NEWS & REVIEW

1. Editorial

I first ran into Bob "Jake" Jacobsen in the mid 1960's when I was a Post-doc with the late Ellis Lippincott at the University of Maryland in College Park. I was there for the summers of 1964 and 1966 and met up with many wonderful people including Peter Klaboe, Derek Steele and Linda and Malcolm Whatley. I got to know Jake at the Maryland Vibrational Spectroscopy Summer Schools and at Conferences. I was immediately drawn to his polite, easy-going manner combined with first-class ability. Jake was one of the most pleasant gentlemen I have ever met - I mean it, a real gentleman in the best sense of the word.

In September 1966, I returned from the States and two weeks later married my lady wife Janette. Jake and Bill Fateley were in London at the time and came to the festivities.

You will notice that Bill looks a good deal younger than he does now - the Bridegroom has hardly changed! Patrick - How have you managed to keep your svelte figure? - Louise[who would also like to point out that she was only 18 months old at the time!]

I kept up with Jake when I visited the States  and last worked with him a few years ago in Seattle where we both lectured on the vibrational spectroscopy of bio-systems - Jake covered the use of mid infrared and I spoke about Raman.

Jake and Bill somehow acquired a slide of our wedding photograph and for years whenever one of them turned up at a Conference at which I was a speaker, a rogue slide would mysteriously get included in my set. Usually in the MIDDLE! Solution - use overheads!

About a year ago, we decided to dedicate an Edition of IJVS to the memory of Dr Bob Jacobsen and to highlight the work in which Jake was most involved in the latter part of his career - the use of infrared spectroscopy in studying biological systems. To my delight Kai Griebenow agreed to edit this important edition. Kai's review is quite honestly, in a class of its own - please read it - you will be sampling, I am sure, a keynote article.

Editor


It is my pleasure to introduce this special memorial issue to Dr Bob Jakobsen of the Internet Journal of Vibrational Spectroscopy at the end of the year 1999 and on the brink of a new century.  Quite amazing are the advantages that have appeared in vibrational spectroscopy, particularly in the last 20 years when Fourier-techniques have been applied to spectral acquisition and data analysis.  'Jake', who this issue is being dedicated to, has contributed significantly to this process.  The regular editor of IJVS, Patrick Hendra, has taken on the task of describing these contributions.

I do not consider myself a vibrational spectroscopist.  The truth is that I am a scientist who uses whatever method is available to characterize biological systems.   However, many of my scientific works have made extensive use of the power of Raman- and FTIR-spectroscopy.

Based on this, I started thinking about what makes both spectroscopic methods so appealing in the characterization of complex biological systems.  One possible answer is that both can be applied under a variety of conditions that are not easily accessible by other spectroscopic methods.  Secondly, they can be applied non-invasively.  This is what this issue focuses on.

Sockalingum et al. describe the application of surface enhanced FT-Raman spectroscopy (SERS) to study the interactions of drug and target molecules in living cells.  Novel concepts are described in this exciting paper (both instrumentally and conceptually) allowing insight into drug-target interactions with tremendous potential in the area of drug development and testing.

My own group contribute a mini review on the use of FTIR spectroscopy in the development of devices for the sustained release of pharmaceutical proteins from biocompatible polymers.  The strength of the spectroscopic method consisted here in the possibility to investigate samples in very different states (amorphous protein powders, suspensions, and dispersions in the polymer matrix) in combination with the non-invasive character.

The other three contributed articles in this issue use FTIR spectroscopy.  Gao and Ci describe the use of FTIR spectroscopy to characterize breast tissue samples.  They found significant spectral differences between benign and carcinoma tissue.  The findings may become important in the future to further the diagnosis of the disease on the biochemical rather than morphological level.

Sertsou, Agatonovic-Kustrin and Rades offer a paper in pharmaceutical analysis field and in Section 4 Nagasaki and co-workers describe a fascinating application of computer power to FTIR instruments.

I hope you find pleasure in reading the above-mentioned articles, which provide a small insight into the growing world of vibrational spectroscopy applications in biological systems.

Kai Griebenow

Feature Review

2. Application of FTIR spectroscopy
to probe and improve protein structure
in sustained release devices.

Kai Griebenow*,
Ingrid Castellanos and Karen G. Carrasquillo 

* Corresponding author, e-mail griebeno@adam.uprr.pr

Introduction

In our contribution published previously in the IJVS (Griebenow et al., 1999) we have extensively described the application of FTIR spectroscopy to characterize dehydration-induced structural changes in proteins. Included were a brief introduction of the theoretical background, practical considerations of how to determine protein secondary structure from IR spectra, and some examples showing what results can be obtained employing this spectroscopic technique. In this overview article we would like to extent the last section of our prior review and focus on relevant examples of recent applications of FTIR spectroscopy in an important area with medical and biotechnological applications. All examples have in common that protein structure is being investigated under conditions not easily approachable by other techniques. Such conditions include (but are not limited to) suspensions of dehydrated protein powders in organic solvents and dispersions of protein powders in polymers. We will outline how FTIR spectroscopy can be employed in a rational optimization of a modern drug delivery system. We named this approach structure-guided protein encapsulation.

Pharmaceutical proteins and their delivery

Many biopharmaceutical drugs, such as peptides and proteins, currently enter or are in clinical trials. This is largely due to recent advances in biotechnology allowing the mass-production of recombinant proteins [1]. Some examples of promising new protein drugs include insulin-like growth factor for the treatment of juvenile-onset insulin-dependent diabetes [2], tumor-derived heat shock proteins for immunotherapy of tumors [3], and antibodies in cancer treatment [4]. The list of already FDA approved recombinant protein pharmaceuticals is steadily growing [5]. Applications of peptide and protein drugs include hormone treatment (e.g., insulin, growth hormone), immuno- and tumor-treatment (interferon, antibodies, interleukins), cardiovascular and thromobolytic treatment (e.g., tissue plasminogen activator, urokinase), and immunization (e.g., immunoglobulins, antitoxins) [6].

However, the use of proteins as biopharmaceuticals is hampered by the fact that patient-convenient routes of administration (e.g., orally) frequently cannot be used due to the susceptibility of proteins to biological degradation pathways, such as proteolysis. Therefore, most proteins are delivered via parenteral routes to the patient [6]. However even when delivered by parenteral (e.g., insulin) or pulmonary routes (e.g., recombinant human deoxyribonuclease I) proteins often possess low biological halftimes and frequent administrations are required. For example, the half-life of interferon alpha, beta, and gamma varies between 25 minutes and 16 hours in humans [6].

A possible and exciting solution to such problems is the controlled delivery of proteins from biocompatible polymers [7-16]. Since it has been established that large molecules including proteins can be delivered slowly and continuously from biocompatible polymers [17], the sustained release of proteins and peptides field from such polymers has grown immensely [18]. These systems provide many advantages over conventional therapeutic approaches, such as intravenous injection. For example, they can be constructed to deliver their active component at a constant rate for prolonged periods, they can target specific tissues, extend the half-life of the drug, and also enhance its in vivo stability [13,18-20]. In addition, patient compliance and comfort, as well as control over blood levels may be improved with the development of sustained release protein injectables, since regular invasive doses can be avoided [18]. Of tremendous value for medical and humanitarian reasons would also be the development of "one shot" immunization [21,22], e.g., by the sustained release of tetanus toxoid [13,19]. Particularly exciting are developments in single-administration vaccines against HIV-1 infection based on the envelope glycoprotein gp120 (currently in clinical trials) [21]. Success in such endeavors would in particular boost efforts by the WHO to achieve high levels of vaccination in developing countries where frequent medical attendance is still a very serious problem [21].

Protein encapsulation for their sustained release

Even though there is a tremendous potential in delivering proteins from biocompatible polymer devices, there are also tremendous problems involved in the encapsulation process. Proteins have a very fragile three-dimensional structure [23-25] and the protein encapsulation process was believed to cause significant protein structural perturbations [18,26]. Such structural perturbations can lead to the formation of irreversible protein aggregates. Aggregate formation not only leads to a loss of expensive pharmaceutical protein, but also can have severe consequences for the patient. First, protein aggregation adds a hard-to-control variable to the precise prediction of the release profile for the drug, which in turn may prove fatal to the patient. Furthermore, protein aggregates can be immunogenic and possibly lead to shock and death of patients. An ultimate goal in the encapsulation of proteins for their sustained release is therefore to assure the preservation of their native three-dimensional structure upon encapsulation and subsequent delivery to avoid formation of protein aggregates. How challenging this requirement is will be explained following a typical protocol used for the encapsulation of proteins. Figure 1 shows the scheme of encapsulation of a protein by the so-called double-emulsion/solvent-evaporation (a.k.a., water-in-oil-in-water or w/o/w) technique. This procedure is by far the most frequently used encapsulation method [18].

In the w/o/w technique, an aqueous protein solution is introduced into an organic solvent. This solvent is needed to dissolve the hydrophobic polymer [mostly poly(lactic-co-glycolic) acid, PLGA] and is typically methylene chloride because of its excellent properties (good polymer solubility and easy to evaporate) or sometimes ethyl acetate. The first emulsion is formed using a suitable method, e.g., probe sonication or homogenization. During this process a large area of solvent-water interface is formed. It is believed that proteins adsorb into this interface due to their surface activity [18]. Hydrophobic interfaces are known to destabilize the native structure of proteins and this can lead to formation of protein aggregates [18]. In addition, the method of emulsification poses stress upon the protein. After formation of the first emulsion, a second one is formed by introducing the first emulsion into an aqueous solution containing an emulsifier, e.g., polyvinyl alcohol. Again, a homogenization method has to be used. This is next followed by a period of solvent evaporation leading to the hardening of the polymer, typically by stirring for some hours. As a result, the protein solution is trapped into the relative hydrophobic matrix of the PLGA polymer forming small inclusions. Some dehydration is likely to occur due to the solubility of water in the organic solvent (methylene chloride can contain up to 2% of water). Finally, the microspheres formed are collected, washed with distilled water (repeated centrifugation or filtration), followed by dehydration, typically by lyophilization. The final step again imposes significant stress upon the protein because lyophilization causes significant protein structural perturbations (see our previous review in IJVS, Griebenow et al., 1999). In summary, the list of stresses resulting in the destabilization of the three-dimensional structure of proteins is long and includes events such as interfacial-, solvent-, mechanically-, and dehydration-induced protein denaturation. The resulting PLGA-microspheres loaded with protein are shown in Figure 2.

In the context of this long list of events involved in the encapsulation procedure that could indeed impact upon the structure of proteins and also the reports that abound on the incomplete delivery of proteins from such devices, an urgent goal identified in the literature is to learn how the encapsulation procedure affects protein structure. Until very recently NO information was available on the structure of proteins in such microspheres simply because there was no suitable method that could be employed to study them.

Now, thanks to recent instrumental and conceptual advances in the area of FTIR spectroscopy, (Griebenow et al., 1999) this has changed. As a result, the structure of a protein dispersed as an amorphous powder in a hydrophobic matrix such as PLGA has been determined for the first time by Carrasquillo et al. [27]. In parallel, the secondary structure of lysozyme and BSA was determined for the first time within PLGA microspheres by Fu et al. [28]. The outcome of these and related investigations and their impact on protein formulation for their sustained delivery from biocompatible polymers will be described in this article.

Protein structure in PLGA microspheres created by the w/o/w technique

Fu et al. [28] encapsulated the two model proteins - hen egg-white lysozyme and bovine serum albumin (BSA) in PLGA microspheres following a typical protocol involving the w/o/w technique. The structure of the proteins encapsulated in the microspheres was determined by FTIR spectroscopy and the a-helix content was used as the main structural parameter. For both proteins, encapsulation was found to severely impact the protein secondary structure. These effects can be summarized by focussing on BSA. The a-helix content for BSA plummeted from 54% in aqueous solution to 21% in the PLGA microspheres. Interestingly, this drop was significantly larger than that upon lyophilization (to 31%). One must conclude that the complete encapsulation procedure imposes significantly more stress upon the protein secondary structure than does lyophilization alone, the final step of the encapsulation procedure. Thus, the initial steps in the encapsulation protocol probably contributes significantly to the destabilization of protein structure. In particular, the formation of the first emulsion is suspicious in this context, but protein structural data have not been obtained thus far.

The detrimental structural changes could to some extent be prevented by the use of lyoprotectant trehalose in the encapsulation protocol. The a-helix content of BSA encapsulated in microspheres whilst in the presence of trehalose was 30% and thus significantly higher than in the absence of trehalose. It should be noted, however, that trehalose is only an established lyoprotectant that is efficient in preventing dehydration-induced structural changes [27,29,30]. Its potency in preventing protein structural perturbations caused by other stress factors involved in the encapsulation protocol is unclear. The data indicate that trehalose is obviously not efficient in eliminating all of the various stress factors efficiently in the w/o/w protocol.

Qualitatively, similar conclusions have been reached by Yang et al. [31] for recombinant human growth hormone. Therefore, thus far, a native protein structure has NOT been reported for proteins encapsulated in PLGA by the w/o/w technique.

Conclusions FTIR spectroscopy has been successfully employed to characterize the secondary structure of proteins in PLGA microspheres obtained by the w/o/w technique. However, significant procedure-induced structural perturbations have been diagnosed under all conditions. Stabilization of the protein secondary structure has been achieved to some extent by employing stabilizing additives, such as trehalose.

Structure-guided protein encapsulation
using alternative techniques

One can surmise from the above results is that there is an urgent need for the development of alternative procedures to encapsulate proteins in hydrophobic polymers. In the following, we will focus primariyl on those aspects of this work that includes protein structural information.

As one possible solution to the problems decided, we have developed recently an approach that we call structure-guided protein encapsulation [27,30]. Herein, FTIR spectroscopy is used to structurally guide the complete encapsulation procedure and any protein structural changes are systematically eradicated. One problem frequently encountered was the fact that the w/o/w technique does not allow us to encapsulate proteins in a unperturbed structure. Too many potential causes for structural changes have been identified and each of these require the development of a stress-specific stabilization method. We felt that the main factor responsible for the difficulties was the employment of proteins in aqueous solution. Under such conditions proteins are flexible - as demonstrated by their typically moderate temperature of denaturation. Most proteins denature reversibly or irreversibly below 100^oC [32,33]. In contrast, it is well established that dehydrated protein powders and also suspensions of those in a variety of organic solvents denature at quite high temperatures. For example, suspended protein powders can show catalytic activity for days in neat organic solvents at temperatures far above 100^oC [34,35]. This is due to the fact that dehydrated proteins are "rigid" in organic solvents. Thus, even though they can be denatured by the solvents for thermodynamic reasons, due to this increased rigidity they are trapped in their conformation for kinetic ones [36].

A novel concept emerged from the above scenario. If we employed lyophilized protein powders and suspend them in organic solvents, would it be possible to encapsulate proteins within PLGA having a native secondary structure? To test this concept, we employed BSA and recombinant human growth hormone as model proteins. Figure 3 shows the outline of the experiments.

It is important to note that the procedures we describe are not novel themselves. Encapsulation of protein powders in PLGA by simple solvent evaporation was actually one of the very first methods to be employed [17]. It leads to macroscopic delivery devices that are probably suitable for implanting purposes. However, due to their size,  such devices are problematic because the creation of defined release profiles for the drug is very difficult. Nonetheless, the above simple experimental scheme demonstrated proof of concept because, in contrast to investigations using the w/o/w technique to encapsulate proteins in PLGA, it allowed us to investigate the protein secondary structure by FTIR spectroscopy at all stages in the encapsulation process. Thus, every process variable could be investigated separately and the detrimental effects on protein secondary structure minimized or completely circumvented.

Scenario 1: Employing protein in the absence of any stabilizer.
The first set of experiments was designed to analyze the fate of a protein lyophilized without any stabilizer using the above procedure. The protein secondary structure was determined both for the lyophilized powder, the powder is suspensed in methylene chloride and subsequently dried, and finally for the powder dispersed in a PLGA film. The latter was obtained by suspending the lyophilized protein in methylene chloride containing the dissolved polymer PLGA followed by direct casting onto a CaF₂ FTIR window.

Figure 4 shows the fate of BSA when the protocol described above is carried out in the absence of any stabilizing additive. In aqueous solution, BSA has an a-helix content of ca. 57% (Bar 1). When BSA was lyophilized from such a solution, the a-helix content dropped significantly, in the case of our conditions to ca. 29% (Bar 2). If this protein was suspended by homogenization in methylene chloride and dried from this suspension, no further structural changes occurred, the a-helix content increased slightly (but statistically not significantly) to 35% (Bar 3). If this procedure was performed with PLGA dissolved in the solvent (and thus the final drying step caused encapsulation of BSA in the polymer matrix), the BSA structure was the same as it was without the polymer (39% a-helix, Bar 4). From this observation it can be concluded that the non-aqueous encapsulation procedure did not alter the structure of BSA significantly. The major structural changes occur during the lyophilization process. This is in contrast to the results obtained with the w/o/w procedure, where significant structural changes (in addition to those occurring upon lyophilization) are caused by the procedure. We also tested the impact of the method used to obtain the protein suspension, e.g. BSA was suspended by probe sonication in methylene chloride, whence the a-helix content was only 23% (Bar 5). Therefore, this method of creating the suspension of the protein in the organic solvent should be avoided.

From the above results one can surmise that, if lyophilization-induced structural changes are to be avoided, one should be able to obtain a protein with a more "native" secondary structure PLGA as a support.

Scenario 2: Employing protein in the presence of trehalose as a stabilizer.
Various lyoprotectants were tested for their ability to prevent lyophilization-induced structural changes in BSA. These results are available in our previous review (Griebenow et al., 1999). An additional difficulty encountered was that not all efficient lyoprotectants allowed the formation of a homogeneous suspension in the organic solvent. Trehalose proved to be the best additive in this context, even though structural preservation was superior employing sorbitol or glucose.

Figure 5 shows the results of the FTIR investigations. At the top, the spectrum we show the amide I band region obtained for BSA in aqueous solution. As is typical for proteins with a high a-helix content, the spectrum is dominated by a band located at around 1655-1660 cm ^-1. When assigning this band to the a-helix secondary structure, the result of the spectral analysis by Gaussian curve-fitting is in excellent agreement with the X-ray structural data. The next two spectra show BSA lyophilized with and without trehalose after suspension in methylene chloride and subsequent drying. It is important to note that the spectra are extremly similar to those of the preparations formed directly after lyophilization. Therefore, the spectra are representative examples for both situations. For the preparation obtained in the absence of trehalose (the second spectrum), significant spectral changes in the amide I demonstrate severe structural changes. In particular, the amide I band and its component show broadening. However, when the protein is co-lyophilized with trehalose, the same protocol produced a dramatically different result. The a-helix band is again clearly the dominant band of the spectrum and band-broadening is much reduced compared to the former situation. In other words, the secondary structure of the protein was largely preserved. Finally, the ultimate spectrum shows BSA encapsulated in a PLGA polymer film. The spectrum is again very similar to that of the protein in aqueous solution. Therefore, we can conclude that, when the encapsulation procedure is optimized, the protein structure can be significantly improved upon encapsulation in the PLGA polymer matrix.

In the following, the section qualitative observations made above are supported by quantitative data (a-helix content). Figure 6 shows the fate of the BSA when the protocol in Fig. 3 is carried out in the presence of the stabilizing additive trehalose (data from Carrasquillo et al., [27]). The a-helix content of BSA in aqueous solution is shown in Bar 1 and is 51%. Lyophilization of BSA in the presence of trehalose at a 1:4 weight ration (BSA:trehalose) largely prevents lyophilization-induced structural changes and the a-helix content determined was indeed 47% (Bar 2). Suspension of this preparation by homogenization in methylene chloride and subsequent drying did not induce any additional structural changes. The a-helix content was remarkably still 47% (Bar 3). In addition, encapsulation of this formulation in PLGA using homogenization (Bar 4) or a sonication bath (Bar 5) did not produce any significant structural changes.

Conclusions FTIR spectroscopy can be used to follow the fate of the model protein BSA in all the various encapsulation steps described. Combining the potential of novel encapsulation strategies with that of FTIR spectroscopy enable us to establish the structure-guided optimization of encapsulation conditions. Strategies have now been developed leading us to a significantly improved secondary structure of the protein in the PLGA polymer matrix. Similar investigations have also been performed with the model protein recombinant human growth hormone and lead us to identical conclusions [30].

Outlook

The technology we describe is currently being used to microencapsulate model proteins in PLGA. Herein, two immiscible organic solvents are used to create a single emulsion. The procedure is completly non-aqueous. BSA-loaded microspheres have already been obtained. Addional challenges have been encountered, e.g., the low yield of microspheres. In this context, the FTIR data collected previously have demonstrated the feasibility of our approach.

Acknowledgements

The authors acknowledge support by NIH-MBRS program (S06 GM08102-26S1), by the GAANN program of the US Department of Education, and the University of Puerto Rico (FIPI program).

References

1. Thayer, A.M. (1995) Technology-based firms define new business approach to drug development. C&EN, June 5, 17-26
2. Stinson, S. (1997) Growth factor protein may treat diabetes. C&EN, Nov. 24, 67-68
3. Tamura, Y., Peng, P., Liu, K., Daou, M., and Srivastava, P.K. (1997) Immunotherapy of tumors with autologous tumor-derived heat shock protein preparations.  Science, 278, 117-120
4. Dickman, S. (1998) Cancer therapy. Antibodies stage comeback in cancer treatment. Science, 280, 1196-1197
5. Robinson, J.R. (1997) Controlled drug delivery. Past, present, and future. In: Controlled Drug Delivery. Challenges and Strategies (K. Park, ed.), ACS Professional Reference Books, ACS, Washington DC, pp. 1-7
6. Lee, J.J. (1995) Biopharmaceutical properties and pharmacokinetics of peptide and protein drugs. In: Peptide-Based Drug Design. Controlling Transport and Metabolism (M.D. Taylor and G.L. Amidon, eds.), ACS Professional Reference Book, ACS, Washington, DC, pp. 69-100
7. Langer, R. (1990) New methods of drug delivery. Science, 249, 1527-1533
8. Langer, R. (1993) Polymer-controlled drug delivery systems. Acc. Chem. Res. 26, 537-542
9. Langer, R. (1995) Biomaterials and biomedical engineering. Chem. Eng. Sci. 50, 4109-4121
10. Langer, R. (1995) 1994 Whitaker Lecture: Polymers for Drug Delivery and Tissue Engineering. Annals of Biomededical Engineering 23, 101-111
11. Cleland, J.L. and Langer, R. (1995) Formulations and delivery of proteins and peptides: design and development strategies. In: Formulation and Delivery of Proteins and Peptides (J.L. Cleland and R. Langer, eds.), ACS Symposium Series, ACS Books, Washington DC
12. O'Hagan, D.T., Rahman, D., McGee, J.P., Davis, H., and Williams, P. (1991) Biodegradable microparticles as controlled release antigen delivery systems. Immunology 73, 239-242
13. Cohen, S., Alonso, M.J., and Langer, R. (1994) Novel approaches to controlled-release antigen delivery. Int. J. Technol. Assessment in Health Care 10, 121-130
14. Saltzman, W.M. (1993) Antibodies for treating and preventing diseases: the potential role of polymeric controlled release. Crit. Rev. Therapeutic Drug Carrier Systems 10, 111-142
15. Kissel, T., Li, Y.X., Görich, S., and Koneberg, R. (1996) Parenteral protein delivery systems using biodegradable polyesters of ABA block structure, containing hydrophobic poly(lactide-co-glycolide) A blocks and hydrophilic poly(ethylene oxide) B blocks. J. Controlled Release 39, 315-326
16. Pearlman, R. and Nguyen, T. (1992) Pharmaceutics of protein drugs. J. Pharm. Pharmacol. 44 (suppl. 1), 178-185
17. Langer, R. and Folkman, J. (1976) Polymers for sustained release of proteins and other macromolecules. Nature 263, 797-800
18. Schwendeman, S.P., Cardamone, M., Brandon, M. R., Klibanov, A. M., and Langer, R. (1996) Stability of proteins and their delivery from biodegradable polymer microspheres. In: Microparticulate Systems for the Delivery of Proteins and Vaccines (Cohen, S. and Bernstein, H., eds.), Marcel Dekker, New York, pp 1-49
19. Schwendeman, S.P., Costantino, H.R., Gupta, R.K., and Langer, R. (1997) Peptide, protein, and vaccine delivery from implantable polymeric systems. Progress and challenges. In: Controlled Drug Delivery. Challenges and Strategies (Park, K., ed.), American Chemical Society, Washington, D.C., pp. 229-267
20. Arakawa, T., Prestrelski, S.J., Kenney, W.C., and Carpenter, J.F. (1993) Factors affecting short-term and long-term stability of proteins. Advanced Drug Delivery Reviews 10, 1-28
21. Cleland, J.L., Lim, A., Barron, L., Desjardin, N., Duenas, E.T., Eastman, D.J., Vennari, J.C., Wrin, T., Berman, P., Murthy, K.K., and Powell, M.F. (1998) Development of a single shot subunit vaccine for HIV-1. 5. Programmable in vivo autoboost and long lasting neutralizing response. J. Pharm. Sci. 87, 1489-1495; Cleland, J.L. (1999) Single-administration vaccines: controlled-release technology to mimic repeated immunizations. Trends in Biotechnology 17, 25-29
22. Schwendeman, S.P., Costantino, H.R., Gupta, R.K., Klibanov, A.M., and Langer, R. (1997) Peptide, protein, and vaccine delivery from implantable polymeric systems. In: Controlled Drug Delivery. Challenges and Strategies (K. Park, ed.), ACS Professional Reference Books, ACS, Washington DC, pp. 229-267
23. Manning, M.C., Patel, K., and Borchardt, R.T. (1989) Stability of protein pharmaceuticals. Pharm. Res. 6, 903-918
24. Volkin, D.B. and Klibanov, A.M. (1989) Minimizing protein inactivation. In: Protein Function: A Practical Approach (Creighton, Ed.). Oxford University Press, pp. 1-24
25. Schöneich, C., Hageman, M.J., and Borchardt, R.T. (1997) Stability of peptides and proteins. In: Controlled Drug Delivery. Challenges and Strategies (K. Park, ed.), ACS Professional Reference Books, ACS, Washington DC, pp. 205-228
26. Krishnamurthy, R. and Lumpkin, J.A. (1998) Stability of proteins during manufacture and release from biodegradable polymers. BioPharm. 11(1), 32-36
27. Carrasquillo, K.G., Cordero, R.A., Ho, S., Franquiz, J.M., and Griebenow, K. (1998) Structure-guided encapsulation of bovine serum albumin in poly(DL-lactic-co-glycolic)acid. Pharm. Pharmacol. Commun. 4, 563-571
28. Fu, K., Griebenow, K., Hsieh, L., Klibanov, A. M., Langer, R. (1999) Use of FTIR spectroscopy to characterize the secondary structure of proteins encapsulated within PLGA microspheres. J. Contr. Rel. 58, 357-366
29. Costantino, H.R., Carrasquillo, K.G., Cordero, R.A., Mumenthaler,M., Hsu, C.C., and Griebenow, K. (1998) The effect of excipients on the structure and stability of lyophilized recombinant human growth hormone (rhGH). J. Pharm. Sci. 87, 1412-1420
30. Carrasquillo, K.G., Costantino, H.R., Cordero, R.A., Hsu, C.C., and Griebenow, K. (1999) On the structural preservation of recombinant human growth hormone in a dried film of a synthetic biodegradable polymer. J. Pharm. Sci. 88, 166-173
31. Yang, T.-H., Dong, A., Meyer, J., Johnson, O.L., Cleland, J., and Carpenter, J.F. (1999) Use of infrared spectroscopy to assess secondary structure of human growth hormone within biodegradable microspheres.
32. Lapanje, S. (1978) Physicochemical aspects of protein denaturation. Wiley, New York.
33. Privalov, P.L. (1992) Physical basis of the stability of the folded conformation of proteins. In: Protein Folding (Creighton, T.E., ed.), Freeman, New York, pp. 83-126
34. Zaks, A. and Klibanov, A. M. (1984) Enzymatic catalysis in organic media at 100°C. Science 224, 1249-1251
35. Volkin, D. B., Straubli, A., Langer, R., and Klibanov, A. M. (1991) Enzyme thermo-inactivation in anhydrous organic solvents. Biotechnol. Bioeng. 37, 843-853
36. Griebenow, K. and Klibanov, A.M. (1996) On protein denaturation in aqueous-organic but not in pure organic solvents. J. Am. Chem. Soc. 118, 11695-11700

REF:  K.Griebenow, I.Castellanos and K.G. Carrasquillo 
Int.J.Vibr.Spec., [www.ijvs.com] 3, 5, 2 (1999)

Feature Article

3. Raman and SERS spectroscopy
for probing drug-target interactions:
from in-vitro models to intracellular imaging

Ganesh D Sockalingum,
Serguei Charonov,
Abdelilah Beljebbar,
Hamid Morjani,
Michel Manfait & Igor Chourpa*

Unité MéDIAN, IFR 53, UFR de Pharmacie,
Université de Reims Champagne-Ardenne,
51 rue Cognacq-Jay,
51096 Reims cedex,
France

* Laboratoire de Chimie Analytique,
Faculté des Sciences Pharmaceutiques,
Université de Tours,
37200 Tours,
France

Tel: (33) 3 26 91 35 74 or (33) 3 26 91 35 53

e-mail to: ganesh.sockalingum@univ-reims.fr
or to:michel.manfait@univ-reims.fr

Abstract

Keywords

Anti-tumour drugs, Live cancer cells, Raman spectroscopy, SERS, Spectral imaging

Raman spectroscopy, in its simplest forms, is not a sensitive spectroscopic technique since Raman scattering cross-sections for most molecules are very small whereas the extinction coefficients in optical absorption or specific emissivities in emission spectroscopy. Fluorescence from the molecule of interest or other sample components further compromises detection, as the luminescence background often obscures a weak scattering signal. The investigator working at the level of the living cell is severely handicapped by these limitations since on the one hand cell components fluoresce strongly and on the other Raman sensitivity is seldom not enough to detect drugs at physiological concentrations. However, when these barriers to the use of Raman spectroscopy can be overcome, the method becomes a powerful tool for biological and biomedical research owing to its capacity to give molecularly specific and structurally useful information from such samples in a non-destructive and non-invasive manner. It is also one of the few available techniques that allow the selective study of molecular interactions within high-molecular-weight supramolecular complexes. The problem of sensitivity can be addressed either by using resonance Raman (RR), surface-enhanced Raman spectroscopy (SERS) or a combination of both (SERRS). Early studies have shown that RR is very useful because of its selectivity to certain chromophores and sensitivity to monitor drug-DNA complexes [1, 2]. But these advantages are exploitable mainly in vitro at relatively high concentrations and exist in the absence of fluorescence interference. For physiological concentrations of drugs (10^-6 M or less ), RR is not always sufficient, and also not all drugs possess a resonant moiety. SERS is a technique in which lasers are used to excite vibrational transitions in molecules adsorbed on a rough metallic substrate [3, 4]. As a result of large optical fields and resonance-related effects, the Raman cross-section of a molecule on a surface is enhanced by factors of up to 10⁶. When the SERS effect is coupled with an optical resonance in the molecule (SERRS), even larger enhancement can be observed. SERS is quite a good alternative because the process allows us to solve both sensitivity and fluorescence (quenching) problems. Another potential solution to the problem of fluorescence is NIR Fourier-transform (FT) Raman spectroscopy. This technique is less sensitive but offers the advantage of being rapid and causes less sample damage. Combined with the SERS technique, it can be an attractive method for probing the cell. To have access to molecular level information on drug-target interactions, both in-vitro and at the cellular level, our efforts have been mainly directed to the use of SERS, FT-Raman spectroscopy and combinations of both. The question is how can these techniques be adapted to solve specific problems such as drug-target interactions at the cellular level? This issue is important because understanding such molecular interactions help to have a better insight into the mechanisms of action and hence improve the development of more effective drugs.

Thus, there are several considerations to be taken into account prior to the application of SERS to such biological studies. It is well known that the SERS effect exhibits certain properties that distinguish it clearly from the normal Raman effect. First, the Raman cross-sections for adsorbed molecules are generally a factor of 10³-10⁶ larger than those of non-adsorbed ones. Second, the SERS spectrum may differ from that of the non-adsorbed free molecule in the sense that selective enhancement of certain bands or appearance of new bands may occur, often due to the contribution of the chemical component of the SERS effect. Finally, studies at cellular level implies the use of a micro-configuration. When SERS is coupled to a microscope one has in hand a powerful microprobe. This idea was first presented by Van Duyne et al.  [5], who calculated detection limits of less than 1 amol, corresponding to about 10⁵ (SERS) and ca. 600 (SERRS) molecules in the probe beam of their experiments.

Although electrodes are often the substrates of first source for SERS substrates investigations of most biological samples, but they are not ideal. This is mainly because biological systems are more complex, SERS-active electrode surfaces are quite tedious to prepare, and are not stable although they provide a good sensitivity [6]. Thus, other substrates such as colloids and vacuum-deposited metal island films have been developed and tested as a response to the problems arising from biological systems [7-10]. Both colloids and island films exhibit the necessary morphological features that are essential for the enhancement of the Raman cross-section of the adsorbed molecule. Apart from the type and morphology of the metallic surface, which must be rough and must have specific adsorption properties [11], the enhancement of the Raman signal is largely dependent on the excitation frequency [12] and, the chemical nature of the adsorbate. In general, silver colloids provide a reasonable enhancement from the visible to the NIR, silver island films are more suited for the visible, while gold island films and gold colloids work best from the red to the NIR [13].

Chemically prepared colloidal suspensions provide a certain number of advantages for SERS observation, including ease of formation and manipulation, simple characterisation by absorptiometric techniques and a definite dependence of the enhancement on particle size and shape. One major disadvantage is that the SERS activity of a given analyte depends to a certain extent on the method of sol preparation and on the exact protocol used for a given preparation. While the same problem also exists for metal island films, these provide perhaps the most reproducible substrate in terms of enhancement factors. However, since these are static substrates, there is always a risk that the buffer, solvent, or the intense laser fields associated with SERS, destroys the surface morphology that gives rise to surface enhancement.

Here, we give an example of how Raman spectroscopy can be used to selectively probe the interactions when dimethylcrocetin forms complexes with the human retinoic acid receptor RARg . In Figure 1 are displayed the RR (457.9 nm) and micro-FT-SERS (1064 nm) spectra from silver colloids of free DMCRT. Comparison of these spectra show that there are hardly any differences in the main band positions and relative intensities. This is an important observation since it tells us that (a) the adsorbed molecule, the analyte is not perturbed by its interaction with the colloidal particles and (b) spectral modifications observed upon complexation could then be reliably interpreted as a consequence of the drug-target interactions. To monitor the interactions in the DMCRT-RARg complex, we will pay particular attention to the bands positioned around 1540, 1165 and 1210 cm^-1 assigned respectively to u _(C=C) , (indicative of the degree of conjugation), and u _(C-C) motions (inform on the isomerisation state). When the micro-FT-SERS spectra of free DMCRT and those of DMCRT-RARg are compared (see Figure 2), the following differences can be noticed upon complexation: (a) there is a global loss of spectral intensity for the complex, (b) the I₁₅₄₁/I₁₁₆₅ ratio remains constant, (c) the I₁₅₄₁/I₁₂₁₀ decreases, and (d) the band at 1210 cm^-1 is upshifted. Following these observations that can be correlated to the interactions in the complex only, we can now see what happens at the cellular level. Two type of cell lines have been tested: K562, a human leukemic cell line established from a patient with chronic myelogeneous leukaemia in blast transformation and HL60, a human cell line established from the peripheral blood leukocytes of a patient with acute promyelocytic leukaemia. It has been shown that other retinoids such as all-trans retinoic acid induce differentiation of HL60 but not of K562 cells [14, 15]. This response is supposed to be due to the presence of the specific retinoid binding protein also called RAR, present in the HL60 cell nucleus but not in K562. This interaction leads to the formation of a retinoid-RAR-DNA ternary complex and subsequently specific gene expression [16]. The HL60 and K562 cell lines were incubated in the presence of 5 µM of DMCRT for 4 hours and washed twice with 0.9 % NaCl. After centrifugation, the pellets were then deposited on glass slides and left for 20 min at 37° C before being analysed by FT-Raman microspectroscopy. Due to the accumulation of the drug in the cells and the strong signal from the retinoid, it was possible using FT-Raman to record the spectra directly from the pellets. The cellular level data are compared to those of the free drug in Figure 3. It can be clearly seen that the spectrum obtained from the K562 cell corroborates with that of the free drug and that obtained from the HL60 cell can be correlated with that of the in-vitro experiment, with the DMCRT-RARg complex. When all spectral data are compiled in Table I and compared, the same four remarks observed for the in-vitro complex also apply to the HL60 cell experiment.

From these spectra and table, it can be seen that the I₁₅₄₁/I₁₁₆₅ ratio is almost constant throughout the whole set of experiments. For data obtained from the HL60 cells, the I₁₅₄₁/I₁₂₁₀ ratio is similar to that of the DMCRT-RAR in-vitro experiment whereas the I₁₁₆₅/ I₁₂₁₀ ratio decreases but has the same value in both sets. Values obtained for the K562 cells correlates with the free DMCRT in-vitro data both for I₁₅₄₁/I₁₂₁₀ and I₁₁₆₅/ I₁₂₁₀ ratios. These data together with the upshift of the 1210 cm^-1 band, observed in HL60 and DMCRT-RAR in-vitro complex, but not in K562 cells, tell us that there is an interaction, measurable by Raman spectroscopy, between the drug and its cellular target. These results are the outcome of three independent measurements. This study therefore demonstrates that FT-Raman and FT-SERS can be used as powerful probes for investigating the effect of differentiating agents (retinoids and carotenoids) in cancer cells related to the receptor expression.

Now, both K562 and HL60  are leukemic cell lines. Thus, Raman spectroscopy can allow us to distinguish between different cell types and can help to discriminate between sensitive and drug-resistant ones. This was not the aim of this work here. What we have shown is that using two cell lines, one containing a retinoid receptor (HL60) and the other does not (K562, control experiment), Raman spectroscopy can be used to monitor the interaction of the drug with its target. In one case there is a possibility of interaction and in the other there is not. This is observed from the spectral changes. So, the usefulness of the technique here is to (i) predict an interaction, and (ii) to select and improve new retinoids (e.g., synthetic ones) in view of a better interaction with their targets.

Another approach to such studies is to use other types of SERS-active substrates such as gold island films in conjunction with red or NIR excitations and dispersive instruments equipped with CCD detectors. Such systems are very sensitive and appropriate for studies at the cellular level. The absorption frequency maximum of gold island films shifts from the red to the NIR as their mass thickness increases. Although their preparation takes longer in comparison with other substrates, island films offer some control over the morphology and optical properties. This category of SERS substrates could be attractive for applications to biological systems since they can generate a fairly reasonable (but less than with silver colloids) long-range electromagnetic enhancement [13]. This property is important because the signal must originate from the molecule in the vicinity of the metal without being perturbed by the latter.

Figure 4 shows the SERS signal collected from a single HL60 cell treated with DMCRT, placed over a 50Å thick gold island film, and excited with a He/Ne laser. By looking at the spectrum, one can recognise the DMCRT bands already discussed  although they are slightly shifted. It can also be noticed that not only the signal from the  DMCRT is enhanced, but information from the cellular components is also available. For instance, the bands at 999 cm^-1 (phenylalanine), 1022 and 1063 cm^-1 (DNA backbone), 1262 and 1279 cm^-1 massif (amide III), 1485 and 1582 cm^-1 (these may arise from guanine and adenine). This spectrum contains more spectral information than those recorded before but is still dominated by the fluorescence background from the cell. So, the approach using island film SERS substrates (as an area probe external to the cell) for intracellular imaging can be envisaged. However, one has to be careful about the origin of the signal, whether it is intracellular or extracellular, since some drugs can quickly efflux out of the cell. Thus, different parameters have to be controlled before interpreting such spectra.

Instrumental considerations
Spectral imaging combines technologies like conventional imaging, spectroscopies and quantitative analysis to produce images for which the spectral signature is associated with each spatial element or pixel. It thus provides the possibility of using different spectral features and not only the integrated intensity within a given spectral range. A major recent development in spectroscopic instrumentation, that has contributed to spectral imaging, is based on the extension of the principles of confocal microscopy (see also article by Otto and Greeve in Int.J.Vib.Spect., [www.ijvs.com] 2, 3, 8 (1998). This mode gives a remarkably improved submicrometric spatial resolution, mostly in the vertical direction [17].

The scheme of a typical system for spectral imaging is illustrated in Figure 5. To scan a laser beam over a sample, either a motorised XY stage and/or mirror scanners can be used. The illumination schemes that can be applied to generate spatial images can be grouped into four main classes, namely (i) global, (ii) point, (iii) line, and (iv) plane.

The spectral image in this scheme is generated from band-pass filter images recorded at a successive range of wavelengths. The optical path is the same as for a conventional white light image. The investigated part of the sample is illuminated by the laser beam. The global illumination scheme is now almost never used. This is to indicate that the mode exists. A spectrograph filters the light with a band-pass d n which can be tuned and transmits a monochromatic image onto a 2D-detector. The spectral image is then generated from these filtered images and recorded successively for the whole desired wavelength range. The spectral resolution is weak and recording times for each image are very long. A grating spectrograph is used as a wavelength filter and transmits a monochromatic enlarged image onto a 2 dimensional detector. The position of the band-pass can be displaced by turning the grating [18]. The main disadvantages of this scheme are non-confocality, the low spectral resolution and the very large accumulation times for each image. This scheme is not often used for cell studies.

The point illumination scheme is the simplest procedure. Only one spatial point of the sample is illuminated by the laser beam and the spectrum corresponding to this point is recorded. Then, either the motorised sample stage moves the sample under the immobile laser focus or the turning mirror changes the position of the laser beam on the sample to the next investigated point. The main disadvantage of the point illumination scheme is the very low speed of spectral acquisition (tens or hundreds of minutes) which is a limitation to its application.

The illumination method is also possible incorporating a two-dimensional detector for simultaneous accumulation of many spectra [19]. One dimension of the detector is used for spectral information contained in the signal and the second one corresponds to the spatial distribution of the signal. The scanning mirror system should consist of two synchronised scanners. The first one deflects the probe laser beam onto the entrance of the microscope objective and produces a narrow line on the sample. The second one deflects the collected signal onto the detector. Scanning of the other space dimension canbe acheived by moving the sample in a direction perpendicular to laser line using a motorised stage.

Like the line illumination scheme, the plane illumination method makes use of a two-dimensional detector and two synchronised scanners [19]. The scanners must provide the possibility of displacing the probe laser beam in both X and Y space dimensions. It allows us to carry out simultaneous accumulation of spectra from different points on the parallel lines covering the investigated area of the specimen. Displacing the laser beam can be done by turning the scanner along two perpendicular axes. By changing the amplitudes and periods of motion corresponding to these two axes, different space resolutions can be achieved. In this way, the data on the detector are organised as strips of rows. Each strip corresponds to one line on the sample, and each row contains a spectrum originating from one point of this line. The capabilities of the CCD detectors used in modern scanning microspectrometers (~2000 pixels) allow us to accumulate the spectral image ~ 50x50 points. The very important advantage of the plane illumination scheme is the practically simultaneous scanning of the total image, and this significantly reduces the time-caused image distortions in unstable samples.

Intracellular Applications


We have reported recently resonance Raman and SERS studies of antitumor drugs on silver colloids [20-23]. In this section we give an example of how SERS signal of a drug can be collected in the vicinity of colloidal aggregates inside a living cell. This situation is different from that with films, discussed, since now the probe is inside the cell. The drug studied here is mitoxantrone, which is an anthraquinone anticancer drug structurally related to DNA-anthracycline antibiotics [24]. Mitoxantrone is a very potent drug and is used in the treatment of many types of cancer. It is able to cause condensation and compaction of chromatin, to disturb DNA-protein interactions and to trap the cleavable complex between DNA and nucleus-located DNA topoisomerase II. However, before the investigation of mitoxantrone inside a single live cell, we recorded RR and SERS spectra of the drug and its complexes with DNA [see also ref. 21]. The study of drug/target complexes in solution is a necessary prerequisite for the intracellular spectroscopic investigations. In vitro spectral characterisation of the free and complexed drugs is important for a better interpretation of the spectra recorded from cellular compartments.

For mitoxantrone, the experimental conditions used allowed the adsorption of the drug and even of the drug/DNA complexes without detectable perturbations of the drug or of the molecular interactions within the complex (see Figure 10 and comments therein). On the basis of these experiments and on band assignments by comparing RR and SERS, it has been possible to infer a preferential intercalation of rings A and B of the chromophore with the DNA double helix, while ring C remains outside. We have also confirmed this finding by NIR-FT-SERS experiments [25]. In fact, at very far off-resonance conditions of excitation at 1064 nm, CC-N and CC-O deformation modes at 441 and 466 cm^-1 and localised on rings A and C of the chromophore respectively, are strongly enhanced compared to the RR and normal SERS spectra. We observed that in the drug-DNA complex, the band at 441 cm^-1 disappeared, while that at 466 cm^-1 remained the same and was as strong as that near ~1300 cm^-1 band. The latter is the strongest feature in RR, pre-RR, SERS and FT-SERS and has been assigned to the ring stretching mode coupled with C-O stretching motions. As has been observed, this band, is very sensitive to the C-O group in ring A of the mitoxantrone molecule, decreases upon complexation.

Mitoxantrone (MXT)

The next step was to record SERS spectra from a single K562 cell by performing spectral imaging. The procedure is schematically depicted in Figure 11. K562 cancer cells (same as above) were treated with 1 µM drug concentration for 1 hr at 37 °C and washed twice with buffer by centrifugation. For the SERS measurements, the treated cells were then incubated with pre-aggregated silver colloid for 15 minutes. Non-penetrated aggregates were eliminated by successive washing in the buffer. Before the SERS measurements, the cells were tested for viability (95%). After treatment of a cell population with the drug and incubation with colloids (step A), one cell is selected under the microscope and spectra are recorded at regular intervals along a line (step B). This line of spectra is shown in step C, where one axis represents the frequency domain (cm^-1) and the other the points on the line. A different line is then recorded (either by a scanning laser or by moving the XY stage by 1-2 µm intervals). This line will give another pattern as in step C and all the lines are represented in step D, where one square corresponds to a line.

To summarise: the spectral image has been obtained with confocal SERS microspectroscopy from single, live K562 cancer cells treated with an anticancer drug, mitoxantrone, at a concentration of 10^-7M. SERS spectra have been recorded from the inside of the cell in which silver colloidal particles have been introduced to generate the SERS effect. First observation: a satisfactory S/N ratio is obtained although the laser power is incredibly weak (0.1 mW, i.e., 100 times less than in conventional Raman microspectroscopy). This range of power is quite acceptable for live cells, as it does not affect their viability (no hole in the cell!!). Second observation: with respect to classical SERS microspectroscopy, SERS spectral imaging offers better sampling of the spectral information coming from a given localisation. The superposition of the spectral image with a conventional image allows to attribute the spectral information to different cell compartments. The spectral image in this example was constructed following the distribution of the intensity of the mitoxantrone band at 1300 cm^-1 enhanced by the colloidal particles. The advantage of spectral imaging is that other representations can be used: band integral intensity, band ratios, functional groups etc.., and each time the information can be retrieved easily by clicking on any part of the image. The use of pseudo-colours can facilitate the image representation and in this case it is easier to see how the drug behaves in different cell compartments, e.g., bound to DNA in the nucleus, free in the cytoplasm…The potential of confocal SERS microspectroscopic imaging at the single cell level is very important for the selective analysis of drugs inside these cells.

The high sensitivity and molecular specificity of SERS microscopy and the confocal property of the instrument give us the possibility of generating multidimensional (space, frequency, time…) spectral images with high spatial resolution. Each dimension of the recorded data corresponds to a specific investigated feature, e.g., localisation of a drug (x, y, z), characterisation (u ), kinetics (s) or temperature (T). However, the SERS approach to intracellular investigation can be improved. So far, we have seen that the colloid particles can be introduced inside the cell by endocytosis or mixing. This introduction is random and can not be totally controlled by the investigator. Another possibility is to use point probes and to deposit, by perforation, the colloidal particles in any cell compartment using a micro-capillary. Cells are not irreversibly damaged by inserting the micro tip. Microelectrodes can also be used as point probes as they can easily perforate the cell and the SERS signal could then be recorded from any given point. SERS spectral imaging is done in the vicinity of the point probe and this enables a statistical analysis of the spectra. SERS spectral imaging could be done on larger areas by using area substrates. Thus using vacuum-deposited metal island films where, in principle, scanning the laser over the whole cell should result into enhanced signals from any point in the cell; in opposition to colloids, which are point probes, and their distribution being random (unless deposited by a microcapillary), enhanced signals are collected only from their vicinity. As we have explained before, island films can be potential candidates for such studies since they act as an external area probes. The combination of gold island films and NIR lasers (dispersive instruments and CCDs) should suit cellular level studies if the chromophore can benefit from long range enhancement. However, as we have mentioned care should be taken to ensure the origin of the SERS signal. Finally, the availability of PMTs which work in the NIR will breath new life into the use of Raman and increase the sensitivity of the technique for cell studies.

1. M. Manfait, L. Bernard and T. Theophanides, J. Raman Spectrosc., 11, 68 (1981).
2. G. Smulevich, A. Feis, A.R. Mantini, M. P. Marzocchi, Indian J. of Pure and Applied Physics, 26, 207 (1988).
3. M. Fleischmann, P. J. Hendra, and A. J. Mcquillan, Chem. Phys. Lett., 26, 123 (1974).
4. D. J. Jeanmaire and R. P. vanDuyne, J. Electroanal. Chem., 84, 1 (1977).
5. R. P. Van Duyne, K. L. Haller and R. I. Altkorn, Chem. Phys. Lett., 126, 190 (1986).
6. K. L. Norrod, L. M. Sudnik, D. Roussel, and K. L. Rowlen, Applied Spectrosc., 51, 994 (1997).
7. T. M. Cotton, J. H. Kim, and G. D. Chumanov, J. Raman Spectrosc., 22, 729 (1991).
8. M. Manfait, H. Morjani, R. Efremov, J. F. Angiboust, M. Polissiou, and I. Nabiev, in The Royal Society of Chemistry, ed. by R E. Hester and R. B. Girling, York, UK, 303-304 (1991).
9. Nabiev, H. Morjani, and M. Manfait, Eur. Biophys. J., 19, 311 (1991).
10. G. D. Sockalingum, I. Chourpa, J. M. Millot, I. Nabiev, S. Sharanov, and M. Manfait, in Biomedical applications of Spectroscopy, ed. By R. J. H. Clark and R. E. Hester, Wiley, Chichester, 49-87 (1996).
11. P. C. Lee and D. Meisel, J. Phys. Chem., 86, 3391 (1982).
12. J. A. Creighton, C. G. Blatchford, and M. G. Albrecht, J. Chem. Soc. Faraday II, 75, 790-798 (1979).
13. A. Beljebbar, G. D. Sockalingum, H. Morjani, J. F. Angiboust, M. Manfait, Spectrochim. Acta Part A, 53, 123, (1997).
14. T. R. Breitman, , S. E. Selonick, , and S. J. Collins, Proc. Natl. Acad. Sci., USA, 77, 2936 - 2940 (1980).
15. T. R. Breitman, , S. J. Collins, and B. R. Keene, Blood, 57, 1000 - 1004 (1981).
16. M. Petkovich, N. J. Brand, A. Krust, and P. Chambon,. Nature, 330, 444 (1987).
17. C. Favard, P.Valisa, M.Egret-Charlier, S.Sharanov, C. Herben, M. Manfait, E. da Silva, P. Vigny, Biospectroscopy, 5, 101, (1999).
18. P. Dhamelincourt, J. Barbillat and M. Delhaye, Spectroscopie Europe, 5/2 ,16, (1993).
19. S. Sharonov, I. Chourpa, P.Valisa, F.Fleury, A. Feofanov, M. .Manfait, Miscroscopy and Analysis, 4, 23, (1994 ).
20. I. Chourpa, H. Morjani, J.F. Riou, M. Manfait, FEBS Lett., 397, 61, (1996).
21. I. Nabiev, A. Baranov, I. Chourpa, A. Beljebbar, G.D. Sockalingum, M. Manfait, J. Phys. Chem., 99, 1608, (1995).
22. I. Chourpa and M. Manfait, J. Raman Spectrosc., 26, 813, (1995).
23. I. Nabiev, I. Chourpa, J.F. Riou, C.H. Nguyen, F. Lavelle, M. Manfait, Biochemistry, 33, 9013, (1994).
24. J. R. Brown and S. Heider Imam, Prog. Med. Chem., 21, 196, (1984).
25. A. Beljebbar, G. D. Sockalingum, J. F. Angiboust, M. Manfait, Appl. Spectrosc., 50, 148, (1996).

REF:  G.D. Sockalingum, S.Charonov, A. Beljebbar, H. Morjani, M. Manfait
& I. Chourpa Int.J.Vibr.Spec., [www.ijvs.com] 3, 5, 3 (1999)


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