Applications of Raman spectroscopy in aqueous environments

February 1, 2007
Thomas Rades, Clare J. Strachan, Marja Savolainen, Anne Saupe, Niklas Sandler
Pharmaceutical Technology Europe
Volume 19, Issue 2

Raman spectroscopy has become a commonly used technique for physicochemical analysis that possesses many advantages over other analytical techniques. It is a very attractive characterization tool, not least because it enables measurements in water. However, very few examples of its application in an aqueous environment exist in literature. This paper provides some recent applications of Raman spectroscopy in pharmaceutical material and process characterization when water is present.

Different spectroscopic techniques are widely used to help understand the physical solid-state characteristics of drug substances and formulations — and Raman spectroscopy has become a commonly used technique that possesses many advantages over other analytical techniques.

The principles and applications of Raman spectroscopy have been recently well-addressed in this journal.1,2 Benefits of the technique are, for instance, that sample preparation is unnecessary or minimal; data acquisition is rapid and nondestructive; and measurements in aqueous environments are possible because water is a weak Raman scatterer (Figure 1). These benefits also allow bulk and final products to be tested directly in their packaging, such as glass bottles.3

Figure 1 Examples of Raman spectra of Indomethacin (a) and lactose monohydrate (b) measured from a powder and suspension. For comparison, a spectra of only water is also shown.

Numerous papers in literature emphasize that Raman spectroscopy is a very attractive characterization tool, specifically because it enables measurements in presence of water, but, surprisingly, few studies have applied Raman spectroscopy in an aqueous environment. This paper focuses on recent uses of Raman spectroscopy to characterize pharmaceutical materials and processes in water.

Probing critical solid-state processing steps and tablet dissolution

One of the earliest studies in a pharmaceutical setting demonstrating the use of Raman spectroscopy in crystallization was performed by Findlay and Bugay.4 They studied crystallization dynamics by measuring supersaturated menthol solutions with variable temperature (VT) Fourier Transform (FT)-Raman spectroscopy. Elkordy et al.5 reported the measurement of crystallized and spray-dried lysozyme, a model protein, by FT-Raman spectroscopy in 8% (w/v) solutions. They used a relatively high concentration to detect well-resolved Raman bands. After a water background had been subtracted, aqueous state FT-Raman spectra revealed protein conformation changes.

Raman spectroscopy has been particularly valuable for controlling wet granulation processes. Jørgensen et al.6 successfully employed charged coupled device (CCD) Raman and near infrared (NIR) spectroscopy to monitor polymorphic transitions of caffeine and theophylline during wet granulation. They concluded that the symmetric vibrations in the drug molecules could be readily detected and analysed with Raman spectroscopy. As Raman bands were much narrower than those in the NIR spectra (which has dominating OH bands) it was easier to observe the changes in the Raman spectra.

Wikström et al.7 successfully interfaced in-line Raman spectroscopy to a high-shear granulation process and also monitored pseudopolymorphic conversions of anhydrous theophylline. In a pelletization study, Raman spectroscopy, NIR spectroscopy and X-ray powder diffraction (XRPD) were used to characterize process-induced polymorphic changes.8 Samples were collected at the end of each processing stage (blending, granulation, extrusion, spheronization and drying). Raman and XRPD measurements confirmed the expected pseudopolymorphic changes of the active pharmaceutical ingredients (APIs) in the wet process stages. However, a relatively low Raman signal of theophylline (5% [w/w] in the formulation) complicated the interpretation.

In a recent study by Aaltonen et al.9 Raman spectroscopy was elegantly used to measure the solid state properties in situ during intrinsic dissolution measurements. They showed that the solid-state transformation from anhydrate to hydrate can be quantitatively measured for both theophylline and nitrofurantoin tablets using a Raman probe (Figure 2).

Figure 2 A schematic of the experimental setup used in a solid-state dissolution study. (Figure modified from reference 9.)

They concluded that by combining dissolved drug concentration with solid-state measurements, a deeper understanding of the solvent-mediated phase transformation phenomena during dissolution is achieved.

Controlling the properties of emulsions and suspensions

Islam et al.10 demonstrated the use of Raman to monitor a bench-scale formulation process of topical formulations. They heated the aqueous gel and mixed it with an oil phase to produce an emulsion, showing that a Raman shift of a peak at 925 cm-1 was linked to an ionic interaction between Carbopol and triethanol amine. The height of the 925 cm-1 peak relative to a band at 845 cm-1 increased with greater neutralization, as measured by the pH.

The authors, therefore, concluded that these peak height ratios can be used to control the amount of neutralizer used during manufacture. Raman spectroscopy in combination with stepwise linear regression has also been successfully used to confirm the polymorphic stability of benzimidazole in a complex suspension emulsion formulation.11 The formulation consisted of active drug suspended in oil droplets, which are emulsified in an aqueous matrix. No polymorphic transition was noted during the duration of the study.

However, a small sampling area within the heterogenous matrix caused some measurement variability. Thus, when measuring heterogenous samples multiple spectra should be recorded with a sufficiently large sample area to minimize variable and unrepresentative sample analysis.

The polymorphic composition of L-glutamic acid crystals has been monitored during batch cooling of saturated solutions using Raman spectroscopy in situ.12 By using peak height ratios, the α and β forms were quantified and at 25 °C in aqueous solution, the α form dominated. However, a solvent mediated transformation to the β polymorph followed, with the conversion accelerated at higher temperatures. This in situ quantification allows the optimization of both crystallization kinetics and polymorphic purity.

Raman spectroscopy offers an attractive possibility for direct and nondestructive quantitative measurements in an aqueous environment. Raman spectroscopy has been successfully used to determine the amount of medroxyprogesterone acetate in an aqueous suspension.13 Besides the fact that water does not interfere, an advantage of Raman spectroscopy is that it allows direct measurement of suspended drug concentrations, without the need to know the exact volume or density of the sample suspension, as would be the case with HPLC. Thus compared with HPLC, Raman spectroscopy provided a more reliable quantitation method for the amount of medroxyprogesterone acetate in a suspension.

Raman spectroscopy can also be used to detect changes in dissolved compounds. Li et al.14 used FT-Raman spectroscopy to examine the acid-base characteristics of various citrate buffer systems. The results indicated that the ratio of ionized and un-ionised species remained the same in the solution, the frozen solution and the amorphous state. Quantitation was based on the FT-Raman peak intensity ratio of the -COO to -COOH carbonyl stretching vibrations at 1400 cm-1 and 1720 cm-1, respectively. The signal to noise ratio was largest in the frozen samples, which was probably because of the shorter acquisition times that were necessary to prevent the samples melting.

Studying water sorption and hydration phenomena

Raman spectroscopy has been widely used to detect changes in the solid-state of a drug because of water sorption into the material. Airaksinen et al.15 used Raman spectroscopy to determine the effect of excipients on hydrate formation of theophylline during wet granulation. Despite the excipients in the formulation, Raman spectroscopy could still detect the theophylline anhydrate to monohydrate conversion without excipient interference.

A blue shift of the O=C-N bending band at 550 cm-1 and changes in the C=O stretch region at about 1700–1650 cm-1 were used to detect the hydrate formation. Raman spectroscopy has also been used in-line for quantitation of changes in the amorphous content of spray-dried amorphous lactose and trehalose that were stored in high relative humidity (RH [Figure 3]).16 The quantitation was based on multivariate analysis. Both of the amorphous sugars recrystallized during storage at 85% RH. Amorphous materials are very hygroscopic and absorb moisture when stored in high RH. Unlike with NIR, the absorbed water did not interfere with the Raman spectra and thus Raman spectroscopy was more suitable than NIR spectroscopy for the crystallinity quantitation.

Figure 3 Following the conversion of amorphoustrehalose to trehalose dihydrate at high relative humidity (85% RH) using in-line Raman spectroscopy; (a) illustrates the Raman spectra and (b) shows the increase in crystallinity determined by a PLS model. (Data modified from reference 16.)

Raman spectroscopy has been used to study the gelatinization and liquefaction (hydrolysis) process of starch.17 An aqueous starch suspension was heated above the gelatinization temperature and spectra recorded in-line. During gelatinization, the uptake of water into the crystalline starch caused inter- and intramolecular hydrogen bond breakage and new hydrogen bonds were formed with water molecules.

This could be observed in the Raman spectra because the bands at 1633 cm-1 and 3213 cm-1 increased while the others decreased. The band at 3213 cm-1 was attributed to hydrogen bonding with water and the band at 1633 cm-1 to decreased starch crystallinity. After gelatinization, the liquefaction was initiated by addition of α-amylase. Significant changes in the spectra were used to monitor the liquefaction. The formation of free hydroxyl groups because of the hydrolysis reaction was noted by the increase of the bands in the region of 1057 cm-1 and 1082 cm-1 (C-O-H bending).

In addition to solid-state transitions in aqueous environments, Raman spectroscopy can be used to study the molecular interaction between water and different substances. Raman spectroscopy has been used to determine the changes in the strength of hydrogen bonding between water and different pharmaceutical polymers.18 Raman spectroscopy could be used to study the hydrogen bonding as the studied polymers, PVP, PVAc and PVP/VA were unable to form intra- or intermolecular hydrogen bonds because of the lack of acidic protons. Therefore, any changes in the spectra of these polymers, when in solution, were as a result of hydrogen bonding with water molecules. However, the spectral changes were because of not only the changes in the hydrogen bonding, but also the plasticizing effect of water, which increased the molecular mobility. The increased molecular mobility affected the C-H and C-C vibrations. These groups form the backbone of the polymers.

Characterization of colloidal systems

In studies on nanostructured lipid carriers it has been shown that Raman spectra of lipids are sensitive to conformational, packing and dynamical changes involving hydrocarbon chains.19 Therefore, this technique can be applied to provide more detailed information on the microstructure of colloidal lipid dispersions such as nanoemulsions (NE), solid lipid nanoparticles (SLN) and nanostructured lipid carriers (NLC). Investigations on SLN based on glycerol behenate and NLC (mixtures of glycerol behenate and medium chain triglcyceride [MCT]) resulted in the conclusion that MCT loading did not lead to changes in order or packing behaviour of the lipid chains, meaning that oil incorporation could not be confirmed.20

When investigating the same systems, but using cetyl palmitate as solid lipid and MCT as oil component, Raman spectroscopy revealed that the NE was characterized by random coiled chains, while SLN and NLC formulations clearly showed sharp bands at 1128 cm-1 and 1062 cm-1 indicating the high conformational order of the acyl chains.

To determine whether the NLC-formulation spectrum is a simple mixture of spectra of SLN-formulation and nanoemulsion, a spectrum was calculated as a linear combination of 0.9 times the pure SLN spectrum plus 0.1 times the pure nanoemulsion spectrum. This calculated spectrum was compared with the NLC spectrum. The main peak positions did not change and subtraction of the normalized spectra revealed no major differences between the experimental and calculated spectra, suggesting that the SLN in the NLC particles is completely unperturbed by the presence of the oil.21


This paper has demonstrated the versatile use of Raman spectroscopy in pharmaceutical analysis. The technique offers a major advantage that spectra can be obtained noninvasively in the presence of water. However, the full potential of Raman spectroscopy analysis in aqueous environments is still to be realized, and it is envisaged that the number of application and studies in this area will grow rapidly in the near future.

Niklas Sandler is a senior scientist at AstraZeneca R&D (UK).

Marja Savolainen is a research scientist at the Division of Pharmaceutical Technology, Faculty of Pharmacy, University of Helsinki (Finland).

Anne Saupe is a research scientist at the School of Pharmacy, University of Otago (New Zealand).

Clare Strachan is chief research scientist at the Division of Pharmaceutical Technology, Faculty of Pharmacy, University of Helsinki (Finland).

Thomas Rades is a Professor at the School of Pharmacy, University of Otago (New Zealand).


1. S. Webster and K.J. Baldwin, Pharm. Technol. Eur., 17(8), 30–35 (2005).

2. S. Webster and K.J. Baldwin, Pharm. Technol. Eur., 17(6), 46–52 (2005).

3. T. Vankeirsbilck et al., Trends in Analytical Chemistry, 21(12), 869–877 (2002).

4. W.P. Findlay and D.E. Bugay, J. Pharm. Biomed. Anal., 16, 921–930 (1998).

5. A.A. Elkordy, R.T. Forbes and B.W. Barry, Int. J. Pharm., 278, 209–219 (2004).

6. A. Jørgensen et al., Pharm. Res., 19(9), 1285–1291 (2002).

7. H. Wikström, P.J. Marsac and L.S. Taylor, J. Pharm. Sci., 94(1), 209–219 (2005).

8. N. Sandler et al., AAPS PharmSciTech, 6(2), E174–E183 (2005).

9. J. Aaltonen et al., J. Pharm. Sci., in press (2006).

10. M.T. Islam et al., Pharm. Res., 21(10), 1844–1851 (2004).

11. B. De Spiegeleer et al., J. Pharm. Biomed. Anal., 39(1), 275–280 (2005).

12. T. Ono et al., Cryst. Growth Des., 4, 465–469 (2004).

13. T.R.M. De Beer et al., Eur. J. Pharm Sci., 23(4–5), 355–362.

14. J. Li et al., J. Pharm. Sci., 93(3), 697–712 (2004).

15. S. Airaksinen et al., J. Pharm. Sci., 92(3), 516–528 (2003).

16. M. Savolainen et al., J. Pharm. Pharmacol., in press (2006).

17. K.C. Schuster et al., Vibrational Spectroscopy, 22, 181–190 (2002).

18. L.S. Taylor, F.W. Langkilde and G. Zografi, J. Pharm. Sci., 90(7), 888–901 (2001).

19. S. Wartewig and R.H.H. Neubert, Advanced Drug Delivery Reviews, 57(8), 1144–1170 (2005).

20. K. Jores et al., Pharm. Res., 22(11), 1888–1897 (2005).

21. A. Saupe, K.C. Gordon and T. Rades, Int. J. Pharm., 314(1), 56–62 (2006).