSGSGGS

18.7 20.2 21.2 18.2 18.1 20.3

∗

The caspase-3, enteropeptidase, and TEV-protease substrates have also been tested with their respective native proteases.

fusion and the CyPet-P2 fusion resulted in a 14-fold ratio change relative to YPetMona incubated with CyPet alone (XY unpublished data). This large dynamic range enabled high-throughput detection of FRET in bacteria, yeast, and mammalian cells (XY unpublished data), allowing high-resolution screening for partners of a given bait, analogous to the yeast two-hybrid system [31]. The FRET approach should prove useful for interaction screening since it offers the potential to quantitatively screen for reversible interactions and directly determine binding constants. Fusion of CyPet and YPet to maltose binding protein (MBP) as previously described [10] resulted in a significant signal change improvement upon addition of maltose. The CFP–MBP–YFP fusion yielded a RRC of 1.04, while CyPet–MBP– YPet yielded a ratio change of 1.59 over the maltose concentration range of 0.001– 10000 μM (Fig. 5). The dynamic range typical of CFP–YFP periplasmic binding

Fig. 5 Shift in FRET signal using a MBP indicator. CFP and YEP (squares) or CyPet and YPet (circles) were used to create maltose indicators that shift fluorescence with the addition of maltose


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protein fusions hinders utilization of these sensors because the noise in the detection system must be very low to allow detection of the signal change. Substitution of CFP–YFP with CyPet–YPet improves signal-to-noise ratios significantly, enabling detection of subsaturating levels of analyte with greater accuracy.

4 Using FRET to Screen Libraries High-throughput screening based on fluorescent protein FRET has been used to detect known interactions, but could potentially be applied for library screening and directed evolution. Fluorescence-activated cell sorting (FACS) is one of the fastest and most quantitative methods to search through cell-based protein libraries for a specific function. A high-end cell sorter can sort individual cells based on fluorescence at a rate of 100,000/s [32]. Requirements of FACS include a traceable link between phenotype and genotype and a detectable fluorescence signal change indicating function. Compartmentalization within a single cell is a simple way to contain the gene, protein, and fluorophore, though innovative methods such as emulsion encapsulation have also been used [33]. In many library screening applications, an intracellular fluorescent probe is used to screen variants in a high-throughput manner. Yet there are a limited number of probes that have been developed with these traits. In many cases, this problem can be solved using fluorescent protein FRET probes. For example, introduction of a protease cleavage site between two fluorescent proteins capable of FRET creates a fluorescent protease probe that remains within the cell allowing FACS. Until recently though, attempts at using fluorescent protein FRET in combination with cytometry have indicated that the dynamic range of CFP–YFP was not sufficient for FACS-based screening in bacterial or yeast cells and suboptimal for mammalian cell screening (Fig. 4). FRET has been used for colony and microtiter plate imaging and screening, though only a small number of variants are typically analyzed. In one example, described above, a linker library consisting of 88 linker lengths between CFP and YFP was screened using colony imaging [30]. This resulted in the CFP7Venus clone with an improved RRC (Table 1). A proof-of-principle experiment using a CFP–YFP caspase probe in Jurkat cells has also been used to screen 480 known compounds for apoptosis-inducing activity in microtiter plates. The CFP– YFP sensor was sufficiently robust when compared with more typical in vitro peptide cleavage assays and provided the advantages of cell-based screening [34]. Additionally, novel apoptosis inhibitors were identified using Hela cells in 96well plates [24]. Two compounds were identified that inhibit apoptosis, where one inhibits caspase activity directly and the other is thought to inhibit an upstream apoptosis-inducing event. The directed evolution of CyPet and YPet is the first example of using FRET to screen large libraries by FACS. This screen involved the interrogation of greater than 108 individual cells, each expressing a different variant. Though sorting of the CFP–YFP fusions for improved FRET was possible, the sought-after signal changes

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were quite small when compared with many FACS experiments. Each round of mutagenesis and screening resulted in only about twofold improvements in FRET signal (Fig. 3). Sorting the leading edge of the population, regrowing and repeating multiple times was required for successful enrichment [35]. And, importantly, this strategy should be useful in a wide variety of screening applications where signals differ by as little as twofold.

5 Future Opportunities for FRET-Based Screening 5.1 Protease Evolution Sorting of improved FRET pairs was the first step toward using fluorescent protein FRET as a screening method for detection and evolution of proteins of interest. Specifically, application of fluorescent protein FRET for FACS-based protease assays could be a substantial improvement over current methods. This approach could be used to redesign protease specificity or turnover rate or screen for new protease activities from a naïve library. Protease screening is currently limited by available protease probes. Inserting a protease susceptible linker between CyPet and YPet provides a simple and inexpensive means to create a probe compatible with FACS, fluorescence spectroscopy, or fluorescence microscopy. And while construction of such substrates should be generalizable, potential caveats should be considered. The protease of interest must be compatible with a dumbbell-like substrate, flanked by two-folded fluorescent proteins. Thus, for example, the enzyme should not require threading of small substrates into the protease structure [36]. Furthermore, it is necessary that the protease be expressed inside of a host cell in an active form. Some proteases would be expected to lead to cellular toxicity, in which case PCR can be used to amplify variants of interest if the cells remain intact [37]. The detection of caspase-3 activation in human embryonic kidney cells with the CyPet–YPet substrate (Fig. 4) indicates that the signal change is sufficient for screening, via FACS, mammalian cells that possess caspase activity. Smaller cell types including yeast or bacterial cells present unique challenges. Since FRET is highly distance dependent, high concentrations of donor and acceptor molecule can result in non-specific FRET even when they are expressed independently. At the same time, a sufficient level of fluorescence is required in each cell for cytometry detection. In mammalian cells, this balance is achieved and with moderate levels of fluorescent protein expression; the signal is detectable and non-specific FRET does not occur in the FRET-off state. However, a bacterial cell (approximately 3 × 1 μm [38]) is on the order of 1000-fold smaller volume than a typical mammalian cell (assuming a 20 μm diameter). Thus, expression of detectable levels of fluorescent protein while maintaining low FRET signal in the FRET-off state is challenging. To investigate whether FACS could be used as a tool to screen protein libraries in microbial cells, we compared FRET signal of E. coli cells that express either CFP–YFP (Fig. 6b) or CyPet–YPet (Fig. 6d) fusions to that of bacteria that express


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Fig. 6 Comparison of FRET signals between CFP–YFP (a, b) and CyPet–YPet (c, d) expressed in the cytoplasm of E. coli. Fluorescent proteins were coexpressed independently (a, c) or as a fusion (b, d) within E. coli

unfused partners (Fig. 6a and c) using flow cytometry. This approach indicates the maximum signal change, representing complete substrate cleavage. Using the parental CFP–YFP variants, the two-parameter fluorescence distributions of cells expressing fused partners significantly overlaps with that of cells expressing CFP and YFP independently. The populations are not sufficiently discriminated for FACS. On the other hand, using CyPet and YPet, a greater than 40-fold increase in FRET positive cells is observed (Fig. 6c and d) using optimized expression conditions. These data indicate that CyPet–YPet FRET pairs will enable screening of cell-based libraries by FACS.

5.2 Protein Interaction Screening Fluorescent protein FRET is frequently used as a tool to investigate known protein interactions in their native cellular context, using gene fusions to the interaction partners. Thus far, however, these studies have been limited to validation of known or putative interactions. The development of improved FRET partners, as described above, offers the potential to extend this approach to the screening of libraries to identify novel binding proteins for a given target. Such an approach may prove useful for large-scale interactome screens in a variety of cell types. Our preliminary studies in mammalian, yeast, and bacterial cells indicate that FRET efficiency

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is sufficient to enable FACS-based screening random libraries to identify novel protein–peptide interactions (unpublished data). Investigation of protein–protein interactions in the intracellular environment is advantageous since it can provide insight into co-localization and physiological relevance that a test tube cannot.

5.3 Small Molecule Indicator Development Significant interest exists in creating new FRET-based small molecule binding probes enabling quantitative intracellular measurements of concentration dynamics and localization in the context of a native cell. One application, for example, is to change the binding specificity of periplasmic binding proteins such as maltose binding protein (MBP) in order to recognize a small molecule of interest. The conformational change that occurs upon binding of the analyte must be retained to ensure that a FRET signal change still occurs. Mutagenesis of MBP, followed by cytometric screening for FRET in the presence of a different small molecule, could be used to create new indicators. Since the dynamic range of the CyPet–MBP–YPet maltose sensor is significantly improved when compared with the CFP–MBP–YFP probe, this signal change may be sufficient for screening large cell populations using FACS. Experiments with the CyPet–YPet pair in protease and protein–protein interaction studies indicate that larger FRET signal changes should be obtainable if the distance change between CyPet and YPet upon binding can be increased. Thus, evolution of MBP toward larger RRCs with CyPet and YPet provides a potential means to improve and expand this approach for construction of new small molecule indicators. Mutagenesis of MBP and alternating screening in the presence and absence of maltose for high FRET signal and low FRET signal, respectively, should enable isolation of improved MBP sensors. These enhanced FRET indicators could, then, be used to screen for new binding activities toward other small molecule ligands.

5.4 Long Wavelength FRET Pair Optimization Another important goal for FRET screening applications is the development of longer wavelength FRET pairs. Though cytometry and microscopy instrumentation manufacturers have begun to incorporate violet light sources for CFP excitation into their platforms, a need for tissue penetration and excitation at the common 488 nm wavelength persists. The procedure used for evolution of CFP and YFP for higher efficiency FRET could also be applied to a GFP–RFP pair. Though a dimeric RFP that already functions as a FRET acceptor for GFP could be used as a parent in directed evolution, the identification of highly efficient monomeric acceptor may prove challenging. Instead, one could begin from a monomeric RFP with little FRET signal from GFP and evolve toward improved acceptor brightness in the FRET-on state. The evolution of an efficient GFP–RFP pair would ease experiments involving


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labeling with two sets of FRET probes in the same cell or heterotrimer formation [39] wherein FRET occurs from CyPet→YPet→RFP.

6 Conclusions The use of fluorescent protein FRET as a tool in molecular biology studies is rapidly growing. One further indication that the full potential of the tool has not yet been achieved is that many studies are focused on improving FRET interactions and FRET detection methods. High-throughput screening on the basis of fluorescent protein FRET is now possible and new detection methods, such as fluorescence polarization [40], may enhance separation of FRET-on and FRET-off states further. The pioneering steps that were made in screening hundreds of protease inhibitors using FRET [24] can now be expanded to screen over a billion proteins of interest for a desired function. Flow cytometric screens for protease activity or protein–protein interactions can both be performed on the basis of FRET between CyPet and YPet. These advances will enable large-scale studies of binding and protease specificities, enhance small molecule detection technologies, and provide new opportunities in protein evolution.

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Imaging Protein Interactions in Living Cells Using the Fluorescent Proteins Richard N. Day, Ammasi Periasamy, and Ignacio Demarco

Abstract Over the past decade, the visible fluorescent proteins (VFPs) have become an essential tool for the study of cell biology and physiology. The goal of this chapter is to describe the characteristics of the FPs and discuss the limits of their use as probes for noninvasive imaging of protein behavior in living cells. We also described live-cell imaging techniques used to monitor the subcellular dynamics and interactions of proteins in living cells. We then focused on the method of fluorescence resonance energy transfer (FRET) microscopy to quantify the interactions of proteins inside living cells and consider some of the factors that limit this approach.

1 Introduction It would be very difficult to overstate the impact that the fluorescent proteins (FPs) have had on the biomedical sciences. The diminutive green glow of the jellyfish Aequorea victoria has sparked a revolution in studies of cell biology and physiology. The glow comes from an autofluorescent protein produced by the jellyfish that is a companion for a photoprotein, the bioluminescent aequorin. This jellyfish green fluorescent protein (GFP) was first isolated and characterized by Shimomura [1] and the gene encoding Aequorea GFP was cloned in 1992 by Prasher et al. [2]. Gene expression experiments in bacteria, animal, and plant cells proved the utility of the expressed protein as an intravital marker [3–5]. What’s more, their successful integration into a variety of transgenic organisms has demonstrated that the FPs are well tolerated in living systems (reviewed by Hadjantonakis[6]). For example, the stable expression of the FPs in targeted cells in transgenic mice has allowed repeated microscopic imaging of the cells over periods of days to weeks [7–9]. Moreover,

A. Periasamy (B) Keck Center for Cellular Imaging, Department of Biology, Gilmer Hall, University of Virginia, Charlottesville, VA 22904 e-mail: [email protected] This chapter is dedicated to our colleagues at Tulane University who lost so much in the aftermath of hurricane Katrina, August 29, 2005. C.D. Geddes (ed.), Reviews in Fluorescence 2008, Reviews in Fluorescence 2008, DOI 10.1007/978-1-4419-1260-2_14, C Springer Science+Business Media, LLC 2010

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the FPs are routinely used in combination with a variety of different imaging methods to track the functional recruitment, co-localization, and interactions of specific protein partners within subcellular compartments in living cells. These cellular imaging methods are now an indispensable complement to the biochemical techniques that are traditionally used to define protein interactions. Importantly, these live-cell imaging approaches extend quantitative measurements of protein behavior to specific subcellular sites within the living cell (reviewed in [10, 11]). The modifications to the Aequorea GFP and the cloning of new color variants that are discussed below have supplied the research community with genetically encoded proteins that emit light from the blue to the red range of the visible spectrum. The goal of this chapter is to describe the characteristics of the FPs and discuss the limits of their use as probes for noninvasive imaging of protein behavior in living cells. We will describe live-cell imaging techniques used to monitor the subcellular dynamics and interactions of proteins in living cells. We then focus on the method of fluorescence resonance energy transfer (FRET) microscopy to quantify the interactions of proteins inside living cells and consider some of the factors that limit this approach.

2 Spectral Variants of the Fluorescent Proteins The characteristic glow of the FPs requires that the proteins form an 11-strand beta-barrel structure that positions a three amino acid chromophore, serine-65tyrosine-66-glycine-67, at its core. Nearly the entire 238 amino acid sequence is necessary for the protein to adopt this structure (reviewed by Tsien [12]). Many different laboratories contributed to the extensive engineering of the Aequorea GFP, and this yielded new proteins with improved characteristics that emit light from the blue to the yellowish green range of the visible spectrum (reviewed in [13, 14]; see Table 1). For example, changing the Aequorea GFP serine at position 65 to threonine promoted ionization of the chromophore, shifting the peak absorbance from 395 to 489 nm, and resulted in a more rapidly maturing protein [15–17]. Additional modifications to incorporate preferred human codon usage and to improve the maturation and stability of the protein at physiological temperatures resulted in proteins called the “enhanced” FPs (e.g., EGFP). Color variants of GFP were generated by mutagenesis of residues in the chromophore and the surrounding beta-barrel. The availability of new color FPs expanded their repertoire of uses to include multicolor imaging of protein colocalization and, importantly, their application in FRET microscopy to measure protein–protein interactions (these applications are discussed below). For example, a blue color variant (BFP) resulted from substitution of the tyrosine at position 66 with histidine [17, 18]. While BFP has a low quantum yield and is prone to photobleaching, it remains the most blue-shifted FP and has proven useful in both multicolor and FRET imaging [19, 20] (Table 1). The substitution of tyrosine at position 66 with tryptophan in combination with mutations in several other residues within the surrounding beta-barrel structure yielded a cyan FP (CFP) that provided an alternative to the dim and photolabile BFP [15, 17].

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Table 1 The characteristics of the Aequorea-based FPs, DsRed, mRFP, and the new generation of FPs based on mRFP FP

Ex (nm)

Em (nm)

Photobleach sensitivity

Intrinsic brightnessa

Distinctive characteristics

Aequorea-based FPs BFP

382

446

High

7b

ECFP

434

476

Low

10

EGFP

489

509

Low

33

Venus

514

527

Moderate

52

Dim, spectrally distinct, FRET donor Brighter, complex Ex state, FRET donor Bright, stable, FRET donor or acceptor Very bright, FRET acceptor

DsRed-based FPs DsRed

558

583

Low

45

mRFP

584

607

Moderate

11

Tetramer, slow maturation, green intermediate Monomeric red, nonfluorescent intermediate

mRFP-based FPs mOrange

548

562

Moderate

49

tdTomato

554

581

Low

95

mCherry mPlum

587 590

610 649

Low Low

16 9c

a Intrinsic b Cellular

Very bright, FRET acceptor for GFPs Brightest of current FPs, obligate dimer Bright red, fast maturing, stable Deepest red, largest Stokes shift, stable

brightness is the product of quantum yield × extinction coefficient. autofluorescence, except in the cell nucleus is high at these wavelengths, reducing signal

to noise. c Cellular autofluorescence is low at longer wavelengths, improving signal to noise.

The longest wavelength emission of the Aequorea-based FPs, the yellowish FP (YFP), resulted from the targeted mutation of the beta-barrel at position 203, which was identified in the crystal structure as being near to the phenol ring of the chromophore [21]. The enhanced YFP (EYFP) produces bright fluorescence, but its utility is limited by pH and halide sensitivity [12]. Starting with the EYFP, Nagai et al. [22] discovered that substitution of the phenyalanine at position 46 with leucine improved the maturation efficiency and reduced the halide sensitivity. Several other substitutions lead to further improvement, resulting in the protein called Venus, the brightest and most red shifted of the mutant variants based on the Aequorea GFP available (Table 1). Many marine organisms exhibit fluorescence and most of the natural FPs that have been characterized are either dimers, tetramers, or higher order complexes [23, 24]. Although the Aequorea GFP could be crystallized as a monomer, the proteins can form dimers at high concentrations [12, 16, 21]. The dimerization domain is a hydrophobic patch that is found at the carboxyl-terminus of all the Aequorea-based

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FPs [25]. The dimerization of the Aequorea-based FPs is not typically observed when the proteins are free to diffuse within the cell. For instance, recent fluorescence fluctuation spectroscopy measurements of EGFP expressed in the cytoplasm of living cells over the concentration range of 50 nm to 10 M (M - molar) showed no evidence for dimer formation [26]. The tendency to form dimers, however, becomes an important consideration when the FP-fusion proteins are expressed at high concentrations in a restricted volume, such as in the two-dimensional space of biological membranes. Here, the dimerization of the FPs can cause the fused proteins to form atypical complexes [27]. Because of these concerns, Zacharias and colleagues [25] showed that substitution of the hydrophobic patch residues in the beta-barrel with positively charged amino acids prevented the dimerization of the FPs. For example, the substitution of alanine at position 206 with lysine blocked dimer formation without changing other characteristics of the FPs, leading to the development of the monomeric Aequorea-based FPs [25].

2.1 The Red Fluorescent Proteins Most of the colors that we see in reef corals come from GFP-like proteins. This observation led to the cloning of several different red fluorescent proteins (RFP) from Indo-Pacific corals that added to the spectrum of available FPs [24, 28, 29]. The first of the commercially available RFPs, called DsRed, was cloned from the mushroom anemone Discosoma striata. When it is expressed in mammalian cells, DsRed is an obligate tetramer that requires nearly 20 h to fully mature [30]. In addition, there is a tendency for DsRed to form oligomers, and this can lead to the aberrant behavior of the proteins that are fused to DsRed. What’s more, a green intermediate is formed during the slow maturation, and this has limited the utility of DsRed for co-localization and protein interaction studies. Several different laboratories used mutagenesis to overcome the deficiencies of DsRed. Using a strategy of both random and directed mutagenesis, Bevis and Glick [31] were able to solve the problem of slow maturation, generating the fast maturing ( (2-F)Tyr-protein > (3-F)Tyr-protein. This corresponds well with the pKa values of free amino acids, Tyr (10.0) > (2-F)Tyr (9.04) > (3-F)Tyr (8.5) [67]. It is worth noting that the pKa value of fluorescence in (3-F)Tyr-EYFP is considerably decreased by about 1.2 pH units which should be at least in part attributed to the stacking interactions with (3-F)Tyr203. There are several possible effects of the substitution of fluorine on the behavior of fluorinated chromophores in addition to a decrease in pKa . The most important one seems to be the observed blue-shifts of (2-F)Tyr-variants, and red-shifts for (3-F)Tyr-variants in both EGFPs and EYFPs. The measured spectral red- and blue-shifts can be regarded as intrinsic features of the fluorinated chromophores with significant spectral enhancements observed in the EYFP variants at a neutral pH [25]. The complex photophysics of the chromophore is usually explained by the threestate photoisomerization model assuming that the chromophore exists in a neutral (395 nm, AH) or in an anionic (475 nm, A− ) state; the latter state exists in a thermodynamically unstable intermediate form (493 nm, I-state) and in a stable low-energy form [53]. Spectroscopic analyses and quantum mechanical calculations suggest that they represent chromophores with protonated and deprotonated tyrosyl hydroxyl groups, respectively [69]. Fluorine being the most electronegative element is expected to exert strong inductive electron-withdrawing effects on the fluorinated chromophore ring. By this, it becomes relatively electron deficient, and more hydrophilic, which makes its hydroxyl moiety more acidic. Both excited and ground states of the phenolic oxygen of Tyr66 would be certainly influenced by such almost isosteric H→F single atom exchange. This is because the electron-withdrawing fluorine atom in the meta-position is positioned in the vicinity of the phenolic

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Fig. 4 Structural and spectroscopic aspects of Tyr-fluorination of EGFP. Global replacement of Tyr-residues in both proteins by (2-F)Tyr, and (3-F)Tyr, respectively, results in concomitant fluorination of the chromophores. Upper part – left: The general architecture of the avGFPs is based on the 11 stranded β-barrels that rigidly hold a chromophore anchored at a coaxial helix within the protein core. Upper part – middle: Difference Fourier maps of (3-F)Tyr-chromophore in (3-F)Tyr-EGFP superimposed on its continuous electron density revealed the existence of two conformers (“major” and “minor”). In contrast, X-ray crystallographic structure analysis of (2-F)Tyr-EGFP mapped a single conformer state of (2-F)Tyr-containing chromophore. Upper part – right: Global replacement of EYFP with (3-F)Tyr and (2-F)Tyr yielded red- and blue-shifted chromophore. Spectral properties are the following: wt-EGFP – λmax(ex) = 488 nm, λmax(em) = 510 nm; (2-F)Tyr-EGFP – λmax(ex) = 482 nm, λmax(em) = 504 nm; (3-F)Tyr-EGFP – λmax(ex) = 485 nm, λmax(em) = 514 nm; wt-EYFP – λmax(ex) = 514 nm, λmax(em) = 527 nm; (2-F)Tyr-EYFP – λmax(ex) = 504 nm, λmax(em) = 520 nm; (3-F)Tyr-EGFP – λmax(ex) = 518 nm, λmax(em) = 533 nm. Lower part: Schematic representation of the protein–matrix interactions in native and fluorinated EGFP variants. Note novel interactions in the chromophore microenvironment brought by the fluorine atoms

hydroxyl group and enhances the acidity of (3-F)Tyr-chromophore to a larger extent than (2-F)Tyr-chromophore. This is indeed plausible, since pKa of free (3-F)Tyr is 8.5, which is 1.5 pH units lower than that of Tyr (10.0) [67]. Following this logic of reasoning, it is obvious that a deprotonated anionic A− of the chromophore is much more favored upon its fluorination at meta-position than its protonated or neutral (AH) form. Therefore, red-shifted spectra of (3-F)Tyr-containing proteins (4–6 nm, see Fig. 4) can be attributed to their enhanced preference for the anionic A− state

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of the chromophore. However, one might expect the same effect for (2-F)Tyrcontaining proteins, but to a lower extent. Instead, spectra of these proteins are blue-shifted (6–7 nm, see Fig. 4), and there should be other reasons for these observations. In addition to the inductive electron-withdrawing interactions, the resonance effect induced by fluorine as well as by the OH group might also influence electronic properties of the aromatic ring in chromophore. This aromatic ring is influenced by the electron-donating resonance effect of the hydroxyl group as well. Such resonance effect of both groups is reflected in the distribution of their lone pairs to the ring carbons. These effects together are responsible for significantly altered biophysical properties of (3-F)Tyr relative to those of Tyr. On the other hand, a combination of these effects in (2-F)Tyr-chromophores most probably causes a different balance that is responsible for their intrinsic spectral blue-shifts. Furthermore, it is likely that inductive electron-withdrawing effects caused by the (2-F)Tyr in the chromophore outweighs the resonance effects to a lower extent than does (3-F)Tyr. Thus, it should not be surprising that biophysical properties of (2-F)Tyr and related chromophores are also much more similar to the parent structures. The most interesting observation from the available structure of (3-F)Tyr-EGFP is the existence of two conformers of the fluorinated Tyr66, in the context of chromophore [68]. A rather rigid internal architecture in the crystalline state of the fluorinated chromophore and its surrounding residues is reflected in the average crystallographic B factors for atoms in the chromophore which are 15 Å2 , while the crystallographic B factors of the surrounding residues are from 5 to 25 Å2 . In such a rigid environment (3-F)Tyr145 has only one conformer which is plausible, since it is involved in interactions with other residues and the solvent water. The crystallographic occupancy (i.e., intensities of electron densities) is higher (about 60%) for one conformer (assigned as “major”) than for the other (40%, “minor”) (Fig. 4). The hydroxyls of both (3-F)Tyr in the chromophore conformers interact directly with Thr203 and His148 through hydrogen bonds. Fluorine, when fitted in the “minor” (3-F)Tyr-chromophore conformer is in hydrogen-bond contact (2.82 Å) with the hydroxyl of Ser205 and conserved solvent water (S316, 2.74 Å). On the other hand, Cγ2 of Thr203 is 3.87 Å distant from the fluorine of the “major” conformer indicating a possible weak –CF–HC- hydrogen bond (Fig. 4). In the light of these observations, it is difficult to explain the higher crystallographic occupancy of the fluorine atom position in the “major” conformation. The crystal structure of (3-F)Tyr-EGFP [68] enabled “visualization” of the positions of some of the hydrogen atoms in the chromophore due to their replacement by fluorine atoms (Fig. 4). Does it mean that at least in part, the chromophore of the wt-EGFP flips in its cavity? Modeling and molecular dynamic simulation studies indicate that avGFPs have a fairly large central cavity in the protein core, which allows the planar chromophore to have some rotational freedom, especially in the ϕ dihedral angle. Electronic excitation (SO →S1 ) was found to alter the conformation of the chromophore. It was speculated that in the excited state, (responsible for the characteristic fluorescence), the conformation may be twisted relative to the ground state. The cis–trans photoisomerization cannot occur at a 180◦ rotation

380


of the τ dihedral angle of the chromophore [70]. Thus, it might well be that the observed conformers represent Tyr-flipping “caught” or “frozen” by the chromophore formation and maturation. Other alternatives include the possibility that the measured electron difference maps indeed represent an isomerization of a mature chromophore. The crystal structure of (2-F)Tyr-EGFP at 2.2 Å is the first known protein structure substituted with this analog [25]. This has surprisingly shown that all (2-F)Tyr-residues are present in only one conformeric state. The single conformer of the (2-F)Tyr-chromophore in (2-F)Tyr-EGFP can be plausibly explained by taking into account sterical considerations. Fluorine, when fitted in the “second” orthoposition of (2-F)Tyr-chromophore induces direct clashing with the hydrogen of the ring nitrogen (in fact, there is a distance of about 2.1 Å between the nitrogen and the fluorine). This might offer a plausible explanation for the exclusive presence of the single conformer, not only in (2-F)Tyr-chromophore but also of (2-F)Tyr-residues in (2-F)Tyr-EGFP. Although the crystal structures of fluorinated EYFP variants are not available, it is reasonable to suppose that the most of the fluorotyrosyl residues behave in the same way as those in fluorinated EGFPs. However, the most interesting question is regarding the behavior of fluorinated chromophore and Tyr203. Results of modeling and energy minimization procedures performed with planar chromophore of EYFP show a very tight packing. In other words, there is no rotational freedom for flipping of the (3-F)Tyr-EYFP chromophore in the same manner as for the (3-F)Tyrchromophore in EGFP despite the fairly large central cavity in the protein core (Fig. 5). The main restraint on any flipping is the nearby positioning of Tyr203 which is in stacking interaction with the chromophore of EYFP.

Fig. 5 Structural and spectral responses of avGFPs upon Tyr-fluorinations. Left: Modeled chromophore cavities of (3-F)Tyr-EGFP and (3-F)Tyr-EYFP (chromophores and amino acids Thr203 and (3-F)Tyr203 are shown as stick representations). Right: Fluorescence emission profiles of wt and substituted ECFP, EYFP, and EGFP at neutral pH. Note that ECFP was used as a control. It contains the same number of Tyr-residues (10) as EGFP while the 11th one is replaced with Trp at position 66 (chromophore). As expected, global fluorination with both (3-F)Tyr and (2-F)Tyr influences neither absorbance nor fluorescence properties of ECFP


381

8.3 Fluorination of the Chromophore Environment in EYFP By virtue of its sequence mutation Thr203Tyr, EYFP is known to be the most redshifted mutant designed by conventional methods (λmax(em) = 527 nm) [12]. The fluorination of Tyr203 in EYFP offers a fairly good model to study effects caused by fluorinations of its microenvironment on the spectral functions of the chromophore. Fluorinated EYFPs (having the chromophore in stacking interaction with Tyr203) offer a framework to study the nature of the lowering of pKa upon fluorination. It is reasonable to expect that inductive effects exerted by fluorine nucleus on the chromophore or fluorine atoms from its microenvironments are responsible for such behavior. Thus, the fluorination of Tyr203 is most probably responsible for the enhanced blue- and red-shifts in the EYFP variants (Fig. 5). The Tyr-side chain at position 203 stacks next to the chromophore to induce additional polarizability in its microenvironment; π–π interactions are expected to reduce excited-state energy and enhance both excitation and emission wavelengths [71]. Undoubtedly, all these interactions are complicated and are in addition influenced by the introduction of the fluorine nucleus. (3-F)Tyr203 and (2-F)Tyr203 are mutually isosteric, and the differences observed in the related protein mutants may rely not only on the different physicochemical properties but also on the different stereochemical positions of fluorine atoms relative to the chromophore protonated/deprotonated hydroxyl moiety. As shown in Fig. 5, a closer inspection of the chromophore of (3-F)TyrEGFP in the cavity with Thr203 revealed that there are no steric hindrances in the accommodation of the fluorine atom (purple) in the cavity, and even for (3-F)Tyr-chromophore flipping. The model of (3-F)Tyr-EYFP was constructed by the replacement of a hydrogen by a fluorine at Tyr203 and Tyr66 in wt-EYFP (PDB entry: 2YFP), and by energy minimization using the standard modeling procedures. Although fluorine atoms can easily be accommodated in the chromophore cavity of EYFP, it is obvious that the flipping of the fluorinated chromophore should encounter steric hindrance even in the native EYFP structure. The coplanar position of both Tyr203 and (3-F)Tyr203 that is about 4 Å distant from the centre of chromophore should be the main barrier for possible flipping motions.

9 Perspectives and Outlook Chromophore-building residues of GFP are almost ideal models for the incorporation of noncanonical amino acids since the produced effects are immediately observable in the optical properties. Till now, extensive variations of the chromophore itself as well as its protein matrix by classical protein engineering approaches were performed. Indeed, classical protein engineering yielded several classes of avGFP mutants. In order to expand the scope of these experiments, classical methods (site-directed mutagenesis, guided evolution, random mutagenesis coupled with the screening, etc.) should be supplemented with methods and

382


approaches of the expanded genetic code. The most conceivable way to achieve this goal is to combine production of the mutant proteins in high-throughput manner which include the possibility for canonical → noncanonical amino acid replacements during protein expression. Although different methods and amino acids are available for such experiments, the most promising approach is the use of amino acid analogs and surrogates which mimic their natural counterparts. The example provided in this study suggests the possibility for a rational strategy in designing avGFPs with improved fluorescence and increased stability of the monomeric state. One might expect that future experiments on substitution with aminotryptophan analogs on known red-emitting FPs like DsRed will result in further red-shifted mutants. Furthermore novel generations of red-shifted autofluorescent proteins are also conceivable by the introduction of the novel substituents like cyano, nitro, nitroso, etc., in chromophores formed by indole, imidazole, phenol, and other aromatic rings. The translation of these substances in protein structure in combination with classical random and combinatorial mutational approaches may well pave the way toward novel families of tailor-made autofluorescent proteins. The design of GdFP might be seen as paradigm for the generation of novel avGFP classes by an expanded amino acid repertoire; GdFP itself might serve as an excellent model for photophysical studies of chromophore dynamics. However, its broader in vivo use as a marker for cellular biology is currently limited due to the special incorporation protocols. On the other hand, their intracellular synthesis could be assisted by supplementation of the system with plasmids encoding modified translation components (aminoacyl-tRNA synthetases, tRNAs, etc.) which are compatible with endogenous host transitional apparatus. Recent progress on the development of orthogonal aminoacyl-tRNA synthetases as well as their cognate tRNAs pairs for use in yeast [72] and even mammalian cells [73] is an example for efforts in this direction [74]. It is well known that ideal fluorophores are excitable with a single excitation color, while being easily separable in their emission wavelengths. This goal can be achieved for semiconductor nanocrystals which possess a common broad absorption and emission band depending only on their sizes. The biofunctionalization of these particles however still poses major problems, thus preventing their broad usage. avGFP mutants on the other hand can easily be fused to target proteins and subsequently be expressed in prokaryotic and eukaryotic cells. With the avGFP mutants available to date, however, only a pair of two variants can be excited with one color and detected at two different wavelengths. The three populations of Trp-auxotrophic E. coli ATCC49980 strain expressing ECFP, EGFP, and GdFP, respectively, clearly show how to overcome these limits (Fig. 2). This should open new ways for multiple labeling applications, design of FRET partners that enable measurements of molecular scale-distances or concentrations through changes in fluorescence. In the coming decades, tailor-made autofluorescent proteins like GdFP employing noncanonical amino acids will certainly be valuable as genetically encoded physiological indicators and tools.


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Fluorescent Proteins in Transgenic Plants Reginald J. Millwood, Hong S. Moon, and C. Neal Stewart Jr.

Abstract Fluorescent proteins (FPs) have revolutionized many areas of biological research. In particular, plant biotechnology has been significantly advanced by harnessing the power of FPs. Aequorea victoria, green fluorescent protein (GFP), has been the most studied of the proteins, but many new FPs are discovered each year. We provide here a timely review of the current uses of FPs in whole plants and we look at the color palette of candidate proteins. Lastly, we discuss current instrumentation and methods for detection and quantification of FPs in plants.

1 Introduction For nearly two decades, fluorescent proteins (FPs) have been invaluable tools in basic and applied scientific research. However, FPs are not new to science. In fact, they have been studied for more than 50 years. The most widely studied FP is the green fluorescent protein (GFP) from the bioluminescent jellyfish Aequorea victoria. GFP was first isolated by Osamu Shimomura at the Friday Harbor Laboratories, Washington, USA [1]. Shimomura was not interested in GFP as a biotechnology tool. Rather, his research interest was to understand the chemistry and biochemistry of A. victoria’s bioluminescence. It was not until Doug Prasher et al. [2] cloned the gfp gene that the utility of GFP was fully realized, which enabled the award of the Nobel Prize in Chemistry (2008) to three scientists (Osamu Shimomura, Martin Chalfie, and Roger Tsien), but ironically, not to Doug Prasher. For the first time researchers had accessibility to a marker gene that was useful in vivo, could be detected in real-time, and required no substrates or cofactors. Subsequently, the gene was expressed in Escherichia coli and Caenorhabditis elegans [3]. Shortly thereafter, the gfp gene was subjected to mutagenesis and several new variants R.J. Millwood (B) Department of Plant Sciences, The University of Tennessee, Knoxville, TN, USA e-mail: [email protected]


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were created ranging from blue to yellow [4, 5]. These events marked the beginning of what has often been called the “Green Revolution” of biotechnology. The foundation built with GFP research has allowed others to search for new FPs in other organisms [6–12]. At present, scientists have a wide range of FPs to choose from that work for many applications. Here we discuss a brief history of FPs, candidate FPs for plant expression, and FP applications in whole plants. Also included is a description of current instrumentation and methodologies of FP detection.

2 FPs in Model Organisms Because of established transformation protocols, model organisms were the first to be transformed with GFP. E. coli and C. elegans [3] were followed closely by fruitfly (Drosophila melanogaster) [13] and subsequently mammalian cell lines [14] and yeast (Saccharomyces cerevisiae) [15] . Larger organisms, such as tobacco (Nicotiana tabacum) [16] , mouse (Mus musculus) [17], zebrafish (Danio rerio) [18], frog (Xenopus laevis) [19], rhesus monkey (Macaca mulatta) [20], and pig (Sus domestica) [21] were transformed with GFP variants yielding visible green fluorescence. In addition to the many GFP organisms, a red FP (DsRed2) from the coral Discosoma sp. has been expressed in the domestic cat (Felis catus) [22]. The utility of FPs was quickly realized for applications outside scientific research. Two transgenic organisms in this area are particularly intriguing, the R first of which is a commercial ornamental transgenic organism, the “GloFish” (www.glofish.com), marketed by Yorktown Technologies (Fig. 1a). Under the control of a strong muscle-specific (mylz2) promoter, transgenic zebrafish were transformed with the green fluorescent protein (EGFP) and a red fluorescent protein (dsRed) [23]. GloFish appear brightly fluorescent green, red, or orange (GFP+RFP) when in the presence of an ultraviolet (UVA) aquarium lamp. The original intention for creating transgenic zebrafish was to use them as biosensors for pollutants. For example, the fish appear normal when no pollutants are present, but after pollutant exposure the fish would express an FP to give a visual signal. Zebrafish biosensor research is ongoing, but none have been deployed to date. However, the commercial value was immediately realized when their brilliant fluorescence was observed in the presence of a UVA lamp; these fish are sold to people interested in hobby aquaria. The second and most stunning of the transgenic FP organisms, “Alba,” the GFP bunny commissioned by the artist Eduardo Kac, was the central piece of the Eighth Day art exhibit (www.ekac.org/gfpbunny.html#gfpbunnyanchor). Alba, an albino rabbit, was transformed to express GFP and yield a remarkable fluorescent phenotype (Fig. 1b). This caused quite a stir because typically transgenic organisms are not created as works of art but to answer scientific questions. Alba added fuel to the ongoing moral and ethical debate over transgenic organisms. Nevertheless, R and Alba have exposed the lighter side of science and hopefully they add GloFish to greater acceptance of genetically modified organisms.

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R Fig. 1 Transgenic fluorescent organisms. A Glofish , expressing GFP, RFP, and GFP + RFP, marketed by Yorktown Technologies, L.P. B Eduardo Kac, GFP Bunny, 2000, transgenic artwork. Alba, the fluorescent rabbit. C Tobacco leaves expressing mGFP4. The top leaf is wild type and all others are expressing varying amounts of the protein

3 GFP in Transgenic Plants Plant scientists were excited by the success of GFP in other organisms. However, they were met with disappointment because wild-type GFP expression was found to be variable in plants. Wild-type GFP expression was first confirmed in plant cells and not in intact plant tissues. GFP fluorescence was observed in sweet orange [24] and maize (Zea mays) [25, 26] protoplasts. Transient expression in intact Arabidopsis thaliana (Arabidopsis) roots and leaves [26] was also observed but not in Arabidopsis protoplasts [25]. Additionally, stable transformation was confirmed in Arabidopsis but no fluorescence could be detected [27]. With limited success in GFP expression, researchers realized that substantial improvements needed to be made to the wild-type gene.

4 GFP Variants for Plant Expression Much of the credit for stable GFP expression in plants goes to Jim Haseloff. Haseloff et al. [28] reported that aberrant splicing of wild-type gfp mRNA occurred in plant

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cells due to a cryptic intron between nucleotides 380 and 463. In Arabidopsis, this 84 nucleotide deletion resulted in a truncated, non-fluorescing protein. Silent mutations were introduced into the splice recognition sites to remove the intron. Two promising variants were produced; mGFP4 and mGFP5 [29]. Expression of mGFP4 was observed in soybean (Glycine max) cells [30], Arabidopsis [28], tobacco [31], and other plants. However, mGFP4 did not exhibit stable fluorescence under field conditions even as the protein was expressed in the plant at levels that should have yielded visible green fluorescence [31–33] (Fig. 1c). In contrast, the mGFP5 variant with an endoplasmic reticulum (ER) targeting peptide (mGFP5-ER) [28, 29] showed improved levels of fluorescence, improved thermostability, and dual excitation in UV (395 nm) and blue light (473 nm) of almost equal amplitude. Subsequently, under field conditions mGFP5-ER was found to be expressed twice as much as mGFP4 with higher levels of fluorescence [33]. In plant biology, the S65T mutant contained the most significant chromophore modification to GFP. This change created a single blue excitation peak (489 nm optimum) and slight red shift excitation maxima from 507 to 511 nm [34]. Codon optimization was subsequently performed and up to a 100-fold increase in fluorescence was observed in plant cells [16]. There are two S65T variants that have been widely used in plant science. Haas et al. [35] created sGFP which is a synthetic S65T gene with the cryptic intron removed. The other widely used S65T GFP variant is the commercially available EGFP (Clontech). EGFP includes S65T, as well as the F64L and the Y145F mutations and is human codon-optimized [36]. Many GFP variants were made, but not all mutations improved expression levels or brightness. Researchers wanted colors other than green and shortly after the cloning of GFP, blue [4] and cyan [5] variants were produced. Until recently no GFP variant has been produced with an emissions maxima exceeding 529 nm. Mishin et al. [37] reported the first GFP mutant to form a red chromophore. However, only 1–3% of the protein produced matures to the red-emitting state.

5 Other Colors, Other Organisms With the lack of colors (emission) beyond 529 nm, researchers began searching for other marine organisms that possess GFP homologues. Mikhail Matz et al. [6] were able to clone red fluorescent proteins from non-bioluminescent Anthozoa species, most notably, DsRed [38], which has a number of commercially available variants (Clontech). The Anthozoa FPs have become widely utilized because they are spectrally diverse, ranging from blue to far red. Mutagenesis has been performed on these which has lead to stable, bright, and a few monomerized FPs; most of which have not been tested in plants [39, 40] (Fig. 2). Additionally, Matz and colleagues have recently discovered and characterized many new proteins from corals [12]. Table 1 contains a list of the most promising FPs available to researchers, but discussions on partially characterized FPs (those without published brightness information) are not included.


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Fig. 2 Examples of FPs expressed in plants. The images were taken with an epifluorescent stereoscope with 90 s exposure, 530/40 nm excitation light and 600/50 band-pass emission filter. The top leaf is wild type and the lower three are transgenic. A White light photo. B From left to right pporRFP, mOrange, and tdTomato

6 FP Toxicity and Allergenicity When a novel protein is introduced into an organism, toxicity is a concern and it has been suggested that GFP is cytotoxic to plant cells [27, 28, 41]. The argument was made that GFP fluorescence caused a photonic disturbance that created free radicals and eventual oxidative damage. However, many researchers failed to observe this toxicity in plants that were clearly expressing high levels of GFP [16, 32, 42, 43– 46], but see Liu et al. [47] who showed GFP toxicity to mammalian cells. To address the toxicity issue at the whole plant level, Harper et al. [33] tested tobacco in the field using three different GFP variants (mGFP4, mGFP5-ER, and sGFP). Over two growing seasons, seed yield and biomass were recorded. In this study, there was no cost to yield or biomass; therefore, it was concluded GFP is not toxic to plant cells. Many organisms have been transformed with FPs and these show no measurable host cost. It is well documented that plants have many characteristics to deal with excess light that could be damaging to cells.

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Table 1 Properties of potentially useful FPs in plants with published extinction coefficients and quantum yields. Many of these have not been tested in plants

Host Species

Excitation max. (nm) (Extinction coefficient)

Emission max. (nm) (% quantum yield)

Oligomerization (M = monomer, D = dimer, T = tetramer) Refs

CYAN AmCyan mmilCFP anobCFP2 MiCy

Anemonia majano Montipora millepora Acropora nobilis Acropara sp.

458 (40) 404 (90) 477 (86) 472 (27)

486 (24) 492 (43) 495 (28) 495 (90)

T Unknown Unknown D

[56] [12] [12] [71]

GREEN pplu GFP2 pmeaGFP2 pmeaGFP1 wtGFP mAG Azami Green (AG) ZsGreen pporGFP EGFP GFP Emerald GFP S65T cmFP512 plamGFP Kaede eechGFP3 GFP YFP Topaz GFP YFP Venus phiYFP

Pontellina plumata Pontella meadi Pontella meadi Aequorea victoria Galaxeidae sp. Galaxeidae sp. Azumi Zoanthus sp. Porites porites Aequorea victoria Aequorea victoria Aequorea victoria Cerianthus membranaceus Platygira lamellina Trachyphyllia geoffroyi Echinophyllia echinata Aequorea victoria Aequorea victoria Phialidium sp.

482 (70) 487 (98) 489 (99) 395 (27) 492 (72) 492 (72) 497 (36) 495 (54) 488 (56) 487 (58) 489 (55) 503 (59) 502 (96) 508 (99) 512 (45) 514 (94) 515 (92) 525 (115)

502 (60) 502 (72) 504 (74) 504 (79) 505 (81) 505 (67) 506 (63) 507 (98) 508 (60) 509 (68) 510 (64) 512 (66) 514 (99) 518 (80) 524 (120) 527 (60) 528 (57) 537 (60)

M Unknown Unknown M M T M Unknown M M M T Unknown T Unknown M M D

[72] [72] [72] [64] [67] [67] [56] [12] [64] [64] [64] [68] [12] [70] [12] [64] [65] [72]

RED mKO mOrange tdTomato DsRed amilRFP pporRFP mStrawberry mCherry eqFP611 t-HcRed1 mPlum

Fungia concinna Discosoma sp. Discosoma sp. Discosoma sp. Acropora millepora Porites porites Discosoma sp. Discosoma sp. Entacmaea quadricolor Heteractis crispa Discosoma sp.

548 (52) 548 (71) 554 (138) 558 (75) 560 (49) 578 (54) 574 (90) 587 (72) 559 (78) 590 (160) 590 (41)

559 (60) 562 (69) 581 (69) 583 (79) 593 (91) 595 (95) 596 (29) 610 (22) 611 (45) 637 (4) 649 (10)

M M D (tandem) T Unknown Unknown M M T T M

[71] [62] [62] [62] [12] [12] [62] [62] [69] [66] [63]

DUAL COLOR EosFP

Lobophyllia hemprichii

[73]


D

[73]

mEosFP


516 (70) 581 (55) 516 (66) 581 (60) 516 (64) 581 (66)

T

d2EosFP

506 (72) 571 (41) 506 (84) 569 (33) 505 (67) 569 (37)

M

[73]


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Additionally, when a novel protein is introduced into a plant that is intended for use as food or feed, human and animal health issues should be addressed. Before any crop expressing GFP could be deregulated, this issue would have to be probed extensively. However, there is evidence that GFP is neutral with regard to oral toxicity and allergenicity. Richards et al. [48] fed purified GFP and pelletized feed made from GFP expressing canola (Brassica napus) to rats. At physiological relevant levels GFP was completely digested and there were no allergenic features associated with the protein. However, when rats were fed amounts exceeding physiological levels (1 mg/day purified GFP) it altered the spectral properties of their feces – yielding green fluorescent poop (GFP). Aside from this novelty, the protein had no measurable effects on growth. These findings suggest that GFP and GFP-like proteins are likely safe with regards to oral toxicity and allergenicity.

7 FPs in Plant Research FPs have become integral tools in developmental biology and functional genomics. They have been fused to numerous proteins to monitor subcellular localization and to tag subcellular structures [49]. Included in a review by Mathur [49] is a useful list of targeted FP probes available for plants. Furthermore, in functional genomics research FPs have been used to assay promoter activity and to clone regulatory elements (as reviewed in Ayalew [50]). Several novel promoters have been characterized by expressed fluorescence. For example, the taro bacilliform virus promoter has been characterized by GFP fluorescence in banana and tobacco [51]. GFP has also been used to study RNA interference in plants. The studies were designed to examine patterns of gene silencing [52, 53].

8 Whole Plant FP Applications 8.1 Plant Zygosity Determination Using GFP as a Genetic Marker One difficulty when working with transgenic plants is transgene zygosity determination of dominant or semi-dominant traits. Using FPs, zygosity status can be determined in two ways. Halfhill et al. [54] demonstrated that heterozygous whole plant GFP fluorescence is approximately half the fluorescence of homozygous plants (Fig. 3a). This finding suggests that zygosity status can be determined by fluorescence alone. In two other studies, tobacco [55] and canola [56] plants expressed GFP under the control of the LAT59 pollen-specific promoter. This promoter was originally isolated from tomato and allows high GFP expression in pollen. The zygosity of these plants was determined based on the ratio of GFP-expressed to non-GFPexpressed pollen. T1 generation plants were grown and successfully categorized into homozygous (Fig. 3b), heterozygous (Fig. 3c), and isogenic plants for the transgene, according to the relative frequency of GFP-expressed pollen grains [56]. These data

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Fig. 3 Trangene zygosity determination. A Average GFP fluorescence at 508 nm for wild-type canola (Wt) and three GFP expressing transgenic lines (GT1, GT2, and GT3). Plant material was excited with 385 nm light. B Canola pollen from GFP homozygous individual. C Canola pollen from GFP heterozygous individual. Zygosity determination is based on the ratio of GFP-expressed pollen. Pictures were taken with blue excitation filter 360/30 nm, 510 long-pass emission filter, 1.54 s exposure time, at 200× magnification

indicate that FPs could be used for ecological monitoring of transgenic plants by analyzing hybridization and introgression status.

8.2 Monitoring Transgenic Organisms Over the past two decades, transgenic crops have increased in number and acreage grown worldwide. For this reason, concerns over environmental impacts have been voiced [57]. Since many crops are sympatric with their wild relatives, hybridization is sure to occur. It has been documented that transgene escape occurs via seed dispersal or through pollen movement with subsequent hybridization [58, 59]. Therefore, a transgenic monitoring system utilizing FPs to tag whole plants would be useful. In the past, ecological genetics focused on phenotypic traits or DNA markers. However, with FPs one would only have to look for a fluorescent signature. With


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this type of monitoring system, FPs can be genetically linked so that they are inherited along with transgenes in the event of gene flow. This would allow the tracking of transgene movement and their ecological effects, an important component in biotechnology risk assessment [60]. This approach has been demonstrated as an effective tool when either whole canola plants [58] or pollen alone [55, 56] were tagged with GFP . When whole plants were tagged, GFP was inherited along with the Bt insect resistance gene after transgenic canola was crossed with its wild relative Brassica rapa (field mustard) [58]. It has also been observed that GFP and Bt were still present together after several successive back-crosses to B. rapa [61]. When pollen was tagged with GFP, it was observed that transgenic pollen captured several meters away could be distinguished from non-transgenic pollen [55, 56]. It would be useful to track transgenic pollen in this manner because the information gathered could then be used in risk assessment. It is also important to note that FP-tagged transgenic plants can be monitored non-destructively, in real time [62], and perhaps remotely [63]. This type of monitoring could be extended to insects, domestic animals, and aquatic organisms.

8.3 Environmental Monitoring FPs could be used to monitor environmental conditions; in transgenic plants, FPs could be deployed in phytosensors. A phytosenor is a plant that has been genetically engineered to produce a detectable signal, such as an FP, under a certain condition, e.g., to report on a specific contaminant. Since plants are immobile, they possess biochemical and genetic mechanisms to respond to environmental stresses and xenobiotics. Once it has been determined how a particular plant responds to an environmental stress, the information obtained can then be used in the production of a phytosensor. For example, if a gene is expressed at higher levels under a specific condition, the gene’s promoter could be cloned and fused to an FP. This construct would then be introduced into a plant to be used as a bioreporter, producing an inducible fluorescence signal. There are many examples of potential phytosensors. In one study, GFP was fused to general stress promoters and introduced into Arabidopsis [64]. Plants were grown under normal conditions and subsequently subjected to low atmospheric pressure. Under low atmospheric pressure, GFP expression was induced contrasting with no GFP under ambient conditions [64]. To alert of herbivory damage, GFP was fused to a peroxidase promoter and introduced into tobacco. After caterpillar species were allowed to feed, GFP fluorescence was detectable [65]. In other studies, pathogen-inducible promoters were fused to GFP [66, 67] and introduced into plants. When these plants were exposed to fungal elicitors, gene expression was reported but at low levels. It is clear that real-world applications of phytosensors are possible; however, none have been deployed on a large scale. There is a real need for biosurveillance in the areas of precision farming systems, integrated pest management, soil fertility, and biosecurity. To make phytosensors a reality, additional research needs to be performed

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on plant stress responses and the appropriate FPs need to be determined for optimal expression.

9 Instrumentation and Methods for FP Detection and Quantification in Plants Observing FPs in whole plants is made possible by their unique fluorescence signatures. These plants can be monitored by simple visual observation or through the use of more sophisticated detection devices. The proper instrumentation is critical when analyzing FPs in transgenic organisms. Here we detail current instrumentation and methods for detection and quantification of FPs in plants.

9.1 Visual Detection With certain GFP variants, protein expression can be visualized with the naked eye by using a long-wave hand-held UV spot lamp (e.g., UVP model B 100 AP, UVP, Upland, CA, USA) (Fig. 4a). Many GFP variants, such as mGFP5, have dual excitation peaks: one at 395 nm (UV) and another at 470 nm (blue). The human eye cannot see long-wave UV; therefore, we can visualize GFP expression under UV

Fig. 4 Examples of current FP detection methods in plants. A Visual observation when excited with long-wave UV light (360 nm). B Lab-based detection with a FluoroMax scanning spectrometer. C Processed reading from the portable GFP-Meter. D Stand-off detection with laser-induced fluorescence spectrometry (LIFS) and laser-induced fluorescence imager (LIFI) instrumentation


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excitation. Most FPs, including many GFP variants, cannot be visualized in this manner because their excitation maxima are in the visible range. In these cases, visualization requires an emission filter to remove excitation light. An epifluorescent microscope, coupled with the proper filters, is often the best instrument for FP visualization.

9.2 Lab-Based FluoroMax-4 Spectrofluorometer The FluoroMax-4 (Jobin Yvon and Glen Spectra, Edison, NJ, USA) is a large labbased scanning spectrofluorometer used to detect and quantify GFP fluorescence. With this instrument, a 2 m bifurcated fiber optic cable is used to transmit excitation light and detect emission transmission to and from the sample [68]. For example, GFP can be measured by exciting a leaf sample with 395 nm UV (UV excitation maxima) light and scanning for emission transmission in the range of 440–600 nm. The collected fluorescence signal is displayed in units of photon counts per second (cps) (Fig. 4b). Standardization and normalization must be performed on each scan because each sample varies in background fluorescence [68]. The FluoroMax can be quantitative as well. In one study, a strong correlation was observed between GFP ELISA quantification and the FluoroMax readings [69]. This detection method will work for all FPs, including the ones with a small Stokes shift. There are excitation filters in place to block out bleed-over light and allow for emission scanning close to the excitation peak.

9.3 Portable Hand-Held GFP-Meter FP transgenic plants are often grown in field sites far away from the lab. To use a system like the FluoroMax for FP detection, tissue must be collected and brought back to the lab. However, there are field-portable detection systems available such as the GFP-Meter (Opti-Sciences, Tyngsboro, MA) fluorescent spectrophotometer [68]. The GFP-Meter is small, easy to use, operates on a 12 V battery, and has a data logging system. The principle behind the GFP-Meter is similar to the Fluoromax, except it is not a scanning spectrophotometer. Measurements are displayed as a single number in units of photon counts per second (cps) (Fig. 4c). The instrument has a filtered light emitting diode (LED) to generate excitation light. The excitation light travels to a fiber optic cable and through a band-pass filter until it reaches the sample. An attached leaf clip provides stability by holding the sample. The leaf clip also provides consistency between samples by holding the fiber optic cable at a fixed angle. The light emitted from the sample enters back into the fiber optic cable and through a band-pass filter. Subsequently, the measured fluorescence is shown in real time in a display window. The GFP-Meter comes equipped with a 465/35 nm band-pass excitation filter. Two emissions channels are available. Channel 1 is a GFP channel using a 530/35 nm band-pass filter and channel 2 is a chlorophyll channel using

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a 680/35 nm band-pass [68]. It is important to note that excitation and emission filters can be changed to meet the requirements of any FP. The GFP-Meter is quantitative as well. Regression analyses on GFP expression between the FluoroMax spectrofluorometer and GFP-Meter measurements produced strong positive correlations [68].

9.4 Stand-Off Laser-Induced Fluorescence Detection For FP detection from a distance, a laser-induced detection and imaging system has been developed. Laser-induced fluorescence spectrometry (LIFS) and laser-induced fluorescence imager (LIFI) were described and tested with transgenic canola and tobacco expressing GFP [63] (Fig. 4d). LIFS is a laser-based remote detection instrument that records fluorescence of transgenic organisms. The LIFS system collects fluorescence from a 10 cm diameter centered in the laser-illuminated area. A 3-m fiber optic bundle is a conduit for the light collected transferring to the input slit of a 275 cm focal length spectrograph (Model SP-275, Acton Research, Acton, MA, USA). A gated CCD camera (Princeton Instruments, Trenton, NJ, USA) allows detection of the transferred light at the output side of the spectrograph. LIFI is a remote sensing detection system used to capture images of fluorescent organisms. It uses a gated charge coupled device (CCD) camera system (NVSI Camera Systems, Fayetteville, NC, USA) as an imager to collect images. This camera is connected to an intensifier by a fiber optic taper. Fluorescent images of GFP expressing plants can be captured within the intensifier’s spectral bands. The bands can be extended from 400 to 900 nm and this emission range would capture the fluorescence of any FP. These two systems, LIFS and LIFI, have capabilities for remote detection of GFP expression in plants. Any FP could be used with this detection system as long as the excitation laser used meets the requirements of the FP. This capability makes these two systems good candidates for detection monitors of phytosensor systems.

10 Customized FPs With a variable color palette of FPs characterized, it is no longer novel to clone, mutagenize, and express FPs in transgenic organisms. However, new applications may drive researchers to mutagenize a particular FP to obtain desired spectral characteristics. For example, in one study, mGFP5 was used as a marker for stand-off detection of transgenic plants [63]. A pulsed ND:YAG (neodymium-doped yttrium aluminum garnet) laser with a tripled frequency to 355 nm was used to excite GFP. However, 355 nm has been found to increase the signal to noise ratio about ten times more than the optimal excitation which is 390 nm. Additionally, there is much more endogenous autofluorescence from leaves when excited with lower wavelengths (Fig. 5). When these factors are considered together, GFP fluorescence could be


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Fig. 5 Background autofluorescence in GFP transgenic and non-transgenic tobacco. Plant leaves were excited by either 395 or 525 nm light. Background fluorescence is much less when exciting with 525 nm compared with 395 nm. These data suggest that an FP with a 525 nm excitation and an emission in the range of 575–600 or 625–650 nm would be ideal in plants

masked and more difficult to detect. To decrease background fluorescence, a doubled frequency ND:YAG laser (532 nm) could be used for excitation. This laser would provide twice the power than excitation at 355 nm and it would significantly reduce the amount of background autofluorescence in plants (Fig. 5). However, this wavelength would not excite GFP. For these reasons, GFP is not the optimal FP for this particular application. If the detection system must be constrained, then it might be most cost-effective to tailor the FP to the laser characteristics. Perhaps an FP exists naturally that meets a priori requirements. Unfortunately, there is no wellcharacterized native or monomerized FP that meets these requirements (Table 1). Therefore, one will have to be tailored to the application, but laser characteristics are not the only constraints. Plant autofluorescence should be considered as well. There are spectral peaks that are located at 540 nm, which seem to be associated with general plant stress (Stewart et al., unpublished data), 610 and 680 nm, which correspond to chlorophyll fluorescence (Fig. 5). Therefore, a monomerized FP that is excitable at 532 nm and an emission that avoids plant autofluorescence peaks is desirable. There are a number of FPs listed in Table 1 that would be good candidates for random mutagenesis or directed evolution to acquire these characterizations for maximal detection in plant leaves using well-characterized systems.

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11 Conclusions The suite of FPs has dramatically changed scientific research in the past decade and a half. With the many FPs to choose from we can now do things that were not even a thought a few years ago. For example, “brainbow” is an elegant strategy that utilized multiple FPs to map the neural circuits of mice brains [70]. The pictures of the multicolored mice brains are as stunning as the research behind them. With such tools available to researchers, it is certain that the future will be colorful and bright. Acknowledgements We would like to thank Mary Rudis and Laura Abercrombie for generating Fig. 2 and its data. For Fig. 1, we would like to thank Eduardo Kac for allowing us to use “GFP Bunny, 2000, transgenic artwork. Alba, the fluorescent rabbit” reprinted in this chapter. We would also like to thank Christy Rose, Murali Rao, and Charles Poovaiah for their help and comments. This review was made possible by funding from the University of Tennessee and many agencies, including NSF, USDA, and the US Department of Defense.

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Peptide Foldamers: From Spectroscopic Studies to Applications Lorenzo Stella, Gianfranco Bocchinfuso, Emanuela Gatto, Claudia Mazzuca, Mariano Venanzi, Fernando Formaggio, Claudio Toniolo, Antonio Palleschi, and Basilio Pispisa

Abstract Peptide foldamers are synthetic oligopeptides which attain a few, specific, constrained conformations in solution. Here, we review our contributions to the study of the structural features of several foldamers, comprising Cα -tetrasubstituted aminoacids, by spectroscopic techniques and, in particular, by a combined approach employing time-resolved energy transfer (FRET) experiments and molecular modeling to determine interprobe distances and orientations. Our data show that, for rigid systems, the commonly used assumption of random orientation of donor and acceptor is unjustified, and that in these cases a correct evaluation of the orientation factor is mandatory for meaningful structural determinations. Finally, we illustrate some applications of peptide foldamers in studies on the kinetics of protein folding and on the realization of peptide-based molecular devices.

1 Introduction Nature performs a vast array of complex chemical tasks exploiting the capacity of biopolymers, most notably proteins, to attain a well-defined 3D structure. Synthetic polymeric systems which exhibit this essential property of natural biomolecules have been defined as “foldamers.” They represent optimal scaffolds to realize molecular devices with specific chemical functions, such as catalysis, molecular recognition, and light→energy conversion [1–3]. Even though some foldamers have been prepared from monomeric units which share no similarity with the peptide backbone of proteins, most foldamers try to mimic the chemical architecture of natural systems [2, 4, 5]. However, short peptides based on the 20 protein aminoacids are usually unordered in solution, due L. Stella (B) Università di Roma Tor Vergata, Dipartimento di Scienze e Tecnologie Chimiche, 00133 Rome, Italy e-mail: [email protected] C.D. Geddes (ed.), Reviews in Fluorescence 2008, Reviews in Fluorescence 2008, DOI 10.1007/978-1-4419-1260-2_17, C Springer Science+Business Media, LLC 2010

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Fig. 1 Ramachandran plot for a standard L-α-aminoacid (left panel) and for Aib (right panel)

to their relatively high conformational freedom, exemplified by the large accessible regions in the φ,ψ-space of the Ramachandran plot (Fig. 1a). This is why most research efforts concentrate on peptoids, β- or γ-peptides or nonstandard aminoacids [2, 6, 7]. Nature itself provides alternative building blocks which are much more conformationally restricted than protein aminoacids. Some peptides synthesized nonribosomially by diverse species of soil fungi (such as alamethicin) contain nonstandard residues, including Cα -tetrasubstituted α-aminoacids [8, 9]. Aib (α-aminoisobutyric acid or Cα -methylalanine) is the prototypical example of this class of aminoacids, which also includes Iva (isovaline), (αMe)Val (Cα -methylvaline), among many others (Fig. 2) [7, 10].

Fig. 2 Chemical structures of Aib and (αMe)Val

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Already in 1971, conformational energy calculations indicated that, due to steric interactions involving the gem-methyl groups linked to the α-carbon, the conformational space accessible to Aib is extremely limited and is located in the region of the Ramachandran plot corresponding to helical conformations (Fig. 1b) [11]. For this reason, Cα -tetrasubstituted aminoacids constitute suitable building blocks to synthesize peptide-based helical foldamers. In the last years we have employed spectroscopic techniques, particularly timeresolved fluorescence, for the conformational characterization in solution of several oligopeptides based on Cα -tetrasubstituted aminoacids [12–25]. Some of the peptides studied by us over the years are summarized in Fig. 3. This review describes the methodology developed by us, summarizing the main results (Section 2) and illustrating some of the most recent applications exploiting these conformationally constrained oligopeptides (Section 3).

Fig. 3 Primary structures of some of the peptides investigated and chemical structures of the chromophores (highlighted in red). Cα -Tetrasubstituted α-aminoacids are highlighted in green. All chiral α-aminoacids have the L-configuration, unless indicated otherwise

2 Spectroscopic Studies of Foldamer Structure in Solution 2.1 Secondary Structure In our studies of Aib- and (αMe)Val-based oligopeptides we have consistently found that these systems in solution adopt a very stable 310 -helical conformation,

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even if some of these molecules were formed by only a few residues. The 310 helix [3, 26] is a secondary structure with approximately 3 residues per turn, and H-bonds formed between the CO group of residue i and the NH group of residue i+3 (i+3→i H-bonds), so that 10 atoms are included in the pseudo-ring formed by the intramolecular H-bond (Fig. 4). By comparison, the canonical α-helix has 3.6 residues per turn, i+4→i H-bonds and 13 atoms per pseudo-ring (3.613 helix). The backbone dihedral angles for the 310 -helix fall in the same region of the Ramachandran plot of the α-helix, but, for protein aminoacids, the former secondary structure is slightly less stable than the latter, due to some close van der Waals contacts and a non-optimal H-bond geometry. However, 310 -helices are found rather frequently in the structures of proteins, most often as terminal extensions of an α-helix. The situation is different for peptides formed by Cα -tetrasubstituted α-aminoacids, because the particular steric characteristics of these residues tend to favor the 310 -helix. Evidence for the secondary structure adopted by the peptides investigated in solution came from IR absorption, 1 H-NMR, and circular dichroism (CD) experiments. IR absorption spectra (performed in deuterated methanol solutions) always demonstrated the formation of a stable secondary structure, as indicated by the high ratio between the intensities of the two bands corresponding to the stretching of NH bonds participating in, or free from, intramolecular H-bonds (around 3330 and 3430 cm−1 , respectively), as shown in Fig. 5. These results were even more informative when data from similar analogues of different length were compared. In this case it was possible to show that the intensity of the band corresponding to the “free” NH groups does not depend on the number of peptide bonds, while that of the band of H-bonded NH groups increased significantly [20]. This result indicates that all analogues contain the same number of free NH groups, corresponding to the N-terminal residues, which, in a helical structure

Fig. 4 Comparison between the 310 - (above) and α-helical (below) peptide structures. (A) View down the helix axis, showing the helical periodicity (approximately 3 residues per turn for the 310 helix, and 3.6 for the α-helix). (B) Side view, showing the pattern of H-bond formation (i+3→i for the 310 -helix, i+4→i for the α-helix). (C) Pseudo cycle formed by the intramolecular H-bond, comprising 10 atoms in the 310 -helix, and 13 in the α-helix. C atoms are shown in black, N atoms in blue, O atoms in red, and H atoms in white

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Fig. 5 Typical infrared absorption spectra of a series of peptide foldamers in the N-H stretching absorption region (see text). Data refer to the P-(Aib)n -N analogues and curves are numbered according to the number of Aib residues

cannot act as intramolecular H-bond donors. Therefore, the IR absorption data support the conclusion that all analogues attain a stable secondary structure. 1 HNMR experiments proved that this structure is a 310 -helix. The NH groups free from intramolecular H-bonds were determined by measuring the effect on the NH proton chemical shifts of a change in solvent composition. Our data showed that only the two N-terminal NH groups were not implicated in intramolecular H-bonds (Fig. 6), thus confirming the i+3→i scheme.

Fig. 6 Typical 1 H NMR titration of NH proton chemical shifts of Z-(Aib)6 -N in chloroform solution by addition of dimethylsulfoxide (DMSO). Protons and related curves are numbered from the N-terminus. Only the two N-terminal protons are significantly sensitive to the addition of DMSO, demonstrating that they are not involved in intramolecular H-bonds. This finding indicates an i+3→i H-bonding scheme (taking into account also the CO group of the N-terminal Z-protecting moiety)

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The 310 -helix presents a characteristic circular dichroism (CD) spectrum [27], but in our case this technique could not be used for the assignment of a specific secondary structure, because induced dichroism contributions from the fluorophores present in our analogues often perturbed the CD spectrum in the far-UV region. However, changes in temperature or solvent composition did not significantly affect the CD spectra analogues, confirming the high stability of the helical conformation of the peptide foldamers investigated.

2.2 More than just a Spectroscopic Ruler: FRET as a Structural Technique In order to obtain a detailed characterization of the structural features of the peptide foldamers in solution, we employed a synergistic approach which combines timeresolved fluorescence measurements of Förster resonance energy transfer (FRET) with molecular mechanics calculations. FRET is a well-known phenomenon of transfer of excitation energy from an excited donor fluorophore (D) to a ground state acceptor chromophore (A). The rate of this process depends on the spectral properties of the two chromophores, and, most importantly, on their relative distance and orientation [28]. In order to exploit FRET as a structural technique in our studies of peptide foldamers, we labeled the synthetic analogues with suitable fluorophores which could act as energy transfer donor and acceptors (Fig. 3). In the weak coupling regime, the FRET rate is given by kET =

1 τD0

R60 r6

3 2 k 2

(1)

where τD0 is the fluorescence lifetime of the donor (in the absence of acceptor), r is the distance between donor and acceptor, and R0 (Förster distance) is a parameter depending on the overlap between the donor emission and the acceptor absorption spectra [28]. Finally, the term that takes into account the orientation of the probes is defined as k = cos ϕDA − 3 cos θD cos θA

(2)

where ϕ DA is the angle between the acceptor absorption transition dipole and the donor emission transition dipole, while θ D and θ A are the angles formed by these dipoles with the line connecting the two probes. In Eq. (1), we have chosen a formulation slightly different from the one which is most widely used [28] to separate the orientational parameter k (which depends on the structural features of the sample) from R0 (which depends on spectroscopic variables). kET , or the closely related quenching efficiency E (i.e., the fraction of excited molecules which decay through the FRET phenomenon), can be easily measured

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with time-resolved fluorescence experiments, by comparing the lifetime of the donor in the presence of the acceptor (τDA ) with that of the unperturbed donor (τD0 ): 1 0 = kD + kET ; τDA 1 1 kET = A − 0 ; τD τD τA kET E= 0 = 1 − D0 kD + kET τD 1

τD0

0; = kD

(3)

If multiple conformations are present in a sample, with different interprobe distances and orientations, each of them has a different FRET rate and therefore a different lifetime. In this case a multiexponential decay is expected for the donor fluorescence: I(t) = I(t = 0)

αi · exp ( − t/τi )

(4)

i

If the dynamics between the different conformers is on a timescale longer than the fluorescence decay, then each lifetime τi provides information on the FRET rate of a specific conformer. In this case the pre-exponential factors αi correspond to the relative populations of the various conformers. The fluorescence decays of our systems could always be satisfactorily fitted with one, two, or in some cases three lifetime components (Fig. 7). Statistical analyses showed that a multiexponential model was generally superior to a lifetime

Fig. 7 Typical decay of the fluorescence of the donor in Fmoc-Toac-(αMe)Val(αMe)Val-NHtBu, exhibiting a distinctively bi-exponential decay

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distribution. Most importantly, changes in sample temperature or viscosity did not induce significant variations in the fluorescence decay, demonstrating the absence of any dynamic effect in the fluorescence timescale. These results, dramatically different from those usually obtained in FRET experiments of flexible oligopeptides composed of protein aminoacids [29], confirm the rigidity of our foldamers. Only a few conformers are significantly populated, and interconversion between them is slower than the nanosecond timeframe typical of fluorescence phenomena. These conclusions, however, do not take full advantage of all structural information contained in the FRET rates. As shown in Eq. (1), the FRET rate depends on two structural parameters (distance and orientation) that cannot be determined separately, unless some hypothesis is made on one of them. A common solution to this problem is to assume that donor and acceptor are rotating very rapidly on the fluorescence timescale and that their transition dipoles assume all possible relative orientations. In this case, k2 has the average value of 2/3, the FRET rate depends only on the interprobe distance, and the technique can be used as a “spectroscopic ruler” [28]. However, this is a rather stringent assumption, and if it is not realistic it introduces serious errors in the distance determination (the range of possible k2 values goes from 0 to 4), and hence in the derived structural features, making them unreliable or even meaningless. This problem is particularly relevant for rigid systems, such as those investigated by us, where our data indicate the absence of dynamic averaging effects. However, the small size and little conformational freedom of these peptides made them amenable to a complete conformational characterization by molecular mechanics (MM) calculations. This independent information was used, together with the FRET experimental data, to obtain both the distance and orientation parameters describing a molecular conformation. More specifically, the calculated molecular structures were used to predict the energy transfer rate corresponding to a given combination of interprobe distance and orientation (Eq. (1)). These theoretical rates were then compared with the experimentally determined ones. When agreement was found, the calculated conformers could be deemed reliable. This approach allowed us to determine the most relevant conformational features of the peptides investigated in solution.

2.3 Molecular Mechanics Calculations MM simulations were always performed with a homemade program, taking into account electrostatic, nonbonding, hydrogen-bond, and torsional interactions. Energy minimizations were carried out by starting from different secondary structures and relaxing the fixed geometry for all internal degrees of freedom, also including stretching and bending terms [23]. These calculations consistently showed that for the peptides investigated the 310 helical secondary structure is very stable (Fig. 8) and is characterized by a lower

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Fig. 8 Theoretical structure of analogue Ac-Toac-AibAib-Aib-Trp-Aib-OtBu in solution obtained by our method (left) compared to the crystallographic structure (right)

energy than the classical α-helix, in agreement with the spectral data. Our calculations also indicated that the rigidity of these systems is such that also the mobility of the fluorophores is significantly restricted and only a few conformers are highly populated in solution. This finding is once again in very good agreement with the experimental data.

2.4 Validation of the Computed Structures Several stringent quantitative comparisons could be performed between calculated structures and experimental data. First of all, for each theoretical conformer, the FRET rate and efficiency were calculated according to Eq. (1). These values can be compared with the experimentally determined parameters. Figure 9a reports the comparison between calculated and experimental efficiencies for several peptides studied by our group over the years. An excellent agreement is found only by taking into account the interprobe orientation, while it is completely lost if the commonly used assumption of random chromophore orientation (k2 =2/3) is used (empty symbols). This result highlights the importance of a correct evaluation of the orientation factor in FRET experiments on rigid systems and is particularly relevant in view of the fact that, even if the limitations of the k2 =2/3 assumption are well known, this approach is still widely used, often without a proper critical evaluation of its validity in the specific system under investigation. We think that an a priori assumption of average k2 values should be avoided, since it represents an immediate abdication to the possibility of discovering important structural details. It is worth recalling that several tests can be performed to assess the k2 = 2/3 assumption. Experimentally, fluorescence anisotropy measurements, determination of transfer efficiency as a

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Fig. 9 (A) Comparison between theoretical and experimental quenching efficiencies for several foldamers investigated by our group over the years (see Fig. 3). Full symbols refer to the theoretical efficiency calculated by taking into account both interprobe distance and orientation, while the empty symbols correspond to efficiencies calculated considering a random orientation of the two chromophores (k2 = 2/3). (B) Comparison between theoretical and experimental relative populations for the different conformers

function of temperature and viscosity, and exchange of the two probe positions provide a good insight into the problem. Theoretically, molecular mechanics and dynamics calculations can show the degree of disorder and mobility of the system under study. Where the results show that the assumption of an isotropic averaging of the probe orientation is untenable, the k2 =2/3 model should be discarded to avoid meaningless distance determinations, and the potential of a combined use of experiments and molecular modeling should be exploited to derive as much structural information as possible. Interestingly, following our studies on peptide

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foldamers, recent FRET investigations on nucleic acids [30], block copolymers [31], and proteins [32] have pointed out the same conclusions. A second comparison was performed between the experimental weight associated with each experimental lifetime component (pre-exponential factor α i in Eq. (4)) and the relative population of the different theoretical conformers, as calculated by the Boltzmann weights Pi : exp ( − Ui /RT) Pi = j exp ( − Uj /RT)

(5)

where Ui is the theoretical energy of the ith conformer. Figure 9b shows this comparison for the peptides investigated by our group. Also in this case the agreement is satisfactory, even if the results are somewhat more scattered than in Fig. 9a because of the larger errors in the measured values and, possibly, of the empirical nature of the force field employed. Finally, when X-ray diffraction structures became available for some of our peptides [14, 33], an excellent agreement with our theoretical models was always found (Fig. 8). Since the solution and crystal-state conformations of short peptides comprising only protein aminoacids often differ markedly because of their high flexibility, this result is an indication of both the quality of our results and the rigidity of peptide foldamers. In conclusion, by combining time-resolved FRET measurements and other spectroscopic data with molecular mechanics calculations we were able to identify the most relevant structural features of the peptides investigated in solution. The overall findings provide great confidence in the proposed structural models and indicate that these peptides are indeed foldamers, because they fold in solution in a stable and rigid 310 -helical secondary structure. A variety of other experimental data on similar peptides agree with this conclusion [10]. Therefore, peptide foldamers based on Cα -tetrasubstituted α-aminoacids can be confidently employed in all applications requiring rigid helical segments, as demonstrated by the selected examples presented in the next section.

3 Selected Applications 3.1 Peptide Foldamers as Models of Protein Folding One of the most promising applications of peptide foldamers is the realization of model systems of protein folding. The discovery of several fast-folding proteins, which can attain their native conformation in just a few microseconds, has generated a renewed interest for kinetic studies of fast events in the folding of proteins and model compounds [34]. These investigations can shed some light on the folding process and discriminate between different protein folding theories [35]. Studies on model systems suggested that contact formation between two points in an unordered

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peptide by random diffusional motions takes place in a few nanoseconds [36]. However, little is known about the timescale for the collisional encounter between two helical segments, a fundamental step in the protein folding process, which is of particular relevance because most of the newly discovered fast-folders are helical bundles [34]. Trichogin GA IV (nOct-Aib-Gly-Leu-Aib-Gly-Gly-Leu-Aib-Gly-Ile-Lol, in which nOct is n-octanoyl and Lol is leucinol) is an antimicrobial peptide member of the lipopeptaibol family, a group of membrane-active compounds of fungal origin, characterized by a high content of Aib residues [37–39]. X-ray diffraction studies on trichogin GA IV [40] show that the first few residues at the N-terminus adopt a righthanded 310 -helix, while the rest of the peptide chain forms a distorted, right-handed α-helix (Fig. 10). A NMR investigation indicates that this mixed 310 /α-helical structure is basically maintained in solution [41, 42]. The hypothesis that the flexible -Gly5-Gly6- motif gives rise to a hinge point between the two helical segments has also been advanced [41]. Therefore, trichogin GA IV could represent an interesting model for studying the coalescence of a pair of helical segments to form a helixturn-helix motif. We applied time-resolved optical spectroscopy techniques, from the nanosecond to the microsecond timescale, on an analogue of trichogin GA IV with the sequence Fmoc-Aib-Gly-Leu-Aib-Gly-Gly-Leu-TOAC-Gly-Ile-Leu-OMe denoted F0T8 [43, 44]. F0T8 differs from trichogin GA IV by the substitution of the n-octanoyl lipid chain at the N-terminus by Fmoc (fluoren-9-ylmethyloxycarbonyl), a well-known fluorescent group, the replacement of Aib8 by TOAC (4-amino-1oxyl-2,2,6,6-tetramethylpiperidine-4-carboxylic acid), a fluorescence quencher, and the substitution of the 1,2-aminoalcohol Lol moiety group at the C-terminus by LeuOMe. Photophysical quenching efficiencies were determined with respect to the reference compound Fmoc-Aib-Gly-Leu-Aib-Gly-Gly-Leu-Aib-Gly-Ile-Leu-OMe (denoted as F0), in which the TOAC quencher is missing. CD spectra prove that these analogues exhibit the same conformational properties of the natural peptide. As we pointed out in the previous section, peptide foldamers comprising exclusively Cα -tetrasubstituted α-aminoacids exhibited a fluorescence decay which could always be described by a few discrete exponential components. By contrast, the fluorescence time decay of F0T8 in methanol was accounted for by two relatively broad continuous lifetime distributions (Fig. 11), indicating a higher degree of conformational flexibility, as expected due to the presence of several Gly residues. The two distributions were centered at τ 1 =1.7 ns (relative weight=0.15) and at τ 2 = 5.7 ns

Fig. 10 Crystallographic structure of trichogin GA IV. Aib residues are shown in green, Gly residues in red, hydrophobic residues in blue, and the N-terminal n-octanoyl group in light blue

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Fig. 11 Lifetime distributions for the fluorescence decay of F0T8

(relative weight=0.85), and the corresponding FRET efficiencies were E1 =0.81 and E2 =0.37, respectively (the lifetime of reference compound F0 was 9.0 ns). Interestingly, changes in the solvent composition, viscosity, or temperature (from −4ºC to 40ºC, in glycerol/methanol 8:2, v/v) did not appreciably perturb the fluorescence decay, indicating that no dynamic effect is present in the ns time frame. According to these data, therefore, trichogin populates two families of conformers, characterized by a different distance between fluorophore and quencher, which interconvert on a timescale slower than ns. MM calculations, performed starting from the X-ray diffraction peptide structure, indicate that the E2 quenching efficiency very likely corresponds to a helical conformation. Indeed, the lowest energy conformer has the first three to four residues at the N-terminus in a 310 -helix arrangement, while the rest of the peptide chain forms a distorted α-helix (Fig. 12a). The computed quenching efficiency for this conformation is 0.36, in very good agreement with the experimental value (E2 = 0.37). However, no linear helical conformer was able to reproduce the other quenching efficiency (E1 = 0.81) associated with the shorter lifetime distribution. Blocking the peptide backbone in a rigid, elongated helical conformation and exploring by MM calculations all possible side chain conformations, we found that the minimal interprobe center-to-center distance is 13.4 Å. By using this value and considering the appropriate value of R0 (10.9 Å), the highest experimental quenching efficiency (E1 = 0.81) can be obtained only with k2 = 9.8. This result is obviously untenable, because by its geometrical definition k2 can vary between 0 and 4 only. These results strongly suggested that folded structures, characterized by relatively short throughspace Fmoc/TOAC distances, in which the flexible, central Gly5-Gly6 motif should play a pivotal role, significantly populate the solution. We, therefore, repeated the energy minimization by starting from all possible backbone rotamers for the two central Gly residues. The lowest energy conformers obtained in this way exhibited a

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Fig. 12 Molecular models for the low-energy conformers of F0T8

bent structure, characterized by a compact 3D arrangement, including an N-terminal 310 -helix and a C-terminal distorted α-helix, connected by a turn involving the -Gly5-Gly6- residues (Fig. 12b). The interprobe distance and orientation of these conformers is fully compatible with the higher quenching efficiency experimentally observed (E1 = 0.81). These results suggest that the two families of conformers associated with the two lifetime distributions correspond to an extended helical structure and to a compact helix-turn-helix conformation in which quenching of the Fmoc singlet state is more efficient. No interconversion between these two conformational families takes place in the ns timescale. To extend our investigation to the microsecond timescale we exploited the relatively efficient intersystem crossing between the Fmoc excited singlet and triplet states. Excitation of Fmoc leads to a significant population of the triplet state, which has a lifetime in the microsecond time frame, and can be detected by time-resolved transient-absorption measurements on the Fmoc triplet state. Interestingly, TOAC is an efficient quencher also of the triplet state, as demonstrated by intermolecular quenching experiments performed with Fmoc and TOAC model compounds in methanol. These measurements yielded a bimolecular quenching constant of 1.3 · 109 M−1 s−1 , a value close to the diffusion limit, indicating that, once the two probes attain short separation distances, Fmoc triplet quenching by TOAC occurs almost instantaneously. This finding allowed us to use intramolecular tripletquenching experiments to determine the rate of contact formation between the two probes in F0T8. The observed triplet-decay rate can be written as kobs = k0 +kuni , where k0 is the rate in the absence of quencher and kuni is the unimolecular rate associated with the intramolecular quenching processes. According to the intermolecular quenching results, we could safely assume that kuni is determined by the rate of the process bringing the D/A pair to close separation distances. In our case, this process was represented by the interconversion from the extended helical conformation

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to the helix-turn-helix arrangement. Transient-absorption data yielded a value of kuni = 3.1 · 106 s−1 in MeOH. This value (kuni −1 =320 ns) is in between the diffusionally controlled rate of formation of end-to-end contacts in randomly coiled peptides (20–100 ns) and the 1–10 μs needed for complete folding of an all-helical fast-folding protein. Interestingly, in glycerol/MeOH (8:2, v/v) the triplet-quenching efficiency decreased (kuni = 1.3 · 106 s−1 ), confirming that this process is governed by dynamic events related to the interconversion from the extended helical conformation to the helix-turn-helix arrangement. In conclusion, quenching experiments of the Fmoc singlet and triplet excited states enabled us to study peptide dynamics from the nanoseconds to the microseconds and to determine the characteristic time of the diffusional encounter of two helical segments connected by a flexible peptide segment (320 ns). This phenomenon might actually mimic the early hydrophobic-collapse step of protein folding.

3.2 Peptide Foldamers as Building Blocks for Molecular Devices Another important area of application of peptide foldamers currently investigated by our group is the use of these rigid molecules as building blocks for ordered complex architectures, most notably self-assembled monolayers (SAMs) [45–47]. SAMs of alkanethiols on gold substrates have been extensively studied for potential applications in the areas of biosensing, molecular recognition, optoelectronics, nanopatterning, and soft lithography [48]. Much less attention has been devoted to SAMs based on peptide foldamers, but we argue that these systems deserve further attention, for several reasons: (i) peptides can be easily functionalized; (ii) helical peptides exhibit an intrinsic tendency to self-assemble, as shown by the architecture of several proteins; (iii) the peptide bond is polar (3.6 D), and all these dipoles are aligned when the backbone adopts a helical conformation. The macrodipole generated in this way favors directional electron transfer, and therefore helical peptides represent a useful scaffold for biomolecular electronic devices; (iv) SAMs formed by oligopeptides with a specific conformation have obvious applications in the realization of biosensing devices. Peptide SAMs can be obtained by exploiting the great affinity of sulfur atoms for gold surfaces. For instance, we grafted peptide foldamers to gold by attaching a lipoyl group, bearing a disulfide functionality, to the N-terminus of a fluorescent hexapeptide: Lipoyl-(Aib)4 -Trp-Aib-OtBu(SSA4WA) Simple incubation of a gold substrate with a 1 mM ethanolic peptide solution led to the formation of a densely packed peptide monolayer on the surface, as demonstrated by quartz crystal microbalance, scanning tunneling microscopy (STM), cyclic voltammetry (CV), and X-ray photoelectron spectroscopy experiments. For

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Fig. 13 Typical STM image of the SSA4WA peptide monolayer on gold. Image size: 100 × 100 nm. The depth of the observed holes (about 2 Å) indicates a complete coverage of the surface by the peptide, as shown in the model of a hole reported in the lower panel

instance, one STM image of the peptide monolayer on an atomically flat gold surface is shown in Fig. 13. The height of the observed holes and steps corresponds to a single Au atomic step, indicating a complete coverage of the surface. In addition, CV plots associated with the discharge of the Fe3+ /Fe2+ pair on a bare gold electrode and on gold electrodes coated with SSA4WA (Fig. 14) show a substantial decrease of the electrochemical activity due to the formation of a well-packed peptide monolayer. To illustrate the potentialities of a peptide monolayer for the construction of functional biomolecular electronic devices, we demonstrated that irradiation of the SSA4WA monolayer in the Trp near-UV absorption region leads to the generation of a significant photocurrent. Typical photocurrent intensities of the peptide-modified gold electrode, immersed in a solution of the electron donor triethylethanolamine (TEOA), upon irradiation at 280 nm, are reported in Fig. 15. The measured photocurrent intensity follows reversibly the on/off photoexcitation cycles without a

Fig. 14 Cyclic voltammetry curves associated with the discharge of the Fe3+ /Fe2+ pair on a bare gold electrode (curve A) and on gold electrodes coated with SSA4WA (curve B)

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Fig. 15 Photocurrent curves corresponding to excitation at 280 nm in a TEOA solution for a SSA4WA monolayer on a gold electrode

significant degradation of the anodic current. This current is due to an electron transfer process between the Trp excited state and the gold electrode (Trp∗→Au), followed by an electron transfer from TEOA to the Trp+· radical cation, which gives rise to a net anodic current. These results emphasize the good electron-mediating properties of a helical peptide spacer, suggesting the possibility of developing bio-hybrid molecular devices based on the integration of photoactive peptide layers chemisorbed on conductive supports.

4 Conclusions Our studies contributed to establish the high stability of 310 -helical structures formed by oligopeptides based on Cα -tetrasubstituted α-aminoacids. The road is now open to applications of these rigid, conformationally constrained, systems. Examples of possible applications of peptide foldamers in the realization of molecular devices for electron transport and photocurrent generation and in the design of small systems mimicking the basic elements of protein folding were described in this review. Peptide foldamers are already being exploited in many other applications, such as in the realization of artificial membrane channels, in the design of mini-enzymes, and in biomolecular recognition processes where a helical structure is a key requirement [4, 5, 49].

References 1. Gellman, S.H. (1998) Foldamers: a manifesto. Acc. Chem. Res. 31, 173–180. 2. Cheng, R.P. (2004) Beyond de novo protein design – de novo design of non-natural folded oligomers. Curr. Opin. Struct. Biol. 14, 512–520. 3. Crisma, M., Formaggio, F., Moretto, A. and Toniolo, C. (2006) Peptide helices based on α-amino acids. Biopolymers 84, 3–12.

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4. Cubberley, M.S. and Iverson, B.L. (2001) Models of higher-order structure: foldamers and beyond. Curr. Opin. Chem. Biol. 5, 650–653. 5. Licini, G., Prins, L. and Scrimin, P. (2005) Oligopeptide foldamers: from structure to function. Eur. J. Org. Chem. 969–977. 6. Sanford, A.R., Yamato, K., Yang, X., Yuan, L., Han, Y. and Gong, B. (2004) Well-defined secondary structures. Information-storing molecular duplexes and helical foldamers based on unnatural peptide backbones. Eur. J. Biochem. 271, 1416–1425. 7. Toniolo, C., Crisma, M., Formaggio, F., Peggion, C., Broxtermann, Q.B. and Kaptein, B. (2004) Molecular spacers for physicochemical investigations based on novel helical and extended peptide structures. Biopolymers 76, 162–176. 8. Kleinkauf, H. and von Döhren, H. (1990) Nonribosomal biosynthesis of peptide antibiotics. Eur. J. Biochem. 192, 1–15. 9. Aravinda, S., Shamala, N., Roy, R.S. and Balaram, P. (2003) Non-protein amino acids in peptide design. Proc. Indian Acad. Sci. (Chem. Sci.) 115, 373–400. 10. Toniolo, C., Crisma, M., Formaggio, F. and Peggion, C. (2001) Control of peptide conformation by the Thorpe-Ingold effect (Cα -tetrasubstitution). Biopolymers 60, 396–419. 11. Marshall, G.R. (1971) Studies on the biologically active conformations of angiotensin. In N. Kharasch (Ed.), Intra-Science Chemistry Report. Gordon and Breach, New York, pp. 305–316. 12. Pispisa, B., Palleschi, A., Venanzi, M. and Zanotti. G. (1996) Conformational statistics and energetic analysis of sequential peptides undergoing intramolecular transfer of excitation energy. J. Phys. Chem. B 100, 6835–6844. 13. Pispisa, B., Palleschi, A., Stella, L., Venanzi, M. and Toniolo, C. (1998) A nitroxide derivative as a probe for conformational studies of short linear peptides in solution. A spectroscopic and molecular mechanics investigation. J. Phys. Chem. B 102, 7890–7898. 14. Toniolo, C., Formaggio, F., Crisma, M., Mazaleyrat, J.P., Wakselman, M., George, C., Deschamps, J.R., Flippen-Anderson, J.L., Pispisa, B., Venanzi, M. and Palleschi, A. (1999) First peptide-based system of rigid donor – rigid interchromophore spacer – rigid acceptor: a structural and photophysical study. Chem. Eur. J. 5, 2254–2264. 15. Pispisa, B., Mazzuca, C., Palleschi, A., Stella, L., Venanzi, M., Formaggio, F., Polese, A. and Toniolo, C. (2000) Structural features of linear, (αMe)Val-based peptides in solution by photophysical and theoretical conformational studies. Biopolymers 55, 425–435. 16. Pispisa, B., Palleschi, A., Stella, L., Venanzi, M., Formaggio, F., Polese, A. and Toniolo, C. (2000) Structural features of linear, homo-Aib based peptides in solution. A spectroscopic and molecular mechanics investigation. J. Pept. Res. 56, 298–306. 17. Pispisa, B., Stella, L., Venanzi, M., Palleschi, A., Viappiani, C., Polese, A., Formaggio, F. and Toniolo, C. (2000) Quenching mechanisms in bichromophoric, 310 -helical Aib-based peptides, modulated by chain length-dependent topologies. Macromolecules 33, 906–915. 18. Pispisa, B., Stella, L., Venanzi, M., Palleschi, A., Marchiori, F., Polese, A. and Toniolo, C. (2000) A spectroscopic and molecular mechanics investigation on a series of Aib-based linear peptides and a peptide template, both containing tryptophan and a nitroxide derivative as probes. Biopolymers, 53, 169–181. 19. Stella, L. (2001) Comparisons between time-resolved fluorescence experiments and computer simulations. In G. M. Giacometti and G. Giacometti (Eds.) Spectroscopic Techniques in Biophysics. IOS Press, Amsterdam, 2001, pp. 89–103. 20. Pispisa, B., Palleschi, A., Mazzuca, C., Stella, L., Valeri, A., Venanzi, M., Formaggio, F., Toniolo, C. and Broxterman, Q.B. (2002) The versatility of combining FRET measurements and molecular mechanics results for determining the structural features of ordered peptides in solution. J. Fluoresc. 12, 213–217. 21. Pispisa, B., Mazzuca, C., Palleschi, A., Stella, L., Venanzi, M., Formaggio, F., Toniolo, C. and Broxterman, Q.B. (2002) Structural features and conformational equilibria of 310 -helical peptides in solution by spectroscopic and molecular mechanics studies. Biopolymers 67, 247–250.

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22. Pispisa, B., Palleschi, A., Stella, L., Venanzi, M., Formaggio, F., Toniolo, C. and Broxterman, Q.B (2002) Effects of helical distortions on the optical properties of amide NH infrared absorption in short peptides in solution. J. Phys. Chem. B 106, 5733–5738. 23. Pispisa, B., Mazzuca, C., Palleschi, A., Stella, L., Venanzi, M., Formaggio, F., Toniolo, C., Mazaleyrat, J.P. and Wakselman, M. (2003) Spectroscopic properties and conformational features of short linear peptides in solution. A fluorescence and molecular mechanics investigation. J. Fluoresc. 13, 139–147. 24. Pispisa, B., Mazzuca, C., Palleschi, A., Stella, L., Venanzi, M., Wakselman, M., Mazaleyrat, J.P., Rainaldi, M., Formaggio, F. and Toniolo, C. (2003) A combined spectroscopic and theoretical study of a series of conformationally restricted hexapeptides carrying a rigid binaphthyl-nitroxide donor-acceptor pair. Chem. Eur. J. 9, 2–11. 25. Venanzi, M., Valeri, A., Palleschi, A., Stella, L., Moroder, L., Formaggio, F., Toniolo, C. and Pispisa, B. (2004) Structural properties and photophysical behavior of conformationally constrained hexapeptides functionalized with a new fluorescent analog of tryptophan and a nitroxide radical quencher. Biopolymers 75, 128–139. 26. Toniolo, C. and Benedetti, E. (1991) The polypeptide 310 -helix. Trends Biochem. Sci. 16, 350–353. 27. Toniolo, C., Formaggio, F., Tognon, S., Broxterman, Q.B., Kaptein, B., Huang, R., Setnicka, V., Keiderling, T.A., McColl, I.H., Hecht, L. and Barron, L.D. (2004) The complete chirospectroscopic signature of the peptide 310 -helix in aqueous solution. Biopolymers 75, 32–45. 28. Andrews, D.L. and Demidov A.A. (1999) Resonance Energy Transfer. Wiley, Chichester. 29. Lakowicz, J.R. (1999) Principles of Fluorescence Spectroscopy. Kluwer, New York. 30. Lewis, F.D., Zhang, L. and Zuo, X. (2005) Orientation control of fluorescence resonance energy transfer using DNA as a helical scaffold. J. Am. Chem. Soc. 127, 10002–10003. 31. Yang, J. and Winnik, M.A. (2005) The orientation parameter for energy transfer in restricted geometries including block copolymer interfaces: a Monte Carlo study. J. Phys. Chem. B 109, 18408–18417. 32. Zeng, W., Seward, H.E., Málnási-Csizmadia, A., Wakelin, S., Woolley, R.J., Cheema, G.S., Basran, J., Patel, T.R., Rowe, A.J. and Bagshaw, C.R. (2006) Resonance energy transfer between green fluorescent protein variants: complexities revealed with myosin fusion proteins. Biochemistry 45, 10482–10491. 33. Crisma, M., Deschamps, J.R., George, C., Flippen-Anderson, J.L., Kaptein, B., Boxterman, Q.B., Moretto, A., Oancea, S., Jost, M., Formaggio, F. and Toniolo, C. (2005) A topographic and conformationally constrained, spin-labeled, α-amino acid: a crystallographic characterization in peptides. J. Pept. Res. 65, 564–579. 34. Kubelka, J., Hofrichter, J. and Eaton, W.A. (2004) The protein folding “speed limit”. Curr. Opin. Struct. Biol. 14, 76–88. 35. Snow, C.D., Nguyen, H., Pande, V.S. and Gruebele, M. (2002) Absolute comparison of simulated and experimental protein-folding dynamics. Nature 420, 102–106. 36. Eaton, W.A., Muñoz, V., Hagen, S.J., Jas, G.S., Lapidus, L.J., Henry E.R. and Hofrichter, J. (2000) Fast kinetics and mechanisms in protein folding. Annu. Rev. Biophys. Biomol. Struct. 29, 327–359. 37. Stella, L., Mazzuca, C., Venanzi, M., Palleschi, A., Didonè, M., Formaggio, F., Toniolo, C. and Pispisa, B. (2004). Aggregation and water-membrane partition as major determinants of the activity of the antibiotic peptide trichogin GA IV. Biophys. J. 86, 936–945. 38. Mazzuca, C., Stella, L., Venanzi, M., Formaggio, F., Toniolo, C. and Pispisa, B. (2005) Mechanism of membrane activity of the antibiotic trichogin GA IV: a two-state transition controlled by peptide concentration. Biophys. J. 88, 3411–3421. 39. Gatto, E., Mazzuca, C., Stella, L., Venanzi, M., Toniolo, C. and Pispisa, B. (2006) Effect of peptide lipidation on membrane perturbing activity: a comparative study on two trichogin analogues. J. Phys. Chem. B. 110, 22813–22818.

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40. Toniolo, C., Peggion, C., Crisma, M., Formaggio, F., Shui, X. and Eggleston, D.S. (1994) Structure determination of racemic trichogin A IV using centrosymmetric crystals. Nat. Struct. Biol. 1, 908–914. 41. Auvin-Guette, C., Rebuffat, S., Prigent, Y. and Bodo, B. (1992) Trichogin A IV, an 11-residue lipopeptaibol from Trichoderma longibrachiatum. J. Am. Chem. Soc. 114, 2170–2174. 42. Monaco, V., Locardi, E., Formaggio, F., Crisma, M., Mammi, S., Peggion, E., Toniolo, C., Rebuffat, S. and Bodo, B. (1998). Solution conformational analysis of amphiphilic helical, synthetic analogs of the lipopeptaibol trichogin GA IV. J. Pept. Res. 52, 261–272. 43. Venanzi, M., Gatto, E., Bocchinfuso, G., Palleschi, A., Stella, L., Formaggio, F. and Toniolo, C. (2006) Dynamics of formation of a helix-turn-helix structure in a membrane-active peptide: a time-resolved spectroscopic study. ChemBioChem 7, 43–45. 44. Venanzi, M., Gatto, E., Bocchinfuso, G., Palleschi, A., Stella, L., Baldini, C., Formaggio, F. and Toniolo, C. (2006) Peptide folding dynamics: a time-resolved study from the nanosecond to the microsecond time regime. J. Phys. Chem. B 110, 22834–22841. 45. Pace, G., Venanzi, M., Castrucci, P., Scarselli, M., De Crescenzi, M., Palleschi, A., Stella, L., Formaggio, F., Toniolo, C. and Marletta, G. (2006) Static and dynamic features of a helical hexapeptide chemisorbed on a gold surface. Mater. Sci. Eng. C. 27, 1309–1312. 46. Venanzi, M., Pace, G., Palleschi, A., Stella, L., Castrucci, P., Scarselli, M., De Crescenzi, M., Formaggio, F., Toniolo, C. and Marletta, G. (2006) Densely-packed self-assembled monolayers on gold surfaces from a conformationally constrained helical hexapeptide. Surf. Sci. 600, 409–416. 47. Gatto, E., Venanzi, M., Palleschi, A., Stella, L., Pispisa, B., Lorenzelli, L., Toniolo, C., Formaggio, F. and Marletta, G. (2006) Self-assembled peptide monolayers on interdigitated gold microelectrodes. Mater. Sci. Eng. C 27, 1309–1312. 48. Love, J.C., Estroff, L.A., Kriebel, J.K., Nuzzo, R.G. and Whitesides, G.M. (2005) Selfassembled monolayers of thiolates on metals as a form of nanotechnology. Chem. Rev. 105, 1103–1170. 49. Sia, S.K., Carr, P.A., Cochran, A.G., Malashkevich, V.N. and Kim, P.S. (2002) Short constrained peptides that inhibit HIV-1 entry. Proc. Natl. Acad. Sci. USA 99, 14664–14669.

Circularly Polarized Luminescence (CPL) of Proteins and Protein Complexes Eugene Gussakovsky

Abstract The review considers general theory of circularly polarized luminescence (CPL), instrumentation employed for its accurate measurement including artifacts, calibration, principle parts, and its application to proteins and protein complexes. Circularly polarized intrinsic fluorescence of proteins includes CPL of peptides, poly-α-amino acids, tyrosine and tryptophan residues in proteins, complexes of proteins with functional non-fluorescent agents, and protein conformation perturbation. CPL of fluorescent agents in protein complexes addresses to artificial probes, enzymatic cofactors, bilirubin, lanthanides and light-harvesting chlorophyll-protein complex of photosynthetic photosystem II of higher plants. Abbreviations Ala ANS CD CPL DMSO Eu(facam)3 Glu Gly HSA LHCII NAD NADH PLP poly(NapAla) PEM

alanine 8-anilino-1-naphthalenesulfonic acid circular dichroism circularly polarized luminescence dimethyl sulfoxide tris(3-trifluoroacetyl-D-camphorato)europium(III) glutamic acid glycine human serum albumin light-harvesting complex of Photosystem II nicotinamide dinucleotide reduced nicotinamide adenine dinucleotide pyridoxal 5 -phosphate poly(L-naphthylalanine) photoelastic modulator

E. Gussakovsky (B) National Research Council Canada, Institute for Biodiagnostics, Winnipeg, Manitoba R3B 1Y6, Canada


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TNS Trp Tyr εNAD+

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2-p-toluidinylnaphthalene-6-sulfonate tryptophan tyrosine 1,N6 -ethenoadenine dinucleotide

1 Introduction: What Is CPL? Light absorption and emission correspond to the transitions between ground electronic state S0 and first singlet excited state S1 , S0 → S1 and S1 → S0 , respectively. According to the Franck–Condon principle, the absorption depends on the conformation of the ground electronic state, whereas emission depends on the conformation of the excited state. Circular dichroism (CD) is a difference in absorbance of left- (l) and right-handed (r) circularly polarized light ε = εl – εr and serves as a measure of chirality of the ground electronic state conformation. Respectively, if the excited electronic state conformation is also chiral, then emission should be circularly polarized. The difference in light emission f = fl – fr is a circularly polarized luminescence (CPL). It is obvious that as any electronic transition emission, CPL does not depend on the way by which the chromophore appeared in the excited state. For example, circularly polarized chemiluminescence and bioluminescence were reported during 1977–1980 [1, 2]. Electroluminescence of light emitting diodes may also be circularly polarized [3]. Respectively, if the excitation of photoluminescence is not polarized, then emission becomes circularly polarized because of the chirality of the molecule in its excited electronic state. Quantitatively CPL is expressed by the emission anisotropy factor, gem , which is a weighed difference between the left- and the right-handed circularly polarized components of the total emitted light: fl − fr 2f = f fl + fr /2

gem (λ) =

(1)

where f and f are the total luminescence intensity and the number proportional to its circularly polarized part, respectively. Absorption anisotropy factor has been defined as follows: gab (λ) =

εl − ε r ε = ε (εl + εr ) 2

(2)

where ε and ε are extinction coefficient and CD, respectively, and gab characterizes the weighed circular dichroism. The comparison of the emission and absorption anisotropy factors indicates that CD and CPL methods are in fact complementary.

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Electrodynamical presentation of anisotropy factors (1) and (2) [4]: g

gab (isotropic) = 4 gem (isotropic) = 4

g

Im P( Q0 ) · M( Q0 ) g

[P( Q0 )]2 Im P( Qk0 ) · M( Qk0 ) [P( Qk0 )]2

(3) (4)

where P and M are electric and magnetic transition dipole moments, respectively, g and Q0 and Qk0 are coordinates of the ground-state (g) and excited-state (k) equilibrium conformations, respectively, that illustrate how gam depends on the chirality in the ground state and gem depends on the chirality in the excited state. It clearly shows that the difference in molecular geometry in the ground and excited electronic g states leads to the difference in absorption and emission anisotropy factors (Q0 = Qk0 ⇒ gab = gem ). If fluorescence consists of a single electron transition, then gem is expected to be constant across the emission band [4]. When achiral molecule is in the excited electronic state, chirality may be induced via intermolecular interaction with a chiral molecule. Quantum electrodynamical theory showed that chirality in the achiral molecule is induced dynamically by the coupling of virtual transitions of the chiral molecule to those of the achiral molecule [5]. At this, CPL occurs in the wavelength range characteristic of achiral molecule, whereas chiral molecule does not absorb light in this wavelength. Most experimental results of protein-related CPL can be explained by the idea of induced chirality in the excited electronic state. No signal related to circularly polarized emission reflects the chirality of chromophore. Measured circularly polarized signal may result from artifacts of either the measuring system or macroscopic properties of the sample (see below). Other types of non-natural CPL are magnetic circularly polarized luminescence (MCPL; [6]) and fluorescence-detected circular dichroism (FDCD). MCPL appears as a result of orientation of chromophore in a static magnetic field. This effect is similar to the effect of photoselection when a molecule of some orientation has more chance to be excited by light of given polarization than does a molecule of another orientation. If the excitation light was polarized either linearly, or circularly, or elliptically, the CPL will depend on both chirality of chromophore and excitation polarization. FDCD is an example when the difference between intensities of fluorescence excited by left-handed and right-handed circularly polarized light is a matter of interest. In fact, FDCD is a specific mode of CD detection. In the present chapter, we consider only natural CPL. Both MCPL and FDCD topics are omitted. Several reviews describe the CPL principles in more detail [4, 7–14]. It is implied that CPL of multi-chromophore system Smc consists of CPL of individual chromophores. Regarding proteins, the systems may be multi-tryptophan proteins, proteins forming complexes with more than one ligand such as probes, cofactors, lanthanides, bilirubin, chlorophyll, in our case. How can gem of Smc be presented via gem,i of chromophores?

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Assuming f = fl – fr for Smc and φ i = φ li – φ ri for ith chromophore, it may be shown that f = φ i and f = φ i , which leads to gem = gem,i

φi f

(5)

where the gem factor is determined by Eq. (1). Therefore, gem of the multichromophore system is not a simple sum of gem,i of individual chromophores but a linear combination of gem,i with coefficients which are contributions of related chromophores to the total fluorescence of the system. A chromophore of high gem,i may contribute little to the total gem factor if its contribution to the total fluorescence is low. In particular, gem may equal zero when different non-zero gem,i have different signs. In this case, it is useful to analyze the f signal. Equal gem,i of all chromophores determine the same gem = gem,i of the whole system. Thus, the interpretation of the multi-chromophore CPL in terms of individual CPL is not trivial. Time-resolved CPL of a few chromophores was developed assuming that during the luminescence decay no change in each chromophore gem,i occurs [15]: gem (t) =

αi gem,i exp ( − t/ τi ) αi exp ( − t/ τi )

(6)

where α i and τ i are the pre-exponential factor and the fluorescent lifetime of the ith chromophore, respectively. It is obvious that α i = φ i0 /f0 , where subscript zero denotes t = 0. According to Eq. (6), the luminescent time dependence becomes mostly important for complicated molecular systems of two or more chiral chromophores. This feature supported by a related instrumentation [16] provided an opportunity to determine gem of two different chromophores in two separate samples simultaneously [15].

2 How to Measure CPL? The measurement of CPL signal is not a trivial task because circularly polarized part of the total luminescence is pretty small. CPL signal regularly varies in the range of gem ∼ 10−2 –10−5 . At the error δf of the luminescence intensity measurement of ∼ 10−2 , the CPL f is less than error δf. To extract the CPL signal from the total luminescence with error of δgem ∼ 10−5 , a few approaches are used.

2.1 Instrumentation In contrast to CD, there are no commercial instruments for CPL measurements. All studies are performed with lab-built setups, which may be divided into three major types. Figure 1 shows their general schematics. All types have a common part and a


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Fig. 1 General schemes of three types of CPL instruments. Amp, amplifier; DGG, delay gate generator; Disc, discriminator; DPC, differential photon counter; I, IR , and IL , signals related to the total emission intensity and right- and left-handed circularly polarized emission intensities, respectively; L, lens; Lock-in, lock-in amplifier; M, emission monochromator; P, linear polarizer; PEM, photoelastic modulator; PMT, photomultiplier tube; S, sample. Dashed rectangle limits the common part of all instruments

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part specific to the given type. There may be variations in setup in the frame of the same type. Some details are available in reviews [9–11, 13, 17, 18]. The common part closely resembles the scheme of regular spectrofluorometer: emission may be excited by the light of various sources (lamps, excitation monochromators, lasers, etc.) at various angles with respect to the direction of the emission collection. It may be polarized but depolarization is strongly recommended in order to avoid (i) registration of non-natural CPL, (ii) “contamination” of the signal by parasite polarization, and (iii) photoselection. A lens provides the fluorescence collection and focuses it at the entrance slit of emission monochromator. Photodetector (photomultiplier, in most cases) is placed at the exit slit. Pre-amplifier converts the photocurrent to photovoltage and amplifies it. In addition to this regular scheme, all common parts contain a tandem of quarter-wave 50-kHz modulator (photoelastic modulator PEM was found to be most convenient) and linear polarizer oriented at 45◦ to the PEM optical axis provides modulation of only circularly polarized part of the total emission, leaving the nonpolarized part non-modulated. To extract the modulated photocurrent from the total one, different ways are used. SG instruments. Steinberg and Gafni (SG) at the Weizmann Institute of Science, Israel, in 1972 have described a phase-sensitive scheme, in which the modulated photovoltage is directed to the input of a lock-in amplifier connected with the PEM controller [9, 19] (Fig. 1a). If the frequency and the phase supplying the PEM head by the PEM controller are equal to the frequency and the phase of photovoltage, then the lock-in amplifier will amplify only circularly polarized part of the total photocurrent and provide an output proportional to the CPL intensity. Total photovoltage is proportional to the total fluorescence. Therefore, two simultaneous electrical signals (direct photovoltage and lock-in output) represented as f and f determine gem according to Eq. (1) by means of either an analog circuit or a digital calculation when applied to an A/D converter board in a computer. The time consumption of the measurement is less than 1 min. Most of the protein CPL studies were done using this instrument. Although in the SG instrument the lock-in amplifier output may be unstable because of the electronic ground loops, the instability can be effectively compensated by multiple readings in a few minutes, reducing the error of measurement to δgem ∼ 10−5 . Double modulation (excitation light is chopped at low frequency of 13 or 200 Hz in addition to photoelastic modulation of 50 kHz) [13, 20, 21] has been expected to reduce the instability. SG scheme was the only type of CPL instrument until Schippers, van den Beukle, and Dekkers (SBD) from the University of Leiden, the Netherlands, proposed SBD instruments in 1982 [22] (Fig. 1b). The features of this scheme are photon counting and discrimination between the left- and the right-handed CPL with the help of a differential photon counter (DPC). The discrimination is based on the gate formation related to positive and negative sinusoidal half-wave of 50 kHz modulation. The output of DPC consists of a photopulse number proportional to the total fluorescence intensity f and the number proportional to its circularly polarized part f. This variant of the CPL instrument has been described in detail elsewhere [11, 12, 22].


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In protein research, this instrument was employed at the CPL study of lanthanide– protein complexes (see Table 5). It was reported that the time consumption of data collection of weakly luminescent system might approach 20 h [23]. SSG instruments. Schauerte, Steel, and Gafni (SSG) from the University of Michigan, Ann Arbor, MI, in 1992–1995 reported the detailed schemes allowing to measure time-resolved CPL in microsecond and nanosecond timescale [16, 24] (Fig. 1c). This scheme is also convenient for steady-state CPL measurements. It discriminates between left- and right-handed circularly polarized emissions similar to the SBG instrument, but in contrast to the SBG, left and right CPL signals are analyzed separately, and the difference between them corresponds to real CPL. The time resolution approaches 1–2 ns. The time-resolved instrument built by the optical scheme of SBG instrument [25] provides a 50 ns time resolution. In the last version [16], a time-to-digital converter (TDC) has been applied transforming the instrument from analog to a digital one. The SSG instrument has been constructed to be convenient for protein research.

2.2 Calibration Ideally, the SBG instruments do not need special calibration because gem is determined via photon counting [2]. However, real circumstances introduce some systematic errors in the gem values [22]. The correct gem may be affected by (i) difference in the count rates in left- and right-handed half-periods of the modulation cycle, (ii) dark count rate at low emission intensity, (iii) the non-ideal features of PEM-associated polarizer, and (iv) dimensions of the fluorescent spot in the sample and the light collection angle. To calibrate the CPL instruments, both chemical standards and optical device were proposed. Steinberg and Gafni [19, 26] have developed a device consisting of a pair of quarter-wave and fused silica plates which generate circularly polarized light from non-polarized light. The device artificially converts non-polarized fluorescence (i.e., fluorescein in NaOH) to CPL with a desired gem value in the range of 0.00– 0.045 at 350 nm or 0.000–0.043 at 500 nm when the angle between the normals to the plane of the plates varies from 0◦ to 35◦ [9]. The device is simple and suitable for fast, reproducible, and effective calibration. Bartett et al. [27] have used a CD-active compound instead of such device. The totally circularly polarized fluorescence has been obtained by passing the fluorescein total emission though a Fresnel rhomb [28]. After Luk and Richardson have reported on CPL of d-camphorquinone in ethanol [29], Gafni et al. [16] used (+) and (–)camphorquinone as a stable standard characterized by gem = ±8.4×10−3 with accuracy of 3.5×10−4 . Fluorescence was excited at 300–308 nm. A 418-nm cutoff high-pass filter allowed the measurement of wavelength-integrated fluorescence over the wavelength range of 480–600 nm [29]. Respectively, the gem value is a mean value over this wavelength range. It is

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important to emphasize that, first, Gafni et al. applied depolarized laser excitation, which provides a natural CPL measurement, independent of light source polarization. Second, authors calibrated their instrument with the device mentioned above. Third, shapes of the CD and CPL spectra were found mirror symmetrical and, hence, Qg and Qk coordinates of the ground-state and excited-state equilibrium conformations in Eqs. (3) and (4) were equal [29], resulting in gab = gem . Therefore, gem of camphorquinone in cyclohexane at a certain wavelength λem can be determined by measuring gam at wavelength λam = 2λ0 – λem , where λ0 is the mirror-symmetry axis wavelength (roughly about 510 nm; see [29]; as known, any CD spectrometer may be reliably calibrated with camphorsulfonic acid). Tris(3-trifluoroacetyl-D-camphorato)europium(III), Eu(facam)3 in dimethyl sulfoxide (DMSO) [30], was considered as a stable standard for the absolute CPL determination [12]. Brittain and Richardson [30] measured the Eu(facam)3 gem of –0.84, –1.98, and +0.38 at 588, 595, and 613 nm, respectively. However, Schippers et al. [22] reported gem of –0.25, –0.78, and +0.072 at the same wavelength and did not find the origin of the inconsistency with the Brittain–Richardson’s data, concluding that the use of Eu(facam)3 as a CPL standard is questionable. It is possible that the discrepancy may result from the circularly polarized excitation used by Schippers et al. and be able to affect the measured CPL signal even though the luminescence collection was performed at 90◦ . A calibration was probably not employed. In contrast, Brittain and Richardson [30] have measured CPL with the SG instrument, which implies depolarization of the excitation light and calibration. Because the chemical stability of Eu(facam)3 in DMSO is a serious advantage, this compound seems to be convenient to serve as a CPL standard after performing the additional measurements of its gem value.

2.3 Artifacts In Eqs. (3) and (4), gab and gem relate to isotropic collection of absorbing and emitting molecules, which regularly occurs when Brownian motion is predominant. However, if the excited chromophore orientation distribution is not randomized, then the molecule at some orientation has more chance to be excited by light of given polarization than the molecule at another orientation. This effect is called photoselection and has been described by Steinberg in detail [4, 9]. Ignoring the photoselection may result in the erroneous interpretation of the measured signal as related to natural CPL. To avoid photoselection, a depolarized excitation seems to be effective. However, even at the polarized excitation, photoselection has no effect when electric transition dipole moments involved in the emission and the excitation are parallel or when emission is collected at the magic angles of 54.75◦ or 125.25◦ to the excitation beam. Small inherent birefringence of PEM may also determine some artifact signals that originated from non-ideal operation of lock-in amplifier as discussed elsewhere [12, 31]. This signal was found to be additive with real CPL signal [31, 32]. It


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is possible in practice to eliminate this artifact by using a non-polarized excitation [32]. Although in some specific individual cases it was possible to minimize the artifacts produced by polarized excitation [12], in general, if truly natural CPL which reflects the chirality of the chromophore in its excited electronic state is desired, then one should depolarize excitation light.

3 Circularly Polarized Intrinsic Fluorescence of Proteins Intrinsic protein fluorescence is determined only by tryptophan (Trp) residues in the wavelength range of 300–450 nm when excited at wavelengths higher than 295 nm [33, 34]. At the excitation wavelength of 275–290 nm, a tyrosine (Tyr) component occurs as a supplement to the tryptophan fluorescence in the wavelength range of 300–340 nm. Tryptophan-free proteins show only tyrosine fluorescence. Phenylalanine fluorescence of proteins is too weak to significantly contribute to the total protein fluorescence. Therefore, protein intrinsic CPL is determined by phenol of tyrosine residues and indole of tryptophan residues. Collagen and elastin, as well as green/yellow fluorescent proteins, chromophores of which are not aromatic amino acids residues, were not the subject of the CPL studies. As well, no CPL of tryptophan-less proteins, fluorescence of which is determined by tyrosine residues only, has been considered so far.

3.1 Peptides It should be emphasized that neither free tyrosine nor free tryptophan in solution shows measurable CPL (Table 1). Tyr- or Trp-containing linear peptides show negligible CPL as well [35]. Cyclic dipeptides can exhibit significant gem factor values when they are in highly viscous media (Table 1), while in low-viscosity dioxane or dimethyl sulfoxide, no CPL was detected [35]. In contrast, gab was significant in viscous and non-viscous solutions. When in the viscous media, gab is similar to gem . It was concluded [35] that in viscous medium, relaxation of molecular conformation is probably arrested within period between the excitation and the emission. In other words, in this case, the asymmetry of aromatic amino acids in the excited electronic state copies the asymmetry in the ground electronic state even in the randomized environment.

3.2 Poly(α-amino acid) To recognize the nature and properties of protein fluorescence, poly-Tyr and polyTrp fluorescence research has played a significant role [36–40]. Similarly, peptide and polypeptide intrinsic CPL would serve as model investigations. Unfortunately,

434

E. Gussakovsky Table 1 CPL of tyrosine- and tryptophan-containing peptides

Peptide

Solvent

Emission (nm)

gem ×104

L-Tyr L-NATA L-TRP-GLY3 -COOH Carbobenzoxy-Gly3 -L-Trp-Gly3 o-Benzyl ester Carbobenzoxy-Gly4 -L-Trp-Gly3 o-Benzyl ester Linear L-Tyr-L-Tyr Linear L-Trp-L-Tyr Linear L-Tyr-L-Trp Linear L-Trp-L-Trp Cyclo(Gly-L-Tyr)

POP-D/Dioxane POP-D/Dioxane

300–340 310–370

0 0 0 0

Cyclo(L Tyr-L-Tyr) Cyclo(L-Trp-L-Val) Cyclo(L-Trp-D-Val) Cyclo(L-Trp-L-Phe) Cyclo(L-Trp-L-Tyr) Cyclo(L-Trp-L-Trp)

0

Dioxane Pop-D Dioxane Pop-D Dioxane Pop-D Dioxane Dioxane Dioxane Dioxane

300–340 300–340 300–340 300–340 310–370 310–370 310–370 310–370 310–370 310–370

0 0 0 0 0 ∼6 3.5–4 4–5 0 2 0 0 0 0

The data are retrieved from [35]; room temperature; excitation at 254 or 275 nm; l-NATA, N-acetylL-tryptophanamide; POP-D, polyoxypropylene in dioxane. Zero values mean |gem | < 3×10–5 .

aromatic amino acid polymers were not a goal of CPL studies related to proteins. The significance of such research is emphasized by reports of Sisido et al. [41–43], where they consider structure, spectroscopy, and CPL of two poly-L-Ala polymers in which alanine residues are modified by 1- or 2-naphthyl chromophores (poly(L-1 or 2-naphthylalanine) or poly(1 or poly-2-NapAla)). Fluorescence spectrum consisted of two bands: monomer fluorescence of naphthyl group at 310–360 nm and excimer emission at 370–450 nm. Neither poly (1-NapAla) nor poly(2-NapAla) shows CPL at the band of monomer emission. The intrigue here is that at the excimer band, poly(1-NapAla) has low gem of about –1×10−4 , while poly(2-NapAla) has much higher absolute gem of –20×10−4 . According to the total fluorescence intensity spectrum, excimer formation in poly(1-NapAla) was negligible in comparison with poly(2-NapAla). Regarding polymer structure, empirical conformation energy calculations and CD data allowed to consider that poly(1-NapAla) in the ground electronic state forms either left-handed 310 -helix or right-handed δ-helix, while poly(2-NapAla) forms left-handed α-helix [42]. Probably, CPL of these polyamino acids is connected with both excimer formation and helix structural parameters. No CPL at the monomer band may result from symmetrical surrounding of single naphthyl groups in the excited electronic state. But when the steric conditions allow forming a dimer


435

of excited naphthyls, this can be monitored by a large CPL signal. Some helical structures probably provide the proper steric conditions: they exist in left-handed α-helix and are absent in left-handed 310 - or right-handed δ-helix. It does not matter that in proteins, left-handed α- or 310 -helices and δ-helices are absent. The crucial point is that the distance between naphthyl chromophores in these helices is varied and affects the eximer formation. Such situation can serve as a model for proteins with multiple tyrosine/tryptophan residues where eximer formation is possible similar to poly-Tyr [37] or D-glyceraldehyde-3-phosphate dehydrogenase [44]. Also, tryptophanyls can form exciplexes in proteins (see Section 3.3).

3.3 Tyrosine and Tryptophan Residues in Proteins Because free tryptophan and tyrosine as well as small peptides in solution do not show measurable CPL, these aromatic amino acids determine protein CPL as a result of induced asymmetry in the excited electronic state. The known intrinsic CPL of proteins is limited mostly by tryptophan fluorescence (Table 2). The exception is subtilisins Carlsberg and Novo, the total fluorescence and CPL of which at 300–340 nm at the 280 nm excitation are significantly (not completely) determined by tyrosine residues. A large number of tyrosine residues in these proteins result, probably, in the dependence of gem on wavelength (increase from –2×10−4 –1×10−4 at 300 nm to 3×10−4 –4×10−4 at 320 nm) and do not allow detailing CPL for each residue. However, the various tyrosine gem values for subtilisins Carlsberg and Novo indicate that the asymmetry of tyrosine residues in the excited electronic state is varied. Intrinsic CPL of single tryptophan protein is expected to be constant across the spectrum, while CPL spectra of multi-tryptophan proteins is structured. Indeed, single tryptophan CPL of azurin, subtilisin Carlsberg, and staphylococcal nuclease is wavelength independent. However, although human serum albumin (HSA) also has one Trp, its CPL is strongly dependent on wavelength, probably, because of the HSA heterogeneity. The experiments with naphthyl CPL of poly(NalAla) (Section 3.2) and acridine CPL of poly-D-Glu/poly-L-Glu (Section 4) showed that CPL is significant when polyamino acids are in the helix form. It can be expected that tryptophan CPL of proteins may depend on whether Trp is in the ordered secondary structure. The answer is evident for single tryptophan proteins. In subtilisin Carlsberg, HSA and staphylococcal nuclease, the single tryptophan residue, is included in the α-helix and their gem factor changes from 2×10−4 to 9×10−4 (Table 2). In azurin and apoazurin, the single tryptophan residue is in the random part of the backbone, but their gem of 5×10−4 and 3×10−4 is in the same range as for α-helical residues. It does not seem that a correlation between tryptophan CPL and involvement of this residue in ordered secondary structure exists.

7 7

S. Nuclease + Ca2+ S. Nuclease + 6 M Gnd

4

8

α-Lactalbumin + Cacl2

GPDH+NAD+

10

17 17 17 7

HSA HAS + 6 M Gnd HAS + 6 M Gnd + DTE S. Nuclease

Multi-Tryptophan Proteins Subtilisin Novo

2 13

2

Single Tryptophan Proteins P.a. Azurin

P.a. Apoazurin Subtilisin Carlsberg

Ntyr

Protein

3

4

3

1 1

1 1 1 1

1 1

1

Ntrp

5.5–5.0 4.2–2.9 2.9–2.5 7.5 7.5

5–8.3

6.8 6.8 6.8 7.0 3.5 1.5 7.0

7.0 1.5 7.0 5–8.3

pH

280 300 280 280 280 280 280

270–295 270–295 270–295 280 300 254, 275 254, 275 254, 275 254, 275 254, 275 254, 275 254, 275 254, 275

Excitation

300–340 360–390 340 340 340 310–320 390

300–350 300–350 300–350 300–340 350–390 320–340 320–340 310–370 310–370 310–360 310–360 310–370 310–370

Emission

Table 2 Circularly polarized intrinsic fluorescence of proteins

(0–3.5)±0.5 –3±0.5 3.0±0.4 1.5±0.3 0.3±0.4 4±1 –12/–16/– 20/–23A

5±0.5 3.5±0.5 3±0.5 (–2 to 2)±0.5 2±0.5 9±1 2.5±0.5 0 3.5±0.5 2.5 2 3.5±0.5 0

gem ×104

[48] [48] [49] [49] [49] [51] [51]

[99] [99] [99] [48] [48] [8] [8] [8] [8] [8] [8] [8] [8]

References

436 E. Gussakovsky

275 275 275 275 275

275 275 275 254, 275 254, 275 254, 275

7.3 7.3 7.3 7.0

Excitation

7.5 7.4 7.4 7.4 7.4 7.4

pH 340 310–360 310–340 310–360 315–360 310–340 310–360 310–360 330–370 330–370 330–370 310–320 320–360 310–410

Emission 5 –5±0.5 5±0.5 –5±0.5 4±0.5 –9±0.5 –12±1 –9±0.5 –(3–5)±0.5 –(4–5)±0.5 –(2–4)±0.5 –3.5 3 0

gem ×104

[60] [53] [53] [53] [53] [52] [52] [52] [54] [54] [54] [8] [8] [8]

References

NAD+ molecules per one GPDH Anti-PA, anti-type III pneumococcal antibody; C. pepsinogen, chicken pepsinogen; DTE, dithioerythritol; Gnd, guanidine hydrochloride; GPDH, lobster glyceraldehyde-3-phosphate dehydrogenase; HSA, human serum albumin; IgA, protein-315, myeloma immunoglobulin A protein; LDH, pig heart lactate dehydrogenase; Ntyr and Ntrp , number of tyrosine or tryptophan residues, respectively; P.a. azurin, azurin from Pseudomonas aeruginosa; S. nuclease, staphylococcal nuclease. All CPL data were obtained with the SG [19] instrument. Some gem factor values were retrieved from figures published in related references. Number of aromatic amino acids is according to Protein Data Bank www.rcsb.org/pdb. Excitation and emission values are in nanometers.

5 5 5

17 17 17

a 0/1/2/3

6

7

LDH Anti-RNAse Anti-RNAse Fab’ Anti-Poly(Ala) Anti-Poly(Ala) Fab’ IgA IgA Fab’ IgA Fv IgA reduced Anti-PA Anti-PA Fab’ Pepsinogen Pepsinogen + 8 M Gnd Pepsinogen + 8 M Gnd + DTE

Ntrp

Ntyr

Protein

Table 2 (continued)

Circularly Polarized Luminescence of Proteins and Protein Complexes 437

438

E. Gussakovsky

On the basis of fluorescence parameters, tryptophanyls were classified according to their location in the protein globule [33, 34, 45]. Is there a correlation between this classification and various gem factors of tryptophanyls? Spectral form A corresponds to the emission of unperturbed indole in the neutral hydrophobic environment inside protein globule. Spectral form S corresponds to the emission of indole located inside the protein globule forming a 1:1 exciplex with neighboring polar groups. Single indole in azurin is assumed to represent both form A and form S [46]. Its gem = 5×10−4 (Table 2). Spectral form I corresponds to the emission of indole inside the protein globule forming a 2:1 exciplex with neighboring polar groups. Spectral form II corresponds to the emission of indole at the protein surface in contact with bound water. Single indole in staphylococcal nuclease or human serum albumin (HSA) has spectral property of form II [46]. Their gem are 3.5×10−4 and 9×10−4 , respectively (Table 2). Single tryptophan of subtilisin Carlsberg determines the fluorescence with maximum at about 350 nm, which with 80% probability [47] corresponds to the spectral form III corresponding to the emission of indole located at the protein surface in contact with free water [34]. This tryptophan has gem = 2×10−4 close to gem of nuclease (form II). All three tryptophanyls of subtilisin Novo belong to the same spectral form II [46], but its gem = −3×10−4 is negative in the wavelength range of 360–390 nm [48]. In contrast, indoles of α-lactalbumin represent two different spectral forms S and I [46], while its gem of 3×10−4 (Table 2) does not depend on wavelength (all gem,i s are the same) [49]. These samples indicate that there is no correlation between spectral forms of tryptophan residues in proteins and the gem spectra, its absolute value and sign. Considering multi-tryptophan proteins, one should keep in mind that not every tryptophan residue can contribute to the total fluorescence and to CPL (cf. Eq. (5)). α-Lactalbumin tryptophan fluorescence is probably determined by three of four residues (Trp-28, -108, and -123) [50]. Sommers and Kronman argued that there is a 100% excitation energy transfer between Trp-28 and Trp-108, and their quantum yields are comparable. From Trp-28 and Trp-108, the excitation energy is channeled to Trp-63 where vicinal disulfide groups quench it. Therefore, Trp-123 contributes more than 50% to the total fluorescence intensity, while Trp-28 and Trp108 contributions are about 20% each. Because both Trp-123 and Trp-28 are buried in a hydrophobic pocket and Trp-108 is partially shielded from solvent, the spectral contribution of each of them to the total fluorescence of α-lactalbumin is similar. CPL of calcium-loaded bovine α-lactalbumin at pH 5.3 is almost wavelength independent in the 320–350 nm range [49], which reflects homogeneity of tryptophan CPL (all tryptophanyl gem,i are equal to 3×10−4 ). At lower wavelengths, the increase in gem probably is connected with tyrosine fluorescence, which occurs at the 280 nm excitation. The tryptophan gem of α-lactalbumin probably results from gem of Trp-123 and the pair of Trp-28+Trp-108. It is unrealistic that Trp-28 and Trp-108 form some excimers because no long-wavelength band, characteristic of such complexes, was observed and the distance of 40 nm between two indoles [50] is too big to allow an excimer formation. No excimer in poly-Trp (a single band with maximum at


439

about 340 nm [38, 40]; spectral form II) supports this conclusion. Hence, CPL of this protein should be determined by the asymmetry of Trp-28, -108, and -123 in the excited electronic state. The absorption asymmetry factor gab of this protein is really wavelength dependent and is negative in contrast to gem . This indicates that the environmental asymmetry of indoles in the ground and excited electronic states is crucially different or, in other words, the asymmetry is excitation sensitive. Schlessinger and Levitzki [51] measured intrinsic CPL of glyceraldehyde-3phosphate dehydrogenase (GPDH) in the wavelength range of 310–390 nm at the excitation wavelength of 280 nm. CPL significantly depends on wavelength, changing from about 4×10−4 at 310–320 nm (tyrosine and tryptophan) to –10×10−4 at 360 nm and –12×10−4 at 390 nm (tryptophanyls only) and showing heterogeneity of three tryptophans in regard to their asymmetry in the excited electronic state. The CPL spectra of immunoglobulin A (IgA) and its fragments were found different [52]. For IgA, gem = –9×10−4 from 305 to 335 nm and gradually changes to –2×10−4 at 370 nm. At the same time, gem of its Fv fragment is constant (gem = –9×10−4 ) across whole wavelength range and that of its Fab’ fragment varies from –8×10−4 to –12×10−4 . Light and heavy constituents of Fv (gem = 0 and gem,max = 4×10−4 , respectively) [52] also cannot be combined into a linear combination to reach the gem Fv value of –9×10−4 . Heavy and light chains of IgA have positive gem in contrast to negative gem of whole IgA [52]. Similar big sign-inversion difference between CPL of intact protein and its Fab’ fragment has been observed for anti-RNase, anti-poly(D-alanine), and “loop” of lysozyme [53]. The difference in CPL of whole protein and its parts has served as an argument supporting the idea of conformational transition during biochemical decomposition of IgA or Fv. Not only conformational transition but also the non-linearity of gem must be taken into consideration because whole protein and its fragments have different sets of fluorescent tryptophanyls which may result in different gem , even though individual gem,i of IgA and its fragments are the same. Fragmentation of immunoglobulin may not necessarily lead to the dramatic variation in CPL: the gem spectra of anti-type III pneumococcal antibody (anti-PA in Table 2) and mouse myeloma protein McPC 603 are similar to those of their Fab’ fragments [53, 54]. Moreover, reduction of disulfide bonds of immunoglobulin, which leads to global protein disordering, determines only minor variation in CPL spectrum [53, 54], which is incomparable with the big sign-inversion differences between whole IgA and its fragments. This finding supports the idea that the sign-inversion changes of gem at the antibody fragmentation are a result of the non-linearity of gem rather than the cardinal perturbation of protein chain conformation.

3.4 Complexes of Proteins with Functional Non-fluorescent Agents The important application of protein intrinsic CPL is the description of protein interactions with functional agents. If the agents are fluorescent, then their CPL can be used to probe the binding sites (see Section 4). Here we consider functional agents

440

E. Gussakovsky

which do not fluoresce measurably and intrinsic CPL of protein may report on their effect on conformational properties of protein. The binding of non-fluorescent agent does not change the chromophore composition of protein but may affect individual gem,i of Trp or Tyr via induction of a conformational transition. In other words, the change in protein gem results from perturbation of local or global protein conformation by the agent if it does not affect the tryptophanyl directly. Glyceraldehyde-3-phosphate dehydrogenase (GPDH) reversibly converts glyceraldehyde 3-phosphate to D-glycerate 1,3-biphosphate reducing nicotinamide dinucleotide (NAD+ ) to NADH. So non-fluorescent NAD+ is a coenzyme which forms a complex with the enzyme. Binding NAD+ significantly decreased negative tryptophan CPL of GPDH from gem = –12×10−4 for apo-enzyme to gem = −23×10−4 for GPDH–NAD+ 3 . The antigen–antibody interaction (anti-RNase with RNase, anti-lysozyme “loop” with “loop” [53]) led to the conversion of the gem spectrum from wavelength independent (all individual gem,i are equal) to S-shaped. For other immunoglobulins [anti-poly(D-alanine), protein McPC 603 [53]], the spectral changes were not so pronounced. Obviously, the antigen–antibody interaction may either induce a heterogeneity of individual tryptophanyl gem,i or slightly modify them. CPL has been used for revealing oligosaccharide-induced structural changes in a homogeneous type II pneumococcal antibody and its Fab’ fragment [54]. Indeed, significant changes were observed in the presence of haptenic oligosaccharides.

3.5 Protein Conformation Perturbation Effect of the protein conformation perturbation on CPL was a goal of many studies. The reasons were based on the obvious idea that asymmetry of the excited electronic state is sensitive enough to respond to small variation in the chromophore (mostly indole of tryptophanyl) environment. It can be proposed that such sensitivity may be even higher than asymmetry in the ground electronic state because of higher susceptibility of the excited state to the environment. Strong denaturing agents like guanidine hydrochloride (GndHCl) and urea of high concentrations are known to significantly destroy protein structure. Respectively, in such cases, tryptophan CPL often decreases to zero (staphylococcal nuclease in 6 M GndHCl, IgA and its Fab’ and Fv fragments in 6 M urea [52], subtilisins Carlsberg and Novo in 8 M GndHCl [8, 48]). For some proteins, full denaturation occurs in GndHCl after reduction of disulfides and leads to zero CPL (chicken pepsinogen, human serum albumin [8]). pH of solutions (excluding strong acids or alkalis) makes less dramatic changes than do GndHCl and disulfide reduction. Tryptophan fluorescence maximum of subtilisin Carlsberg shifts from 350 to 360 nm at the pH change from 5 to 8.3, indicating that more tryptophanyls became exposed to water (re-arrangement of tryptophanyls among spectral forms; see Section 3.2) at conformational transition. However, no changes in gem were detected, obviously, because of no changes in the asymmetry


441

of tryptophanyls in the excited electronic state. This also confirms the absence of correlation between tryptophanyl spectral forms and CPL. As the pH of the bovine α-lactalbumin solution was decreased from 5.5 to 3.5, gab did not change, while gem was reduced from 3×10−4 to 1×10−4 with pKa of 4.9 [49]. The result evidences that there were no changes in the ground-state asymmetry, while the excited-state asymmetry reduced significantly. In other words, gem appeared to be more sensitive to tiny changes in protein conformation affecting the environmentally induced asymmetry of the excited chromophore rather than asymmetry of the ground electronic state. Extra high susceptibility of CPL allowed revealing the existence of premolten globule state of bovine α-lactalbumin, which had been predicted theoretically but was not observed by any other experimental techniques [49].

4 Protein Complexes with Fluorescent Agents Generally, fluorescence of functional or non-functional agents forming the complexes with proteins lies in the visible region. This makes it possible to measure their fluorescence independently on intrinsic tryptophan fluorescence. Regarding CPL, such fluorescence agents studied so far are fluorescent probes 8-anilino1-naphthalenesulfonic acid (ANS), 2-p-toluidinylnaphthalene-6-sulfonate (TNS), acridine orange, 9-amino acridin, and dansyl derivatives; fluorescent functional agents reduced nicotinamide adenine dinucleotide (NADH), 1,N6 -ethenoadenine nicotinamide dinucleotide (εNAD+ ; fluorescent analog of NAD+ ), pyridoxal 5 phosphate (PLP) and bilirubin, and lanthanides which mimic functional cations and have pronounced fluorescence (fluorescent pseudo-functional agents).

4.1 Probes Table 3 summarizes the results of the CPL studies of fluorescent probes of proteins. Schlessinger and Steinberg have pioneered the application of CPL of fluorescent probes to protein research [55]. Anthranoyl group covalently attached to chymotrypsin and TNS non-covalently attached to this protein were used as CPL reporters on active and non-active sites, respectively. TNS has been synthesized as a hydrophobic fluorescent probe for protein conformation research, quantum yield of which varies from almost 0% in water (0.08%) to 18% when bound with chymotrypsin [56]. CPL of both the probes was wavelength independent as expected for a single chromophore. Both gab and gem were negative but not equal. From experiments with TNS it was concluded that the TNS-binding site induces the asymmetry of the TNS chromophore in the ground electronic state. Upon excitation, absolute value of g-factor increases showing that the asymmetry is higher in the excited electronic

442

E. Gussakovsky Table 3 CPL of fluorescent probe complexes with protein

Protein

Probe

pH

Anti-Dansyl

Dansylamide

7.2

α-N-Dansyl-L-Ala

7.2

α-N-Dansyl-L-Ala amide

7.2

ε-N-Dansyl-L-Lys

7.2

Anti-Alanine α-Lactalbumin

1 ANS

Chymotrypsin

TSN Anthraniloyl 2 2 Acridine Orange Acridine Orange 9-Amino Acridine

MOPC-315 IgG Poly-L-Glu Poly-D-Glu Poly-L-Glu

5–6 3–4 8.05 6.8

Excitation Emission (nm) (nm)

gem ×104 References

320 365 320 365 320

480–490 470 470–560 470–480 480–590

7/6/4.5a 11/10/12a 3 7.5 4/3.5b

[59] [59] [59] [59] [59]

365 320 365

470–480 470–480 470–480

365 365

490 490 460–520 420–500 410–490 410–490 510–560 510–560 450–460

11/10b 4 9.5 0 –1.8±0.2 –0.9±0.2 –2.5 –3.5 5.5 0 ∼ 40 ∼ –40 ∼2

[59] [59] [59] [59] [49] [49] [55] [55] [59] [59] [8] [8] [4]

365 365 4.3 4.3 4.6

a 0.25/0.5/1.05

hapten molecule per site hapten molecule per site 1, 1-alanyl-2-dansyl diaminoethane; 2, 1-(2 , 4 -dinitro aniline-2(9 -amino)acridinylethane, a fluorescent derivative of 2,4-dinitrophenyl hapten; ANS, 8-anilino-1-naphthalenesulfonic acid; IgG, normal mice immunoglobulin G; MOPC-315; an MOPC-315 protein; poly-Glu, poly-glutamic acid; TSN, 2-p-toluidinylnaphthalene-6-sulfonate. All data were obtained with the SG-type instrument [19]. b 0.33/1.05

state than in the ground state. Based on the CPL it was concluded that orientation, freedom, and strength of TNS binding might have changed upon electronic excitation. Another example of the efficiency of probe CPL is CPL of ANS [49], which fluoresces intensively in hydrophobic regions of proteins (quantum yield of 30–80% [57]), while in the water surroundings, its quantum yield is close to 0 (0.6%) [57] similar to TNS. This property of the dye provides a simple way to investigate the hydrophobic binding site in protein by observing the ANS total fluorescence. The CPL properties of ANS bound to protein have allowed revealing even more tiny conformational properties of α-lactalbumin [49]. The pH dependence of ANS gem showed an additional conformational transition at pH 4–5, which coincides with pKa of Ca2+ dissociation. It was suggested that in the pH range of 3.7–4.1, α-lactalbumin exists in an intermediate state between native and acid-denatured states and that this state is a premolten globule state characterized by a reduced Ca2+ binding, a native-like tertiary structure, and reduced asymmetric fluctuations of the tertiary structure on the nanosecond


443

timescale. This intermediate resembles the “critical activated state” which was theoretically deduced [58] but never shown experimentally. The results demonstrate the power of CPL in the investigation of folding/unfolding transitions in proteins. Fluorescent quantum yield of dansyl amide, ε-N-dansyl-L-Lys, α-N-dansyl-LAla, and α-N-dansyl-L-Ala amide markedly enhanced when bound to anti-dansyl antibodies in comparison with water solutions [59]. As for other chromophores, free dansyl probes in solution showed no CPL, while the complexes exhibited positive gem , slowly depending on wavelength at the 320 nm excitation. At the 365 nm excitation, the gem (λ) spectra changed as a result of heterogeneity of binding sites. Obviously the probe CPL is induced and reflects asymmetry of the binding sites. No CPL of a complex of anti-alanine antibody with 1-alanyl-2-dansyl diaminoethane, when hapten was bound to antibody via alanine, was detected [59]. Probably in this case, dansyl chromophore does not experience specific interaction with immunoglobulin molecule, which provides its asymmetry in the excited electronic state. Acridine orange and 9-amino acridine were employed as fluorescent probes in poly(glutamic acid) conformation studies [8]. gem of acridine orange vanished in random conformation and increased to 50×10−4 –100×10−4 when poly(L-Glu) is in helical form. At this, handedness of glutamate in polymer inverts the gem sign (Table 3) [8]. Because the large CPL signal is a result of induced asymmetry of the excited chromophore, the inversion of the CPL signal sign evidently reports on the inversion of the asymmetry of the probe-binding site at the transition from left-handed to right-handed glutamate.

4.2 Cofactors and Bilirubin The ability of physiologically active compounds to fluoresce makes them the unique reporters of their own binding sites in proteins. Regarding CPL, complexes of proteins with NADH, εNAD+ , PLP, and bilirubin were studied. The results of the CPL studies of these compounds are compiled in Table 4 . Similar to other fluorescent agents, no CPL was observed for free εNAD+ in solutions [60]. In complexes with dehydrogenases (Table 4), εNAD+ shows pronounced positive gem values, which are almost independent of wavelength (variation of about 1×10−4 is close to error of 0.3×10−4 , which is characteristic of the SG-type instruments [19]) and equal to (4.0–4.5)×10−4 for four of five dehydrogenases studied. It was expected that the adenine subsites in their NAD+ -binding sites would be similar [60] and would provide similar induced asymmetry of εNAD+ in its excited electronic state. CPL of these complexes did not depend on the number of εNAD+ ligands attached to multimer apo-enzymes [60]. Rabbit muscle glyceraldehyde-3-phosphate dehydrogenase (GPDH) complex with single εNAD+ differs from other dehydrogenases exhibiting significant wavelength dependence on negative gem . The binding of the second εNAD+ led to flatting of the spectrum and lowering of gem |. The next two εNAD+ zeroes CPL across the

444

E. Gussakovsky Table 4 CPL of fluorescent cofactor complexes with protein pH

Excitation Emission (nm) (nm)

NADH NADH+IBA NADH

7.4

308 308 335

NADH+IBA

7.4

335

LDH

NADH

7.4

335

LDH GDH ADH SMDH GPDH

εNAD+ εNAD+ εNAD+ εNAD+ εNAD+ εNAD+

7.5 7.5 7.5 7.5 7.5 7.2

315 315 315 315 315 325

NADH PLP

7.2

GPB

355 328 365

HSA

Bilirubin

BSA

Bilirubin

Protein

Cofactor

LADH LADH

6.2 4.1 7.4 9.0 4–9

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