Stopped-flow circular dichroism

As is apparent from previous chapters (Chapters 6, 8 and 9), understanding the kinetics of chemical and biological processes is extremely important. Questions we often consider, explicitly or implicitly, include: Has something happened ‘instantaneously’ or will it take 20 years? Does changing the conditions or available reagents affect either the end product or the rate of a process? What intermediates are produced during a reaction? Can we characterize any intermediates? Do we need to remove them to prevent side reactions? If some or all of the reactants or products are chiral, then circular dichroism (CD) detection may be the ideal tool for following the kinetics of a reaction, and if the half-life of the reaction is of the order of milliseconds to seconds or even minutes then stopped-flow mixing of the reagents will almost certainly be the appropriate choice of sample handling method. For reactions with half-lives of a few minutes to tens of minutes the reagents can be mixed by hand in a normal cuvette and the signal monitored at an appropriate wavelength. CD is not well suited to kinetics on timescales of hours due to the baseline drift that does occur (see Chapter 4, Section 2.5). Some CD spectropolarimeters have the useful facility of being able to perform a wavelength scan at pre-set intervals as well as monitoring continuously (except during the wavelength scan) at a chosen wavelength, thus facilitating the characterization of any intermediates. In this chapter we shall highlight some of the considerations of the stopped-flow technique that are particularly relevant to CD experiments. Particular problems may be encountered when performing CD (as opposed to other detection methods) stopped-flow experiments. The measured signals are very small (typical CD intensities are 0.1% or less of the absorbance signal), and the noise level observed is particularly sensitive to any inhomogeneities or turbulence in the samples. Also, as one of the main applications of stopped-flow CD is in the study of protein folding and unfolding, samples are often very viscous and/or corrosive, have significant absorbances due to buffers etc., and the experiments often require wide and variable mixing ratios.

Fluorescence principles and measurement

Fluorescence spectrometry is the most extensively used optical spectroscopic method in analytical measurement and scientific investigation. During the past five years more than 60000 scientific articles have been published in which fluorescence spectroscopy has been used. The large number of applications ranges from the analytical determination of trace metals in the environment to pH measurements in whole cells under physiological conditions. In the scientific research laboratory, fluorescence spectroscopy is being used or applied to study the fundamental physical processes of molecules; structure-function relationships and interactions of biomolecules such as proteins and nucleic acids; structures and activity within whole cells using such instrumentation as confocal microscopy; and DNA sequencing in genomic characterization. In analytical applications the use of fluorescence is dominant in clinical laboratories where fluorescence immunoassays have largely replaced radioimmunoassay techniques. There are two main reasons for this extensive use of fluorescence spectroscopy. Foremost is the high level of sensitivity and wide dynamic range that can be achieved. There are a large number of laboratories that have reported single molecule detection. Secondly, the instrumentation required is convenient and for most purposes can be purchased at a modest cost. While improvements and advances continue to be reported fluorescence instrumentation has reached a high level of maturity. A review of the physical principles of the fluorescence phenomenon permits one to understand the origins of the information content that fluorescence measurements can provide. A molecule absorbs electromagnetic radiation through a quantum mechanical process where the molecule is transformed from a ‘ground’ state to an ‘excited’ state. The energy of the absorbed photon of light corresponds to the energy difference between these two states. In the case of light in the ultraviolet and visible spectral range of 200 nm to 800 nm that corresponds to energies of 143 to 35.8 kcal mol-1. The absorption of light results in an electronic transition in the atom or molecule. In atoms this involves the promotion of an electron from an outer shell orbital to an empty orbital of higher energy.

Introduction to light absorption: visible and ultraviolet spectra

Light is a form of electromagnetic radiation, usually a mixture of waves having different wavelengths. The wavelength of light, expressed by the symbol λ, is defined as the distance between two crests (or troughs) of a wave, measured in the direction of its progression. The unit used is the nanometre (nm, 10-9 m). Light that the human eye can sense is called visible light. Each colour that we perceive corresponds to a certain wavelength band in the 400-700 nm region. Spectrophotometry in its biochemical applications is generally concerned with the ultraviolet (UV, 185-400 nm), visible (400-700 nm) and infrared (700-15 000 nm) regions of the electromagnetic radiation spectrum, the former two being most common in laboratory practice. The wavelength of light is inversely related to its energy (E), according to the equation: . . . E = ch/ λ . . . where c denotes the speed of light, and h is Planck’s constant. UV radiation, therefore, has greater energy than the visible, and visible radiation has greater energy than the infrared. Light of certain wavelengths can be selectively absorbed by a substance according to its molecular structure. Absorption of light energy occurs when the incident photon carries energy equal to the difference in energy between two allowed states of the valency electrons, the photon promoting the transition of an electron from the lower to the higher energy state. Thus biochemical spectrophotometry may be referred to as electronic absorption spectroscopy. The excited electrons afterwards lose energy by the process of heat radiation, and return to the initial ground state. An absorption spectrum is obtained by successively changing the wavelength of monochromatic light falling on the substance, and recording the change of light absorption. Spectra are presented by plotting the wavelengths (generally nm or μm) on the abscissa and the degree of absorption (transmittance or absorbance) on the ordinate. For more information on the theory of light absorption, see Brown (1) and Chapters 2, 3 and 4. The most widespread use of UV and visible spectroscopy in biochemistry is in the quantitative determination of absorbing species (chromophores), known as spectrophotometry.

The use of spectroscopic techniques in the study of DNA stability

Accurate determination of nucleic acid thermodynamics has become increasingly important in understanding biological function as well as applications in biotechnology and pharmaceuticals. Knowledge of the thermodynamics of DNA hybridization and secondary structure formation is necessary for understanding DNA replication fidelity (1), mismatch repair efficiency (2) and the mechanism of DNA triplet repeat diseases (3). In addition, RNA folding thermodynamics are an important aspect of understanding ribozyme catalysis, as well as understanding the regulation of protein expression, mRNA stability and the mechanism of protein synthesis by the ribosome (4). With the genome sequencing era upon us (5), it will increasingly become important to predict the folding and hybridization thermodynamics of DNA and RNA, so that accurate diagnostic tests for genetic and infectious diseases can be developed. Thus, there is a need to develop a database of accurate thermodynamic parameters for different nucleic acid folding motifs (4). This chapter describes practical aspects of the application of UV absorbance temperature profiles to determine the thermodynamics of nucleic acid structural transitions. Protocols and practical advice are presented for issues not normally addressed in the primary literature but that are crucial for the determination of reliable thermodynamics, such as sequence design, sample preparation, choice of buffer, protocols for determining strand concentrations and mixing strands, design of microvolume cuvettes and cell holder, instrumental requirements, data analysis methods, and sources of error. References to the primary literature and reviews are also provided where appropriate.

Use of optical spectroscopic methods to study the thermodynamic stability of proteins

The biophysical characterization of globular proteins will almost always include some type of study of the unfolding of protein to obtain thermodynamic parameters. The basic idea is that a transition between a native and unfolded state, induced by temperature, pH, or denaturant concentration, can serve as a standard reaction for obtaining a thermodynamic measure of the stability of the native state. For example, the free energy change for the unfolding reaction can be used to compare the stability of a set of mutant forms of a protein (1-4). This type of analysis is based both on assumptions of the thermodynamic model for the unfolding process and on assumptions in the way the data are analysed; some of these assumptions and their limitations will be discussed below. There are a variety of methods that can be used to monitor an unfolding process. A common method is differential scanning calorimetry, DSC, which measures the variation in the specific heat of a protein-containing solution as a protein is thermally unfolded (5-7). DSC is a popular method for this purpose, but optical methods can also provide suitable information for tracking the unfolding of a protein The spectroscopic signals for the native and unfolded states of a protein can give some insight regarding the structure of the states, and often can provide advantages of economy, ease of measurement and amenability to a wide range of sample concentration. The optical spectroscopic methods that have been used most often for this purpose are absorption spectroscopy, circular dichroism and fluorescence, which will be discussed in this chapter. A key to each of these methods and their use in protein unfolding studies is that the signal is a mole fraction weighted average of the signals of each thermodynamic state. That is, the observed signal, S, can be expressed as . . . S = ∑XiSi . . . . . . 1 . . . where Xi is the mole fraction of species i and si is the intrinsic signal of species i. In order for a particular spectroscopic signal to be useful for tracking a N ↔ U transition of a protein, the signal must be sufficiently different for the N and U states.

Steady-state kinetics

All of chemical kinetics is based on rate equations, but this is especially true of steady-state enzyme kinetics: in other applications a rate equation can be regarded as a differential equation that has to be integrated to give the function of real interest, whereas in steady-state enzyme kinetics it is used as it stands. Although the early enzymologists tried to follow the usual chemical practice of deriving equations that describe the state of reaction as a function of time there were too many complications, such as loss of enzyme activity, effects of accumulating product etc., for this to be a fruitful approach. Rapid progress only became possible when Michaelis and Menten (1) realized that most of the complications could be removed by extrapolating back to zero time and regarding the measured initial rate as the primary observation. Since then, of course, accumulating knowledge has made it possible to study time courses directly, and this has led to two additional subdisciplines of enzyme kinetics, transient-state kinetics, which deals with the time regime before a steady state is established, and progress-curve analysis, which deals with the slow approach to equilibrium during the steady-state phase. The former of these has achieved great importance but is regarded as more specialized. It is dealt with in later chapters of this book. Progress-curve analysis has never recovered the importance that it had at the beginning of the twentieth century. Nearly all steps that form parts of the mechanisms of enzyme-catalysed reactions involve reactions of a single molecule, in which case they typically follow first-order kinetics: . . . v = ka . . . . . . 1 . . . or they involve two molecules (usually but not necessarily different from one another) and typically follow second-order kinetics: . . . v = kab . . . . . . 2 . . . In both cases v represents the rate of reaction, and a and b are the concentrations of the molecules involved, and k is a rate constant. Because we shall be regarding the rate as a quantity in its own right it is not usual in steady-state kinetics to represent it as a derivative such as -da/dt.

Stopped-flow spectroscopy

There was a time, fortunately some years ago now, when to undertake rapid kinetic measurements using a stopped-flow spectrophotometer verged on the heroic. One needed to be armed with knowledge of amplifiers, light sources, oscilloscopes etc. and ideally one’s credibility was greatly enhanced were one to build one’s own instrument. Analysis of the data was similarly difficult. To obtain a single rate constant might involve a wide range of skills in addition to those required for the chemical/biochemical manipulation of the system and could easily include photography, developing prints and considerable mathematical agility. Now all this has changed and, from the point of view of the scientist attempting to solve problems through transient kinetic studies, a good thing too! Very high quality data can readily be obtained by anyone with a few hours training and the ability to use a mouse and ‘point and click’ programs. Excellent stopped -flow spectrophotometers can be bought which are reliable, stable, sensitive and which are controlled by computers able to signal-average and to analyse, in seconds, kinetic progress curves in a number of ways yielding rate constants, amplitudes, residuals and statistics. Because it is now so easy, from the technical point of view, to make measurement and to do so without an apprenticeship in kinetic methods, it becomes important to make sure that one collects data that are meaningful and open to sensible interpretation. There are a number of pitfalls to avoid. The emphasis of this article is, therefore, somewhat different to that written by Eccleston (1) in an earlier volume of this series. Less time will be spent on consideration of the hardware, although the general principles are given, but the focus will be on making sure that the data collected means what one thinks it means and then how to be sure one is extracting kinetic parameters from this in a sensible way. With the advent of powerful, fast computers it has now become possible to process very large data sets quickly and this has paved the way for the application of ‘rapid scan’ devices (usually, but not exclusively, diode arrays), which allow complete spectra to be collected at very short time intervals during a reaction.

Quantitative determination of equilibrium binding isotherms for multiple ligand-macromolecule interactions using spectroscopic methods

Thermodynamic studies provide information that is necessary in order to understand the forces that drive the formation of ligand-macromolecule complexes. Knowledge of the energetics of these interactions is also indispensable for characterization of functionally important structural changes that occur within the studied complexes. Quantitative examination of the equilibrium interactions are designed to provide the answers to the questions: What is the stoichiometry of the formed complexes? How strong or how specific are the interactions? Are there any cooperative interactions among the binding sites and/or the bound ligand molecules? Are the binding sites intrinsically heterogeneous? What are the molecular forces involved in the formation of the studied complexes, or, in other words, how do the equilibrium binding and kinetic parameters depend on solution variables (temperature, pressure, pH, salt concentration, etc.)? Equilibrium isotherms for the binding of a ligand to a macromolecule represent the relationship between the degree of ligand binding (moles of ligands bound per mole of a macromolecule) and the free ligand concentration. A true thermodynamic binding isotherm is model-independent and reflects only this relationship. Only then, when such an isotherm is obtained, can one proceed to extract physically meaningful interaction parameters that characterize the free energies of interaction. This is accomplished by comparing the experimental isotherms to theoretical predictions based on specific binding models that incorporate known molecular aspects, such as intrinsic binding constants, cooperativity parameters, allosteric equilibrium constants, discrete character of the binding sites or overlap of potential binding sites, etc. (see below). Any method used to quantitatively study ligand binding to a macromolecule must relate the extent of the complex formation to the free ligand concentration in solution. Numerous techniques have been developed to study equilibrium properties of specific and non-specific ligand-macromolecule interactions in which binding is directly monitored, including equilibrium dialysis, ultrafiltration, column chromatography, filter binding assay and gel electrophoresis (1-6). These direct methods are very straightforward; however, they are usually time consuming and some, like filter binding or gel shift assays, are non-equilibrium techniques which require many controls before the reliable equilibrium binding data can be obtained.

Spectrophotometry and fluorimetry of cellular compartments and Intracellular processes

Optical spectroscopy, Spectrophotometry and fluorimetry can be used to monitor processes occurring in living cells provided that suitable chromophores are present which ‘report’ on the events in which they participate. The advantages of optical techniques are manifold. Firstly they can be fast—with appropriate apparatus events in the pico- and nano-second domains can be studied by fluorescence spectroscopy. Secondly they are continuous—instant feedback from the experimental system can guide the most complex of experimental protocols, and allow the experimenter to adjust system parameters as necessary. Thirdly they are convenient, and most laboratories have access to equipment that can provide quantitative analysis of optical signals; examples include conventional spectrophotometers/fluorimeters, dedicated instruments (e.g. for fluorescence lifetime and polarization measurements), cameras, microscopes and plate readers. Significantly detectors from one apparatus can often be used on others to open up new experimental protocols. Fortunately the principles underlying the use of such a diverse array of optical devices are straightforward and universal—they apply just as much to laboratory ‘work-horse’ instruments as they do to the most specialized, laser-illuminated fluorescence microscope. The availability of fast laboratory computers with large storage capacities means that most modern spectrometers are microprocessor controlled and digitization of signals opens up the full range of possibilities of data accumulation, storage, analysis and interpretation. The main problem with optical measurements is not the acquisition but rather the interpretation of the data obtained. Straightforward analysis of the results depends on the clarity of the experimental design and the appropriate choice of chromophore. This chapter describes some of the problems that can be addressed by spectroscopic techniques and attempts to give guidance on good experimental design. Optical spectroscopy requires either spectrophotometers, to measure absorbance, fluorimeters, to measure fluorescence, or microscopes, which can measure fluorescence or absorbance of single cells or small groups of cells. Fluorimeters and spectrophotometers usually require solutions or suspensions of material in conventional cuvettes; microscopes provide two-dimensional images from smears, slices or surfaces. Other devices that record signals resolved in two-dimensions include gel scanners and microplate readers.

Stopped-flow fluorescence spectroscopy

Biochemical reactions, such as substrate or coenzyme binding to enzymes are usually completed in no more than 50-100 ms and thus require rapid reaction techniques such as stopped-flow instrumentation for their study. Fortunately, many such reactions can be followed by changes in the absorption properties of the substrate, product or coenzyme, and examples of these have been described in Chapters 1, 7 and 8. An alternative possibility is that during the reaction there is a change in the fluorescence properties of the substrate, coenzyme or the protein itself. Some reactions, particularly those involving the oxidation/ reduction of coenzymes, involve both changes in absorption and changes in fluorescence emission intensity. In many cases, the fluorescence properties of the ligand or protein itself may change when a complex is formed, even in the absence of a full catalytic reaction occurring, e.g. the protein fluorescence emission of most pyridine or flavin nucleotide-dependent dehydrogenases is quenched when NAD(P)H or FADH (respectively) binds to them, due to resonance energy transfer from the aromatic amino acids of the protein to the coenzyme. Conversely, the fluorescence emission from the reduced-coenzymes is usually enhanced on formation of the complex with these enzymes (1-3). The principles behind both fluorescence and stopped-flow techniques have been described in preceding chapters (2 and 8, respectively) and therefore readers should familiarize themselves with these chapters for some of the background information. In this chapter, we discuss the use of stopped-flow fluorescence spectroscopy and its application to a number of biochemical problems. A typical stopped-flow system is assembled from modular components of a conventional spectrophotometer/fluorimeter, a device permitting rapid mixing of the components of a reaction and a data recording system with a fast response. Commercially available instruments offer facilities for the observation of changes in absorption and/or fluorescence emission after rapid mixing of the reagents. These measurements can often be made simultaneously due to the different optical requirements of the two spectroscopic techniques. Figure 1 gives a generalized diagram of the geometry of a stopped-flow system able to simultaneously measure changes in absorption and fluorescence intensity of a reaction.