Biomacromolecular Applications of UV-Visible Absorption Spectroscopy

130 BIOMACROMOLECULAR APPLICATIONS OF UV-VISIBLE ABSORPTION SPECTROSCOPY

Spectrometry; Vibrational CD Spectrometers; Vibrational CD, Applications; Vibrational CD, Theory.

Further reading

Figure 14 The unfolding of xanthan as a function of potassium ion concentration. (') 4.3, (s) 15, (h) 30 and (d) 500 mM. Redrawn with permission from data in Norton IT, Goodall DM, Frangou SA, Morris ER and Rees DA (1984) J. Mol. Biol. 175: 371–394 Copyright (1984) Academic Press.

Theory; Induced Circular Dichroism; Macromolecule– Ligand Interactions Studied By NMR; Magnetic Circular Dichroism, Theory; Nucleic Acids and Nucleotides Studied Using Mass Spectrometry; Nucleic Acids Studied Using NMR; ORD and Polarimetry Instruments; Proteins Studied Using NMR Spectroscopy; Peptides and Proteins Studied Using Mass

Adler AJ, Greenfield NJ and Fasman GD (1973) Circular dichroism and optical rotatory dispersion of proteins and polypeptides. Methods in Enzymology 27 part D: 675–735. Breslauer KJ (1987) Extracting thermodynamic data from equilibrium melting curves for oligonucleotide orderdisorder transitions. Methods in Enzymology 259: 221–245. Eftink MR (1995) Use of multiple spectroscopic methods to monitor equilibrium unfolding of proteins. Methods in Enzymology 259: 487–512. Greenfield NJ (1996) Methods to estimate the conformation of proteins and polypeptides from circular dichroism data. Analytical Biochemistry 235: 1–10. Johnson WC Jr (1996) Determination of the conformation of nucleic acids by electronic CD. In: Fasman GD (ed) Circular Dichroism and the Conformational Analysis of Biomolecules, pp 433–468. New York and London: Plenum Press. Perrin JH and Hart PA (1970) Small molecule-macromolecule interactions as studied by optical rotatory dispersion-circular dichroism. Journal of Pharmaceutical Science 59: 431–448. Stevens ES (1996) Carbohydrates. In: Fasman GD (ed) Circular Dichroism and the Conformational Analysis of Biomolecules, pp 501–530. New York and London: Plenum Press. Woody RW (1995) Circular dichroism. Methods in Enzymology 246: 34–71.

Biomacromolecular Applications of UV-Visible Absorption Spectroscopy Alison Rodger and Karen Sanders, University of Warwick, Coventry, UK Copyright © 1999 Academic Press

Introduction Ultraviolet (UV) or visible radiation is absorbed by a molecule when the frequency of the light is at the

ELECTRONIC SPECTROSCOPY Applications

correct energy to cause the electrons of the molecule to rearrange (or become excited) to another, higherenergy, state of the system. Frequency, Q (measured in V –1), wavelength, O (usually measured in nm) and

BIOMACROMOLECULAR APPLICATIONS OF UV-VISIBLE ABSORPTION SPECTROSCOPY 131

energy, E (measured in J) are related by

where h = 6.626 196 × 10–34 J s is the Planck constant and c = 2.997 925 × 1017 nm s–1 is the speed of light in units to match the choice of nm for O. Absorption can be pictorially viewed as either the electric field or the magnetic field (or both) of the radiation pushing the electron density from a starting arrangement to a higher-energy final one. The direction of net linear displacement of charge is known as the polarization of the transition. The polarization and intensity of a transition are characterized by the so-called transition moment, which is a vectorial property having a well-defined direction (the transition polarization) within each molecule and a welldefined length (which is proportional to the square root of the absorbance). The transition moment may be regarded as an antenna by which the molecule absorbs light. Each transition thus has its own antenna and the maximum probability of absorbing light is obtained when the antenna and the electric field of the light are parallel. Absorbance is defined in terms of the intensity of incident, I0, and transmitted, I, light:

In most experiments we use the Beer–Lambert law to relate the absorption of light, A, to the sample concentration, C:

where l is the length of the sample through which the light passes and H is the molar absorption coefficient (extinction coefficient); if l is measured in cm and C in M = mol dm–3, then H has units of mol–1 dm3 cm–1. The Beer–Lambert law breaks down when the sample absorbs too high a percentage of the incident photons for the instrument to measure the emitted photons. An absorbance of 2, for example, means that 99% of the photons are absorbed. Biological samples present additional challenges to the Beer– Lambert law: if there are local high concentrations of sample (as in vesicles for example) or if molecules interact and perturb the spectroscopy of the isolated molecule, then the Beer–Lambert law becomes invalid.

The most common application of UV-visible absorption spectroscopy is to determine the concentration of a species in solution using the Beer–Lambert law. Other applications follow because the energy of UV-visible light is usually sufficient only to excite valence electrons which are the ones involved in bonding. Thus any UV-visible absorption spectrum is directly related to bonds and hence the structure of a molecule. The challenge is then to relate the plot of absorbance versus wavelength, which the spectrometer produces, to the structure of the molecules in the cuvette. With complicated systems, such as biological macromolecules and their complexes with small molecules, we usually interpret UV-visible spectra by considering changes in the spectrum as a function of a variable such as temperature, ionic strength, solvent, concentration, etc. Alternatively the absorption spectra data are used as input for interpreting other spectra such as fluorescence, circular dichroism (CD) or linear dichroism (LD).

Biological macromolecule structure and UV spectroscopy Proteins and DNAs are linear polymers where a limited set of residues are joined together by, respectively, the amide or phosphodiester bonds. The situation is similar for carbohydrates though the linking options are more varied. To a first approximation the absorbance spectrum expected for a biomolecule is therefore the sum of the spectra for the component parts. In the case of nucleic acids (DNA and RNA) the UV absorbance from 200–300 nm is due exclusively to transitions of the planar purine and pyrimidine bases (Figure 1). (The backbone begins to contribute at about 190 nm.) The accessible region of the spectrum (nitrogen purging is required below ~200 nm as oxygen absorption interferes with the spectrum) is therefore dominated by π → π* transitions of the bases. The UV spectra of the bases (Figure 2) look as if there are two simple bands; however, each ‘simple’ band observed is a composite of more than one transition. This makes detailed analysis of DNA absorption difficult, but usually ensures that the absorption spectrum changes when the system is perturbed. Thus absorption spectroscopy is a useful qualitative or empirical probe of structural changes. Typical DNA UV absorption spectra are illustrated in Figure 3. The base transitions are significantly perturbed by the so-called π–π stacking interactions and so both wavelength maxima and transition intensities vary depending on the base sequence and structure adopted (cf. Table 1).

132 BIOMACROMOLECULAR APPLICATIONS OF UV-VISIBLE ABSORPTION SPECTROSCOPY

Figure 1

Structural formula of DNA.

Figure 3 Absorbance spectra in 1 cm pathlength cuvettes of 190 µM poly[d(G-C)]2 (dotted line) and 240 µM poly[d(A-T)2] (dashed line).

Table 1 Long-wavelength absorbance maxima and extinction coefficients for some DNAs. Calf thymus DNA is ~60% A-T

DNA

Wavelength of Amax(nm) Hmax (mol–1 dm3 cm–1)

Calf thymus

260

6 600

Poly[d(A-T)]2

262

6 600

Poly(dA).poly(dT)

260

6 000

Poly[d(G-C)]2

254

8 400

Poly(dG).poly(dC)

253

7 400

(A = adenine, T = thymine, G = guanine and C = cytosine)

Figure 2 UV spectra of the DNA nucleotides: deoxyadenosine 5′-monophosphate (A), deoxyguanosine 5′-monophosphate (G), deoxycytidine 5′-monophosphate (C) and thymidine 5′-monophosphate (T). The spectrum of uracil is almost indistinguishable from that of thymine.

In the case of peptides and proteins the spectroscopy of the amide bonds, the side chains and any prosthetic groups (such as haems) determines the observed UV-visible absorption spectrum. However, as with DNA, intensities and wavelengths can be perturbed by the local environment of the groups. UV spectra of proteins are usually divided into the ‘near’ and ‘far’ UV regions. The near-UV in this context means 250–300 nm and is also described as the aromatic region, though transitions of disulfide bonds (cystines) also contribute to the total absorption intensity in this region. The far-UV (< 250 nm) is dominated by transitions of the peptide backbone of the protein, but transitions from some side chains also contribute to the spectrum below 250 nm.

The aromatic side chains, phenylalanine, tyrosine and tryptophan all have transitions in the near-UV region (Figure 4). At neutral pH, the indole of tryptophan has two or more transitions in the 240–290 nm region with total maximum extinction coefficient Hmax(279 nm) ~5000 mol–1 dm3 cm–1; tyrosine has one transition with Hmax(274 nm) ~1400 mol –1 dm3 cm–1; phenylalanine also has one transition with Hmax ~190 mol –1 dm3 cm–1; and a cystine disulfide bond absorbs from 250–270 nm with Hmax ~300 mol–1 dm3 cm–1. Although tryptophans have by far the most intense transitions, many proteins have few tryptophans compared with the other aromatic groups, so the near UV is not necessarily dominated by tryptophan transitions. The peptide chromophore (Figure 5) which gives rise to the transitions observed in the far-UV region (180–240 nm) has non-bonding electrons on the oxygen and also on the nitrogen atoms, π-electrons which are delocalized to some extent over the carbon, oxygen and nitrogen atoms, and σ bonding electrons. The lowest energy transition of the peptide

BIOMACROMOLECULAR APPLICATIONS OF UV-VISIBLE ABSORPTION SPECTROSCOPY 133

Figure 4 Aromatic absorption spectra of tryptophan, tyrosine and phenylalanine. Note the different concentrations required to ensure absorbance maxima of 1 absorbance unit.

In an α-helix, the coupling of the π → π* transition moments in each amide chromophore results in a component at about 208 nm which contributes to the characteristic α-helix CD spectrum. For UV-visible spectroscopy, however, the far-UV spectroscopy is usually of little use either for concentration determination or structural analysis as the accessible region (above 200 nm) is almost a linear plot of increasing intensity with decreasing wavelength. Carbohydrate UV-visible spectroscopy is essentially that of any substituents that have spectroscopy; a simple sugar system such as starch has no spectroscopy above 200 nm. Carbohydrates are typically derivatized with thiols or aromatic chromophores for UV spectroscopy. The spectroscopy of these compounds is largely determined by that of the derivatives. These data may be found in UV-visible spectroscopy atlases.

Wavelength scanning chromophore is an n → π* transition analogous to that in ketones, and the next transition is π → π*. As in the carbonyl case, the n → π* transition is predominantly of magnetic transition dipole character and is thus of low intensity (H ~100 mol–1 dm3 cm–1), though it is not as low as for a simple ketone; it occurs at about 210–230 nm (depending mainly upon the extent of hydrogen bonding of the oxygen lone pairs) and its small electric character is polarized more or less along the carbonyl bond. The π → π* transition (H ~7000 mol–1 dm3 cm–1) is dominated by the carbonyl π-bond and is also affected by the involvement of the nitrogen in the π orbitals; its electric dipole transition moment is polarized somewhere near the line between the oxygen and nitrogen atoms, and it is centred at 190 nm.

Figure 5

Simple wavelength scans of biological macromolecules

Nucleic acids (DNA and RNA), proteins and peptides absorb very little light above 300 nm in the absence of ligands or prosthetic groups with chromophores (absorbing units). However, it is usually wise to collect a simple UV scan of a sample from about 350 nm. If the spectrum is not flat between 350 and 310 nm then the sample has condensed into particles whose size is of the order of the wavelength of light; therefore what is being observed is scattering of the incident light rather than absorption. UV absorbance is most commonly used to determine the concentration of a sample and also to give an indication of its purity.

Schematic illustrations of (A) n → π* and (B) π → π* transitions of peptides.

134 BIOMACROMOLECULAR APPLICATIONS OF UV-VISIBLE ABSORPTION SPECTROSCOPY

To perform a wavelength scan proceeds as follows: 1. First choose the parameters. The wavelength range will usually be 350–200 nm, unless the buffer cuts out the low-wavelength end (see below). A data interval of 0.5–1 nm is adequate as the bands are all broad. Shorter data intervals will not improve the spectrum you plot but will fill up the computer memory. (If data is being collected to aid interpretation of LD spectra, ensure that the data interval is the same in both cases.) The bandwidth is the wavelength range of the radiation at any specified wavelength. If data is being collected every 0.5 nm, then a bandwidth of 0.5 nm is appropriate. A larger value will lead to a greater averaging of data than the data interval suggests. Do not choose a much smaller bandwidth as the incident light intensity will be reduced and the spectral noise will increase without improving the quality of your data. The data averaging time determines the signalto-noise ratio and affects the scan speed required to produce undistorted spectra. 0.033–0.1 s is usually a good choice; longer times are required if the absorbance is very small (less than 0.1 absorbance units) or small differences in absorbance are being determined (as for DNA melting curves; see below). 2. Choose the solvent or buffer for your experiment and fill a pair of matched cuvettes of the desired pathlength with it. Either switch off the baseline correction or use an air/air baseline in the instrument. Place one cuvette in the sample holder (usually the front position) and leave the reference holder empty. Run a spectrum over the desired wavelength range to check that the solvent/buffer spectrum is not significant over the wavelength range of interest. If it is then you need to change the solvent/buffer. In choosing your buffer note phosphate buffers are essentially spectroscopically invisible over the wavelength range usually used; however, you need to ensure that phosphate does not interact in any way with your sample. Cacodylate and ammonium acetate have a window down to ~210 nm and have the added advantage of preventing bacterial growth (whereas phosphate promotes it). Chloride ions begin absorbing just above 200 nm so high salt spectra cannot be collected at the lower-wavelength end of the spectrum. Higher buffer concentrations can be accommodated in shorter pathlength cuvettes (see

Beer–Lambert law discussion above), so if it is not possible to dilute or change the buffer try a smaller pathlength cuvette. (5 mm cuvettes will stand in a normal sample holder, 1 mm cuvettes will need spacers to hold them vertical – if they are not held vertical then you will be working with a variable pathlength. Smaller pathlength cuvettes will need special holders.) UV cut-off wavelengths for solvents are readily available in liquid chromatography texts. 3. Place the matched solvent/buffer cuvettes in both the reference holder (usually the rear position) and the sample holder and perform a baseline accumulation. An alternative to having the solvent/ buffer in the reference beam is to use a reference cell of water and collect a spectrum of the solvent/ buffer which is subsequently subtracted from all spectra. This method may be preferable if the solvent/buffer has a significant absorbance as it is easier to determine whether an apparent peak or dip in the spectrum is due to the buffer in some way. 4. Place the sample in the sample cuvette in the sample holder and record the spectrum. The absorbance at ~260 nm (or wherever the maximum is for a particular molecule) is generally used to determine nucleic acid concentrations using the Beer–Lambert law. As noted above, for DNA samples the linear relationship between concentration and absorbance seems to break down when the absorbance of a 1 cm pathlength solution exceeds ~1.5–2 absorbance units. Some DNA extinction coefficients are given in Table 1. Protein absorbances will be dominated by tryptophan residues (if there are any) and will have a maximum at 280 nm. The other aromatic residues also absorb at 280 nm. Absorbance at 280 nm may therefore be used to give an estimate of protein concentrations. At 280 nm a 1 mg cm–3 protein solution in a 1 cm pathlength cell often has an absorbance of ~1 absorbance unit. This is because many proteins have a similar percentage of ‘aromatic’ amino acid residues. However, the A280 (1 mg cm–3) can vary from 0.3 to 1.8. For example, the A280 (1 mg cm–3) for bovine serum albumin is ~0.66. In cases where the protein amino acid content and molecular weight are known then a reasonably accurate estimate of H can be made using the above H values for the residues (instead of assuming A280 = 1 for 1 mg cm–3) and then the Beer–Lambert law applied. The Beer–Lambert method for concentration determination of nucleic acids and proteins is based on the assumption that the samples are pure. Nucleic

BIOMACROMOLECULAR APPLICATIONS OF UV-VISIBLE ABSORPTION SPECTROSCOPY 135

acids, if present, will interfere with the protein concentration determination because they also absorb at 280 nm, and conversely. In these cases the following formula permits a rough estimate of protein concentration in the presence of nucleic acids (for a 1 cm pathlength cuvette):

It is also common practice to report the A280/A260 ratio as an indication of purity for a given sample. Wavelength scans of derivatized protein samples for concentration determination

Although proteins have different percentages of the different amino acids, each residue has an amide bond linking it to the next residue in the chain. A number of concentration-determination methods have thus been developed that involve derivatizing the amides and spectroscopically determining the concentration of the derivatives. The three methods mentioned below all rely on a standard of known concentration to enable a calibration curve to be plotted. The calibration is not necessarily linear. Biuret method This method is simple and reasonably specific as it depends on the reaction of copper(II) with four N atoms in the peptide bonds of proteins. Compounds containing peptide bonds give a characteristic purple colour when treated in alkaline solution with copper sulfate. This is termed the ‘biuret’ reaction because it is also given by the substance biuret NH2CONHCONH2. For a wide variety of proteins, 1.0 mg of protein in 2 cm3 of solution results in an absorbance at 540 nm of 0.1. Many haemoproteins, for example, give spurious results due to their intrinsic absorption at 540 nm, but modifications which overcome this difficulty are known (either removal of the haem before protein estimation or destruction of the haem by hydrogen peroxide treatment). The protein content of cell fractions such as nuclei and microsomes can be estimated by this method after solubilization by detergents such as deoxycholate or sodium dodecyl sulfate. The biuret reagent may be made by placing CuSO4.5H2O (1.5 g) and sodium potassium tartrate.4H2O (6.0 g) into a dry 1 dm 3 volumetric flask and adding about 500 cm3 of water. With constant swirling, NaOH solution (300 cm 3, 10% w/v) is added and the solution made to volume (1 dm3) with water. The reagent prepared in this manner is a deep blue colour. It may be stored indefinitely if KI

(1 g) is also added and the reagent is kept in a plastic container. The protein solution (x cm3, where x < 1.5) is then mixed with water [(1.5 – x) cm3] to make a total volume of 1.5 cm3 to which 1.5 cm3 of biuret reagent is added. The purple colour is developed by incubating for 20 min at 37°C. The tubes must then be cooled rapidly to room temperature and the absorbance at 540 nm determined. The colour of the solution is stable for hours. Folin–Ciocalteau or Lowry method While the biuret method is sensitive in the range 0.5 to 2.5 mg protein per assay, the Lowry method is 1 to 2 orders of magnitude more sensitive (5 to 150 µg). The main disadvantage of the Lowry method is the number of interfering substances; these include ammonium sulfate, thiol reagents, sucrose, EDTA, Tris, and Triton X-100. The final colour in the Lowry method is a result of two reactions. The first is a small contribution from the biuret reaction of protein with copper ions in alkali solution. The second results from peptide-bound copper ions facilitating the reduction of the phosphomolybdic-tungstic acid (the Folin reagent) which gives rise to a number of reduced species with a characteristic blue colour. The amino acid residues which are involved in the reaction are tryptophan and tyrosine as well as cysteine, cystine and histidine. The amount of colour produced varies slightly with different proteins. In this respect it is a less-reliable assay than the biuret method, but it is more reliable than the absorbance method since A280 may include contribution from other species, and also the absorption of a given residue is dependent on its environment within the protein. Two solutions are required for the Lowry method. For the alkaline copper solution, mix 50 cm3 Na2CO3 (2% w/v) in NaOH (0.1 M) with 1 cm3 of CuSO4.5H2O (0.5% w/v) and 1 cm3 of sodium potassium tartrate (1% w/v). This solution must be discarded after 1 day. The Folin reagent (phosphomolybdic-tungstic acid) may be made by diluting the concentrated Folin reagent obtained from e.g. Sigma with an equal volume of water so that it is 1 N (i.e. 1 M H+). To perform an assay add x cm3 of sample (where x < 0.6) containing 5–100 µg of protein as required to (0.6 – x) cm3 of water. Then add 3 cm 3 of the alkaline copper solution. The solutions must then be mixed well and allowed to stand for 10 min at room temperature. 3.0 cm 3 of Folin reagent is then added and after 30 min the absorbance at 600 nm is determined. Coomassie blue dye binding assay This proteindetermination method involves the binding of

136 BIOMACROMOLECULAR APPLICATIONS OF UV-VISIBLE ABSORPTION SPECTROSCOPY

Coomassie brilliant blue G-250 to protein. The protonated form of Coomassie blue is a pale orange-red colour whereas the unprotonated form (Figure 6) is blue. When proteins bind to Coomassie blue in acid solution their positive charges suppress the protonation and a blue colour results. The binding of the dye to a protein causes a shift in the absorption maximum of the dye from 465 to 595 nm and it is the increase in absorbance at 595 nm that is monitored. The assay is very reproducible and rapid with the dye binding process virtually complete in ∼ 2 min with good colour stability. The reagent is prepared as follows. Coomassie brilliant blue G-250 (100 mg) is dissolved in 50 cm3 95% ethanol. To this solution phosphoric acid (100 cm3, 85% w/v) is added and the solution diluted to 1 dm3. To perform the assay, x cm3 of the sample containing 5–100 µg of protein is placed in a clean, dry test tube. (0.5 – x) cm3 water and 5.0 cm3 of diluted dye reagent are added and the solution mixed well. After a period of from 5–60 min, A595 is determined. The only compounds found to give excess interfering colour in the assay are relatively large amounts of detergents such as sodium dodecyl sulfate, Triton X-100 and commercial glassware detergents. Interference by small amounts of detergent may be eliminated by the use of proper controls. The assay is non-linear and requires a standard curve.

binds to the DNA then the spectrum of the complex will be different from the sum spectrum. One should note that the observed spectrum is probably a complicated mixture of that due to bound and unbound ligand and free and complexed DNA. When a planar aromatic molecule binds intercalatively (sandwiched between two base pairs) to DNA there is usually a characteristic decrease in the ligand-absorbance signal (this can be up to 50%) and a shift to the red (bathochromic shift) of between ∼ 2 nm and 20 nm as illustrated in Figure 7. The DNA spectrum is also affected by any molecule such as an intercalator that causes a structural change. This makes such spectra a useful probe of DNA/drug interactions but renders absorbance useless for concentration determinations unless the perturbed extinction coefficients are known.

Simple wavelength scans of macromolecules with bound ligands

When a ligand is added to, for example, a DNA solution, if it does not bind to the DNA then the UVvisible spectrum will simply be the sum of the DNA spectrum and the ligand spectrum. If the ligand

Figure 6

Coomassie blue.

Figure 7 (A) 5 µM anthracene-9-carbonyl-N 1-spermine in water. (B) 2 µM, 4 µM, 5 µM, 7 µM, 10 µM and 13 µM anthracene-9-carbonyl-N 1-spermine in water with 200 µM calf thymus DNA. Note broadening and magnitude decrease of 250 nm band absorbance; this molecule intercalates between DNA base pairs.

BIOMACROMOLECULAR APPLICATIONS OF UV-VISIBLE ABSORPTION SPECTROSCOPY 137

Titrations

The label ‘titration’ is used to cover experiments where spectra are collected as a function of concentration, ionic strength, pH, etc. To minimize macromolecule consumption and also (perhaps surprisingly) to minimize concentration errors, the best method is often to add solution to the cuvette. A simple way to avoid dilution effects is to proceed as follows. Consider a starting sample that has concentration x M of species X. Each time y cm3 of Y is added, also add y cm3 of a 2x M solution of X. The concentration of X remains constant at x M. Many variants on this theme may be derived. If a titration series where ligand concentration is held constant while the macromolecule varies has a constant absorbance at a wavelength, Oi, where the ligand absorbs this is called an isosbestic point. Isosbestic points only occur if two species are present in solution and those species have the same absorbance at Oi.

case Stot = n′[M] where n′ is the number of binding sites per protein.) Then

where cb is the concentration of the bound ligand and cf is the concentration of the free ligand. There are a number of methods for determining K using absorbance data. The simplest is the enhancement method. This method is commonly used for fluorescence spectroscopy and may also be used to interpret absorbance data. We write

Binding constants

If the spectral shape of a ligand spectrum remains unchanged during a titration experiment, but the magnitude changes in a manner that is proportional to the concentration of bound ligand then the spectral data can be used to determine the equilibrium binding constant, K. This is because in such a case the ligands are binding in one binding mode or in constant proportions in more than one mode (meaning site, orientation, sequence, etc). The data must be of very high quality for absorbance (or any other spectroscopic data) to be used to determine K. A simple plot of change in absorbance versus either macromolecule or ligand concentration (whichever is being varied) will probably enable the quality of the data set to be determined. It should be a smooth curve. Consider the equilibrium

where Lf is a free ligand, Lb is a bound ligand and Sf is a free site. In the simple case where the macromolecule can be treated as a series of binding sites of n residues in size, then the total site concentration Stot = [M]/n, where [M] is the residue concentration of the macromolecule. (For proteins it is sometimes preferable to think in terms of the concentration of molecules rather than residues. In this

Application of this equation requires knowledge of the absorbance of free and bound ligand. Determining the latter requires measuring an absorbance spectrum under conditions where it is known that all the ligand is bound to the macromolecules. K may then be determined directly. A more accurate value of K will be achieved if the data is used to perform a Scatchard plot. The Scatchard plot is based on rewriting the equation for the equilibrium constant as:

where

So, a plot of r/cf versus r has slope –K and y-intercept K/n. The x-intercept occurs where r = n. Other methods more commonly used with CD or LD data may be used with normal absorption data if the change in absorbance (the absorbance of the DNA/ligand system minus the absorbance of a freeligand solution of the same ligand concentration) is used in the analysis.

138 BIOMACROMOLECULAR APPLICATIONS OF UV-VISIBLE ABSORPTION SPECTROSCOPY

Macromolecule condensation

A UV spectrum may be used to follow the condensation of a macromolecule sample into particles, though, as discussed above, what is being probed is really scattering of the light rather than its absorbance. A monotonic increase in absorbance is observed above 300 nm as condensation takes place. In the case of DNA, the addition of a highly charged DNA binding ligand (such as spermine or [Co(NH3)6]3+) will effect this change. Concomitantly with the increase in absorbance signal above 300 nm, a decrease in the 260 nm DNA absorbance is observed.

DNA melting curves: absorption as a function of temperature If there were no residue–residue interactions in a biomolecule then the UV spectrum would be independent of its geometry and the absorption spectrum would simply be the sum of the contributions from the residues are discussed above. This is particularly true for DNA, where π–π stacking interactions lower the magnitude of the absorbance at 260 nm (the hypochromic effect) and change it at most other wavelengths. The extent of this change depends on the DNA structure. In principle if the DNA is heated enough to disrupt all base structure then the spectrum would become the sum of the base spectra in appropriate proportions. A high enough temperature to achieve this cannot usually be reached in water; however, we can use the change in the UV absorbance signal at a chosen wavelength (usually 260 nm, though at ∼ 280 nm A-T base pairs show very little change in absorbance so this wavelength may be used to probe the role of G-C relative to A-T base pairs) to follow the disruption of base stacking and hence also base-pair hydrogen bonding. The data from such an experiment is usually illustrated as a melting curve or a derivative melting curve and summarized by the so-called melting temperature, Tm (e.g. Figure 8). Tm is the temperature where the absorbance is the average of the duplex and single-stranded DNA absorbances where 50% of the DNA has melted. Thermodynamic data relating to the stability of the duplex may also be extracted from melting curves.

List of symbols A = absorbance; cb,f = ligand transmitted light intensity;

concentration; I = Io = incident light

Figure 8 UV melting curves for 200 µM calf thymus DNA in 10 mM salt (denoted DNA), and also with 20 µM spermine and 20 µM anthracene-9-carbonyl-N 1-spermine (denoted anthsp). Note the premelting transition with anthracene-9-carbonyl-N 1spermine.

intensity; l = sample light path length; H = molar absorption coefficient; O = wavelength of radiation (usually in nm); Q = frequency of radiation (s–1). See also: Biomacromolecular Applications of Circular Dichroism and ORD; Dyes and Indicators, Uses of UV-Visible Absorption Spectroscopy; Macromolecule–Ligand Interactions Studied By NMR; Nucleic Acids and Nucleotides Studied Using Mass Spectrometry; Nucleic Acids Studied Using NMR; Peptides and Proteins Studied Using Mass Spectrometry; Proteins Studied Using NMR Spectroscopy.

Further reading Atkins PW (1983) Molecular Quantum Mechanics. Oxford: Oxford University Press. Atkins PW (1991) Physical Chemistry, 4th edn. Oxford: Oxford University Press. Bradford MM (1976) Analytical Biochemistry 72: 248. Craig DP and Thirunamachandran T (1984) Molecular Quantum Electrodynamics: An Introduction to Radiation-Molecule Interaction. London: Academic Press. Eriksson S, Kim SK, Kubista M and Nordén B (1993) Biochemistry 32: 2987. Gornall AG, Bardawils CJ and David MM (1949) Determination of serum proteins by means of the biuret reagent. Journal of Biological Chemistry 177: 751–766. Hiort C, Nordén B and Rodger A (1990) Enantioselective DNA binding of [Ru(1,10-phenanthroline) 3]2+ studied with linear dichroism. Journal of the American Chemical Society 112: 1971. Hollas JM (1992) Modern Spectroscopy, 2nd edn. Chichester: John Wiley and Sons. Legler G et al (1985) Analytical Biochemistry 150: 278.

BIOMEDICAL APPLICATIONS OF ATOMIC SPECTROSCOPY 139

Marky LA and Breslauer KJ (1987) Calculating thermodynamic data for transitions of any molecularity from equilibrium melting curves. Biopolymers 26: 1601–1620. Michl J and Thulstrup EW (1986) Spectroscopy with Polarized Light. New York: VCH. Newbury SF, McClellan JA and Rodger A (1996) Spectroscopic and thermodynamic studies of conformational changes in long, natural mRNA molecules. Analytical Communications 33: 117–122. Puglisi JD and Tinoco IJ (1989) Absorbance melting curves of RNA. Methods in Enzymology 180: 304– 325. Read SM and Northcliffe DH (1981) Analytical Biochemistry 96: 53.

Rodger A (1993) Linear dichroism. Methods in Enzymology 226: 232–258. Rodger A and Nordén B (1997) Circular Dichroism and Linear Dichroism. Oxford: Oxford University Press. Rodger A, Blagbrough IS, Adlam G and Carpenter ML (1994) DNA binding of a spermine derivative: spectroscopic studies of anthracene-9-carbonyl-N1-spermine with poly(dG-dC)2 and poly(dA-dT)2. Biopolymers 34: 1583–1593. Rodger A, Taylor S, Adlam G, Blagbrough IS and Haworth IS (1995) Multiple DNA binding modes of anthracene-9-carbonyl-N1-spermine. Bioorganic and Medicinal Chemistry 3: 861–872. UV-VIS Atlas of Organic Compounds (1992) 2nd edn. Weinheim: VCH.

Biomedical Applications of Atomic Spectroscopy Andrew Taylor, Royal Surrrey County Hospital and University of Surrey, Guildford, UK Copyright © 1999 Academic Press

The techniques of flame, electrothermal and vapour generation atomic absorption, flame and inductively coupled plasma atomic emission and inductively coupled plasma mass spectrometry for the measurement of minerals and trace elements in biological specimens are described. Situations in which each of the techniques might be employed for the analysis of biomedical samples are reviewed. Interferences associated with the types of samples typically examined are mentioned with accounts of how these are removed in regular practice, either during the sample preparation or by features of the instrumentation. The advantages and disadvantages of these techniques are given with particular reference to sensitivity, single to multielement measurements and special applications such as the determination of stable isotopes. While total analyte concentrations are typically determined, examples of how speciation may be of interest are included. It is seen that for biomedical measurements, atomic spectroscopic techniques are complementary and that each may be appropriate for particular sample types or applications. Except for a few very special purposes these are the techniques of choice for measurement of minerals and trace elements in biomedical specimens. Quantitative analytical techniques included under the general heading of atomic spectroscopy are almost always employed for the direct determination of inorganic elements. For a few biomedical applications the specimen preparation gives indirect

ATOMIC SPECTROSCOPY Applications measurements of molecular compounds, generally by using a metal-based reagent in a separation or extraction step and measurement of the metal as a surrogate for the analyte of interest. The few examples of indirect measurements are included in the extensive reviews of atomic spectrometry published annually as the Atomic Spectrometry Updates (ASU). As described in the articles on ‘Theory’ and ‘Methods & Instrumentation’ in this Encyclopedia, analytical atomic spectroscopy concerns measurements of about 60 elements, generally metals, although consideration of mass spectrometry extends the range to most other elements. Biomedical applications require the measurement of almost all these elements, which are often loosely termed the minerals and trace elements. As given in Table 1, biological specimens contain various bulk elements which fulfill essential structural and functional roles. The trace elements, which individually account for less than 0.01% of the dry weight of the organism, are represented by a series of elements essential to health, development and well-being (Table 2) and others such as lead, present as contaminants from the environment. With sufficiently sensitive analytical techniques almost all elements of the periodic table can be found and are included among the trace elements. Figure 1 demonstrates that minerals and trace elements are relevant to a large number of disciplines which may be regarded as ‘biomedical’. Some of