- Email: [email protected]
Detection of algae and bacterial biofilms formed on titanium surfaces using micro-Raman analysis
Applied Surface Science 256 (2010) 5108–5115
Contents lists available at ScienceDirect
Applied Surface Science journal homepage: www.elsevier.com/locate/apsusc
Detection of algae and bacterial biofilms formed on titanium surfaces using micro-Raman analysis S. Ramya ∗ , R.P. George, R.V. Subba Rao, R.K. Dayal Corrosion Science and Technology Division, Indira Gandhi Centre for Atomic Research, Kalpakkam 603102, India
a r t i c l e
i n f o
Article history: Received 21 December 2009 Received in revised form 17 March 2010 Accepted 17 March 2010 Available online 25 March 2010 Keywords: Raman spectroscopy Colloids Biofouling Biofilms Characterization
a b s t r a c t Biofouling is one of the major impediments in the use of titanium in sea-water cooled condensers of power plants, which is otherwise an excellent material with respect to corrosion resistance. Raman microscopic experiments were carried out on biofilms grown on titanium surfaces to find out the chemical composition of complex extracellular polymeric substances (EPS) in the biofilm. Though the spectral resolution of normal Raman experiments on these systems was very poor, it was improved when microSERS experiments were carried out using mono and bimetallic Ag and Cu colloids. It was observed that spatial distribution of polysaccharides was higher than that of proteins in algae biofilms formed on titanium matrix. Similar experiments were performed on laboratory cultured bacterial films of Pseudomonas aeruginosa. It was evidenced that algal and bacterial biofilms on titanium can be clearly distinguished with the help of Raman mapping coupled with SERS technique using bimetallic Ag/Cu colloids. © 2010 Elsevier B.V. All rights reserved.
1. Introduction Microorganisms in the aquatic systems are susceptible to form biofilms on metallic surfaces. The interaction of these films with metal surface facilitates biofouling [1–7], which is thought to be one of the major problems in cooling water systems of many industries. For instance, titanium suffers serious biofouling, which is otherwise an excellent material for corrosion resistance. The surface coverage and thickness of these films form on titanium are seldom uniform. Since biofouling checking measures involve biofilm characterization, proper monitoring of biofilm formation and the quantification of its extracellular polymeric substances (EPS) are essential to impact the mechanical cleaning and water treatment programs. In this regard, the present work concerns about the detection and the semi quantification of algae and bacterial biofilms on titanium surfaces using Raman microscopy. The application of Raman spectroscopy to complex biological systems is of great importance [8–12] due to highly specific vibrational spectra. However, biological systems suffer overlapping of various spectral signatures and lesser intensity because of their large fluorescence background. Quenching of the fluorescence signals was desirable to enhance and resolve the vibrational spectra of EPS components, and was achieved normally by surface enhanced Raman spectroscopy (SERS) [13–16]. Generally, SERS is a strong tool for the detection of materials present in lower concentration. The
∗ Corresponding author. Tel.: +91 44 27480121; fax: +91 44 27480121. E-mail addresses: [email protected] (S. Ramya), [email protected] (R.P. George). 0169-4332/$ – see front matter © 2010 Elsevier B.V. All rights reserved. doi:10.1016/j.apsusc.2010.03.079
vibrational spectra of the analyte molecules in the proximity of substrate metal colloids can be obtained in SERS experiments where local optical fields of metallic colloids yield enhanced Raman signals of the surrounding environment [17–21]. Generally, natural biofilms can be algae, bacteria or fungi and their cell wall is made up of microfibrillar elements that provide rigidity and strength. The most common components of the microfibrils are polysaccharides, proteins, glycoproteins, lipids and minerals [22]. As far as the algae (plant) cell wall is concerned, the foremost portion constitutes polysaccharides. In the case of bacteria, the cell wall is peptidoglycan or lipoproteinaceous, made from polysaccharides or lipids cross-linked by unusual peptides containing d-amino acids. In other words, polysaccharides are present as major components in algae cell wall whereas proteins are the chief component in bacterial cell wall. The factor that creates the distinction between algae and bacterial biofilms is their composition of the cell wall wherein the polysaccharides to protein ratio varies distinctly. Our earlier study based on chlorophyll estimation indicated higher algal density in the marine water biofilm formed on the titanium matrices [23,24]. However, fast and effective method for monitoring the biofouling was needed. Hence, the single analysis through Raman spectroscopy with simple sample preparation was considered for analyzing the molecular composition in biofilm components. In this regard, Raman imaging was adapted to find out the distribution ratio of protein to polysaccharide counterparts so as to confirm the algal dominance on the titanium surfaces. There are many reports on SERS studies for the characterization of various single cells, proteins, amino acids and bacteria [25–38]. However, the application of bimetallic Ag/Cu colloids as SERS substrate for
S. Ramya et al. / Applied Surface Science 256 (2010) 5108–5115
biofilm detection using laser Raman microscopy was not reported. The reason for selecting bimetallic Ag/Cu colloids over individual Ag and Cu colloids as SERS substrates for biofilm detection was explored in the present task. For the first time, a promising approach to investigate biofouling on titanium surfaces with SERS using mono and bimetallic Ag and Cu colloids was attempted and it was intended to identify the difference between algal and bacterial biofilms based on Raman mapping experiments coupled with SERS technique. Generally, a possible way to measure the relationship between SERS spectra and the nature of the colloid is to perform the measurements for a given simple molecule in presence of different colloids. However, the performance of colloids depends on various parameters such as laser wavelength, particle size and analyte molecules. Proper substrate colloid and excitation wavelength must be chosen for any simple analyte molecule. An extensive study was done by Zeiri and Efrima [21] to study the effect of excitation wavelength and the colloid milieu and they reported that the observation of spectral signatures from different biochemical components of the microorganisms will be facilitated by altering the type of colloid and the laser excitation wavelength. They also stated that the normal Raman protocol can be switched to resonance or surface enhanced Raman by changing the relative spectral contributions of the various cell components and different types of information can be obtained depending on the types of colloid. In this respect, we made an attempt to find out a suitable substrate colloid for biofouling quantification and suggested a use of Ag/Cu bimetallic colloid as a better substrate than individual Ag and Cu systems for biofouling studies on titanium surfaces. Moreover, all our Raman experiments were carried out using red excitation since both the substrate and analyte were shown better stability throughout the analysis under this excitation. 2. Materials and methods 2.1. Algae and bacterial biofilm samples Two different kinds of biofilms on titanium surfaces i.e., algae and laboratory cultured bacterial biofilms were considered for Raman analysis. Commercially pure titanium Grade-2 samples were used in this study. The Ti metal was bought in the form of sheets and the coupons of size 30 mm × 20 mm × 1 mm were cut from it. These specimens were pickled in an acid bath (HNO3 , 400 g/L + HF, 40 g/L + water) to remove any surface scales present. The pickled specimens were then cleaned ultrasonically in a detergent solution, washed in running tap water and finally rinsed in distilled water and air-dried. One set of titanium specimens were suspended in the outfall water from Madras Atomic Power Corporation (MAPS), Kalpakkam. The specimens were withdrawn after 1 week and it was clearly seen that there was a considerable biofilm growth on the titanium surfaces. Based on epifluorescence microscopy and chlorophyll estimation, the algal dominance in the natural biofilm was confirmed (results are not shown here). This algal biofilms were kept for further analysis by Raman microscopy. Another set of titanium specimens were exposed to the culture of Pseudomonas aeruginosa bacteria in 10% nutrient broth for 72 h and withdrawn for biofilm characterization. 2.2. Colloidal milieu The silver and copper colloidal suspensions of 4 × 10−4 M concentration were prepared using the citrate and ascorbic acid reduction methods [39–41], respectively. For the preparation of bimetallic (1:4 molar ratio) Ag/Cu colloidal particles, the copper seed colloidal particles were initially prepared by reducing the
5109
aqueous cupric nitrate using ascorbic acid in presence of poly vinylpyrrolidone (PVP) as a capping agent. The reaction time for the preparation of copper colloid was about 7–8 h [40], followed by the addition of silver nitrate and the reaction was continued for another 2 h to get a reddish brown colloidal suspension. The added ascorbic acid served as a reducing agent for both copper and silver nitrates. Also, the synthesis for copper and Ag/Cu colloids was carried out in an inert atmosphere. The obtained mono and bimetallic colloidal particles were characterized in terms of their respective surface plasmon bands (figure not shown here) using UV–visible absorption spectroscopy. The individual silver and copper colloids have characteristic surface plasmon bands at 420 and 590 nm, respectively. In the present study, the synthesized bimetallic alloy colloid formation was confirmed by the presence bands at 450 and 583 nm due to surface plasmonic oscillations of silver and copper colloids. The shift in the position of the bands when compared to pure colloids was due to the formation of bimetallic alloy colloid of both the silver and copper systems. 2.3. Raman protocol for biofilms The titanium coupons with the biofilms were immersed in the colloidal suspensions and their analysis in Raman microscopic scanning mode were carried out with an HR 800 Jobin Yvon Raman spectrometer equipped with 1800 grooves/mm holographic grating. He–Ne laser of 633 nm was used as an excitation source. The system consists of an Olympus optical microscope mounted at the entrance of the Raman spectrograph and the 10× long distance objective was used to focus the beam on the sample. The laser spot size focused on the surface was approximately 3 m in diameter and laser power was 5 mW at the sample. The slit width of the monochromator was 300 m, which corresponds to a resolution of 4 cm−1 . The back scattered Raman spectra were recorded using super cooled (<−110 ◦ C) 1024 × 256 pixels CCD detector over the range 80–4000 cm−1 . Raman spectra were collected at various points (at least 4) of each sample, and were observed to be reproducible. All the spectra were collected with 4s integration and 20 accumulations. Both integration time and number of accumulations were kept constant for all the experiments. The obtained Raman spectra were further baseline corrected for better comparison. The micro-SERS spectra taken in different colloidal systems showed some minor differences in their peak positions due to slight changes in sampling, morphology and the heterogeneity of the biofilms on titanium surfaces. The spectral maps were obtained by collecting spectra over 200 m × 200 m area with 4 s exposure time and 20 CCD accumulations. Data acquisition was done by a software package (Kaiser-Optical System). 3. Results and discussion 3.1. Normal Raman experiments on algae biofilms on titanium surfaces The main reason for choosing the He–Ne laser with 633 nm as an excitation source was to avoid the degradation and strong fluorescence background of most of the cellular components. Normal Raman experiments (NR) on these biofilms on titanium surfaces did not give much information with respect to EPS of the biofilm. However, some medium and lower intensity peaks were detected (Fig. 1a) in the region below 800 cm−1 and these peaks were assigned to deformations, tetrahedral, ring breathing and stretching vibrations from diatomaceous silicate components in the cell wall [42]. The observed silicate vibrations were supported by the fact that the silicon is the main component of the cell wall of groups of algae [43]. For instance, the cell wall of the some brown and green
5110
S. Ramya et al. / Applied Surface Science 256 (2010) 5108–5115
well resolved Raman spectra. When SERS experiments were carried out using silver colloids, the Raman spectra gained from surface plasmon enhancement. However, at longer excitation wavelengths (here 633 nm) such enhancements were reduced and weaker spectra were obtained owing to the decrease in the effectiveness of silver colloids on some of the components in the cell wall. When considering the chemisorptions of silver with proteins, the bonding can be achieved through Ag–N. In this regard, peaks at lower wave numbers were expected. In the present case, we did not observe lower wavenumber peaks around 200 cm−1 due to the use of longer excitation wavelength and citrate silver colloids [21]. Also, these features were strongly dependent on metal-adsorbate interactions [21]. The micro-SERS experiments using silver colloid gave enhancement of polysaccharide frequencies of algae biofilm. However, Raman signatures from the proteins were not enhanced. Hence, Raman mapping experiments using silver colloids were not carried out to find out the protein and polysaccharide distributions.
Fig. 1. Micro-Raman spectra of the algae biofilms on titanium surfaces; (a) represents the Raman spectra taken in the absence of colloid, (b) represents the Raman spectra taken in presence of Ag colloid, (c) represents the Raman spectra taken in presence of Cu colloid. Laser excitation was 633 nm using He–Ne laser, laser power = 5 mW, exposure time = 4 s, CCD accumulations = 20.
algae is enriched in silicon. Algae diatoms generally take silicon up as silicates [43]. 3.2. Micro-SERS experiments on algae biofilms on titanium surfaces in presence of Ag colloid Normal Raman experiments (NR) on these biofilms on titanium surfaces gave vibrations corresponding to silicates and it was difficult to resolve other spectral signatures with regard to organic components, even by increasing the exposure time and the number of CCD accumulations. Hence, the experiments were carried out in presence of silver colloids to trace out some of the major components. The intensity of micro-SERS spectrum was higher by many orders in magnitude than that of NR experiments. A tentative band assignment was performed by referring to Raman and SERS assignments for biological samples from the literature [25–38,44–59]. A strong peak at 1398 cm−1 must be due to COO− symmetric stretching vibrations from glycoproteins or polysaccharides [48]. The SERS spectra (Fig. 1b) with Ag colloid showed the presence of polysaccharides owing to the presence of some smaller intensity bands 300–600 cm−1 [16,45,56]. Also, the peaks around 970–998 cm−1 were due to the presence of O–CH3 stretching from polysaccharide units [47,57]. In addition to that, a small peak at 1068 cm−1 was seen and attributed to C–O and C–C stretching frequencies of polysaccharides [44,45]. A small intensity peak at 889 cm−1 was assigned to C–C and C–O–C glycosidic linkages [46,49,50]. A strong band was identified around 2900 cm−1 due to C–H stretching vibrations from both polysaccharides and proteins [12,47]. In addition to that, medium intensity peaks were observed at 1504 [56,58] and 1595 cm−1 [44,50,56] due to C C stretching vibrations and aromatic ring stretching frequencies, respectively. The peak at 1660 cm−1 [57] can be assigned to ring conjugated C C stretching vibrations. Though red excitation (633 nm) was used, the micro-SERS spectrum from silver colloid was superimposed on a background, due to the presence of high fluorescence nature of various components in the algae biofilm. Also, features from the protein components were not clearly observed. In addition to that, the silicate peaks observed in NR experiments were not detected in micro-SERS due to the overlapping of silicates and polysaccharide frequencies. It was expected that quenching of fluorescence to tolerable levels by silver colloids will allow the acquisition of
3.3. Micro-SERS experiments on algae biofilms on titanium surfaces in presence of Cu colloid One of the principal features that we wanted to examine was the effect of types of colloids on the SERS spectra. Replacing silver by copper changed the optical properties (copper plasmons are red shifted to 170 nm compared to silver particles) of analyte and substrate. Very strong and informative spectra were obtained with 633 nm excitation using copper colloids. Possible reason that, the excitation wavelength was closely in resonance with copper colloids than silver colloids. Fig. 1c depicts the micro-SERS spectra of algae biofilm in presence of copper colloid, which shows similar features as that of silver colloids in terms of band positions. However, major differences were observed below 800 cm−1 and the Raman signals were about 10 times higher than those with silver colloids and show many additional features with regard to proteins. It was clearly evidenced that copper colloid was found to be a better candidate in resolving the signal of both polysaccharides and proteins of algae biofilm when compared to silver colloid. However, the major difficulty in dealing with the copper colloid was its stability. Especially, when Raman mapping experiments were carried out, copper colloids readily got oxidized due to longer laser irradiation time, and no Raman signals were observed after sometime. Hence, the idea of getting spatial distribution of particular component in the biofilm was merely lost. Though copper colloid gave larger enhancement and better resolution, it suffered greater instability compared to silver colloid. It was concluded that copper colloidal dispersion is a good substrate for the detection of cellular components but not for their quantification. 3.4. Micro-SERS experiments on algae biofilms on titanium surfaces in presence of Ag/Cu bimetallic colloid In the present investigation, we wanted to distinguish between algal and bacterial biofilms based on their polysaccharide and protein distributions. Though micro-SERS with copper colloids gave better results, in order to get quantification in terms of spatial distribution using Raman mapping, the colloidal milieu was modified from Cu to Ag/Cu bimetallic colloids of better stability and the resolving power of both the individual silver and copper colloids was also retained. Fig. 2a shows the micro-SERS spectra of algae biofilm using bimetallic Ag/Cu colloids and the corresponding frequencies were mentioned in Table 1. Even though the intensity of the SERS spectra with this bimetallic milieu was slightly lesser than that of pure copper colloid, the spectral resolution with regard to proteins and polysaccharides was significantly improved. Fig. 3a shows the average micro-SERS spectra taken from the algae biofilm, wherein the presence of dominant polysaccha-
S. Ramya et al. / Applied Surface Science 256 (2010) 5108–5115
5111
Table 1 Assignments of the peaks (cm−1 ) for the micro-SERS spectra of natural algae biofilm in various types of colloids using 633 nm excitation. Peak position (cm−1 ) Ag colloid
Peak position (cm−1 ) Cu colloid
417
356 489 555
619
636 716 755 801
Peak position (cm−1 ) Ag/Cu colloid
560 670
804
889 970 998
910 973 1012
1067
1082
1160
1174
1271
1256 1332 1375
1137
1356
1504
1448; 1452
1595 1660
1580
1570
2901 Other small
2930 peaks
2976
Tentative peak assignments Carbohydrates [16,45,56] Carbohydrates [16,45,56] C–O–C glycosidic ring deformation of polysaccharides and COO− wagging and C–C skeletal [16,44] COO− bending [16,51] CH2 ; Adenine ring stretching [44] Ring breath of proteins [44,56,59] Ring breath of proteins and O–P–O stretching from nucleic acids [44]; Different C–N stretching from proteins [44,58,59] C–C stretching and C–O–C glycosidic linkage [44,59] Different C–N stretching from proteins [44] O–CH3 stretching [47] Carbohydrates [57] C–CH3 deformation from pigments [58,56] C–O; C–C str; C–OH deformation from carbohydrates [44,46] C–C stretching; C–O ring breathing and asymmetric stretching [45,56] CH2 twisting and C–H in-plane deformation [56]; Amide III [44,56] ı(CH) of proteins and carbohydrates [56] COO− symmetric stretching [48,56] COO− symmetric stretching; ␦ (CH2 ) scissoring and CH2 deformation [48] ı(CH2 ) scissoring and CH2 deformation of proteins, lipids and carbohydrates [48,58] C C stretching from pigments [56,58] G-A ring stretching [44,50,56] Ring conjugated C C stretching [57] Ring conjugated C C stretching; C O stretching from lipids or carbohydrates [47,56] Amide I [44,49,56] CH and CH2 stretching [12,47] Unassigned
tribution of polysaccharides than proteins in algae biofilm. The Raman map of polysaccharides was taken from 990–1100 cm−1 (C–O and C–C stretching of polysaccharides). Similarly the map of proteins was taken from 770 to 850 cm−1 (ring breathing and different C–N stretching vibrations of proteins). The mapping experiments were repeated for other spectral regions of polysaccharides and proteins. The observed results were reproducible and in good agreement with the earlier findings. 3.5. Micro-SERS experiments on Pseudomonas aeruginosa bacterial films on titanium surfaces in presence of bimetallic Ag/Cu colloid
Fig. 2. (a) represents the micro-SERS spectra extracted from the Raman map of algae biofilm on titanium surface in presence of bimetallic Ag/Cu colloid. (b) represents the micro-SERS spectra extracted from the Raman map of bacterial biofilm on titanium surface in presence of bimetallic Ag/Cu colloid. Laser excitation was 633 nm using He–Ne laser, laser power = 5 mW, exposure time = 4 s, CCD accumulations = 20.
ride, glycoproteins and lipid frequencies around 1000–1500 cm−1 was identified. Fig. 4a depicts the light microscopic image of the algae biofilm formed on the titanium surface. Raman mapping experiments were performed on the selected region of the algae biofilm. The Raman spectral maps for proteins and polysaccharides (Fig. 4b–d) were obtained by collecting the spectra with 4 s exposure time. The mapping experiments showed the greater dis-
Due to the good performance of bimetallic Ag/Cu colloid on algae biofilm analysis, the colloidal milieu was expected to show similar trend for bacterial biofilm. In this regard, NR and micro-SERS experiments were carried out on laboratory cultured P. aeruginosa bacterial film. The resolution of NR spectrum of P. aeruginosa was very poor and the identification of components in the biofilm was impossible. However, micro-SERS of the P. aeruginosa film in presence of bimetallic Ag/Cu colloid was distinct and well resolved (Fig. 2b). The obtained frequencies for the P. aeruginosa film (Table 2) were in good agreement with the literature findings [51]. Table 2 depicts the tentative assignments of the peaks seen in the micro-SERS spectra of P. aeruginosa. The Raman peaks around 500–1300 cm−1 region were due to various groups including glycosides, nucleic acids, proteins, polysaccharides and unsaturated fatty acids that are present in bacterial cell [51]. The peak at 1655 cm−1 was due to amide I [16]. We did not get specific phenylalanine peak at 1004 cm−1 as reported for many bacterial cultures due to the overlapping of other spectral frequencies. Another pos-
5112
S. Ramya et al. / Applied Surface Science 256 (2010) 5108–5115
Fig. 3. (a) represents the average Raman spectra taken from the algal biofilm on Ti surface. (b) represents the average Raman spectra taken from the bacterial biofilm on Ti surface. Laser excitation was 633 nm using He–Ne laser, laser power = 5 mW, exposure time = 4 s, CCD accumulations = 20. The average spectra were collected over 200 m × 200 m area.
Fig. 4. Distribution of polysaccharides and proteins in algae biofilm formed on titanium matrix. (a) represents the light microscopic image of the algae biofilm obtained by Olympus Raman microscope using 10× long distance objective. The square box inside (a) represents the selected region of interest where Raman mapping was carried out. In (a) dark region corresponds to algae biofilm whereas the bright region corresponds to Ti surface. (b) represents the Raman map of polysaccharides and the mapping region covers 990–1100 cm−1 with a central peak position at 1082 cm−1 . (c) Represents the Raman map of polysaccharides and the mapping region covers 600–700 cm−1 with a central peak position at 670 cm−1 . (d) represents the Raman map of proteins and the mapping region covers 770–850 cm−1 with a central peak position at 809 cm−1 . Table 2 Tentative peak assignments [48] (cm−1 ) for micro-SERS spectra of P. aeruginosa in bimetallic Ag/Cu colloid using 633 nm excitation. Band (cm−1 )
Assignments
360 487 639 767
Carbohydrate [16,45,51,56]. Carbohydrate [16,45,51,56]. C–O–C glycosidic ring deformation of polysaccharides; COO− wagging and C–C skeletal [20,53]. Ring breath of proteins [44,56,59]. Different C–N stretching from proteins [51]. Polysaccharides and C–C, C–O, C–O–H deformation [51]. N–H, C–N, amide III (protein) [51,56]. CH2 bending from proteins and lipids [52,56]. G–A ring stretching [44,50,56]. Amide I [16]. C–H stretching [47,12]. Unassigned.
876 1052 1280 1473 1571 1655 2900, 2953 Other small peaks
sibility is that chemical effects may control the production of SERS-active centers in specific sites within the cellular components [21]. Amides III bands at 1220–1320 cm−1 are absent in the SERS spectra of algae film due to the broad overlapping of other spectral signatures in that region (Fig. 2a). However, it appeared as a weak band in the bacterial SERS spectrum (Fig. 2b). The Raman mapping experiments were carried out on P. aeruginosa bacterial films in bimetallic Ag/Cu colloid to see the spatial distribution of polysaccharides and proteins. Experimental parameters for Raman mapping experiments were kept as same as it was done for algae biofilms. The choice of the frequencies for mapping should be the same for both algae and bacterial biofilms. However, in the present Raman investigation of bacterial biofilms, we did not trace out proper Raman signals for proteins in the region 770–850 cm−1 (chosen for algae mapping). But, distinct signal was obtained at 876 cm−1 and was accounted for proteins based on the literature. Since our aim was to see the differences in the protein ratios in algal and bacterial biofilms, the variations in the spectral positions (due to different Raman active modes) of proteins were considered less
S. Ramya et al. / Applied Surface Science 256 (2010) 5108–5115
5113
Fig. 5. Distribution of polysaccharides and proteins in bacterial biofilm formed on titanium matrix. (a) represents the light microscopic image of the bacterial biofilm obtained by Olympus Raman microscope using 10× long distance objective. The square box inside (a) represents the selected region of interest where Raman mapping was carried out. (b) represents the Raman map of proteins and the mapping was carried out with respect to 1280 cm−1 corresponding to amide III band. (c) represents Raman map of proteins and the mapping was carried out with respect to 876 cm−1 corresponding to different C–N stretching of proteins. (d) represents Raman map of polysaccharides and the mapping was carried out from 990 to 1100 cm−1 corresponding to C–O and C–C stretching of polysaccharides. (e) represents Raman map of polysaccharides and the mapping was carried out from 600 to 670 cm−1 corresponding to C–O–C vibrations of glycosidic ring deformations.
important. In order to overcome this disadvantage, Raman mapping of proteins was repeated at 1280 cm−1 (amide III) for verifying the mapping results. For bacterial biofilms, the features corresponding to proteins and polysaccharides were clearly seen. Hence, the mapping was carried out with respect to those frequencies without any uncertainty. However, the mapping experiments on algal biofilms led to a small confusion in the selection of proper protein and polysaccharide vibrations. Because, most of the protein vibrations including amide bands were absent due to the broad overlapping of other spectral signatures. Hence, the region 770–850 cm−1 was chosen for mapping the protein signatures of algal biofilm. The strong and prominent peaks at 1356 and 1570 cm−1 were not considered for algal mapping. Because, the region 1350–1400 cm−1 was attributed to the presence of COO− symmetric stretching; ı(CH2 ) scissoring and CH2 deformation from proteins, lipids and carbohydrates [48,56,58] and the peaks around 1500 cm−1 were due to aromatic ring stretching frequencies [56,58]. It was observed from the bacterial mapping experiments (Fig. 5) that the spatial distribution of polysaccharides was less compared to that of proteins, which is a characteristic feature of bacterial cell wall. Similar mapping was carried out for other polysaccharides and protein spectral regions and the observed distribution of polysaccharides and proteins ratios were consistent and good agreement with the previous results. The bacterial spectrum was in less coin-
cidence with the algae spectrum. In the Raman spectra of the algae biofilm, signals due to proteins and amino acids were not prominent. Rather, strong signals with comparatively higher background were obtained around 1350–1400 and 1580–1600 cm−1 . The spectral region 1350–1400 cm−1 was attributed to the presence of COO− symmetric stretching; ı(CH2 ) scissoring and CH2 deformation from lipids and carbohydrates [48,56,58]. Some medium intensity peaks were observed around 1500 cm−1 due to C C stretching vibrations and aromatic ring stretching frequencies [56,58]. The Raman mapping experiments on the algae biofilms showed more polysaccharide distribution whereas the bacterial biofilm showed greater protein distributions. The differences between these two biofilms were clearly observed from the Raman mapping experiments. Though the algal and bacterial biofilms showed the presence of basic proteins and polysaccharide signatures as expected, an interesting feature was observed with regard to C–H stretching frequencies around 2900 cm−1. In the Raman spectra of algal biofilm, single peak was observed around 2900 cm−1 . In the case of bacteria, a strong doublet was detected in the same wave number region due to C–H and CH3 stretching vibrations from proteins and lipids. Since Gram-negative P. aeruginosa bacteriam has a characteristic lipoproteinaceous cell wall, these vibrations were probably aroused from the proteins and the lipids of the cell wall. As per literature, algae cell wall consists more of polysaccharides whereas bacterial mem-
5114
S. Ramya et al. / Applied Surface Science 256 (2010) 5108–5115
brane contains more protein (peptidoglycan) units. This is the basis for the present study to detect the micro levels of algae and bacteria on the titanium condensers using SERS coupled with Raman microscopy. Generally, Raman map gives the intensity distributions of one particular spectral range from which the changes in the composition of a specific component can be determined. In our present study, we selected lesser number of data points (36 points) and hence, the image obtained was coarse and not exactly matching with the light microscopic image. Also, the Raman map represented the image distribution of one particular wave number range (correspond to one component) whereas the light microscopic image depicted the distribution of all the components. It was assumed that different protein to polysaccharide ratio will be reflected in the Raman mapping. Since colloids enhance the Raman signals of molecular components in their close proximity even when it is not the major constituent, the titanium coupons with the biofilms were immersed in the colloidal suspensions throughout the Raman analysis. When the Raman experiments were carried out for pure P. aeruginosa bacterial culture, it was observed that both polysaccharide and protein frequencies were enhanced and the idea of doing mapping experiments to find out the protein to polysaccharide distribution was achieved. Hence, the Raman microscopy coupled with SERS using bimetallic Ag/Cu colloid was identified as an effective tool for differentiating algae and bacterial biofilms in order to see the impact of biofouling on Ti surfaces.
lurgy and Materials Group, Dr. A.K. Arora and Dr. M. Rajalakshmi, Materials Science Division, and Dr. Baldev Raj, Director, Indira Gandhi Centre for Atomic Research, Kalpakkam, during the course of this work. The authors thank Dr. M.P. Srinivasan and Mr. P. Chandra Mohan, BARCF, Kalpakkam for their timely help and suggestions. The support of K.S. Krishnan research fellowship is also acknowledged. References [1] [2] [3] [4] [5] [6] [7] [8] [9] [10] [11] [12] [13]
[14]
4. Conclusions The Raman microscopic experiments were carried out on algae and bacterial biofilms grown on titanium surfaces in order to have a proper understanding of the chemical composition of the complex extracellular polymer substances (EPS) in the biofilm. The salient conclusions are as follows: (a) normal Raman experiments on algae biofilms gave information only about silicates and the spectral details corresponding to organic components in the biofilm were not identified due to high complexity of the biofilm. (b) Micro-SERS experiments using silver colloids on these biofilms clearly indicated the contributions of polysaccharides. However, the significance of protein was not identified distinctly. (c) Resolution with regard to both polysaccharides and proteins was remarkably increased when similar micro-SERS experiments were performed using copper colloids. Conversely, it was observed that copper colloid was good for identification of the components but not for their quantification due to their lack of stability while performing mapping experiments. (d) Similar experiments using bimetallic Ag/Cu colloids showed good results and hence, it was easy to resolve the signals of most of the algae biofilm components. The Ag/Cu colloid showed better stability than individual silver and copper colloids and the Raman mapping experiments were conducted to explore the spatial distribution of biofilm components. (e) Raman mapping experiments using bimetallic Ag/Cu colloids on algae biofilm showed greater the distribution of polysaccharides than proteins. (f) The effectiveness of bimetallic Ag/Cu colloid was checked even for the laboratory cultured P. aeruginosa bacterial films. It was concluded that Raman microscopy coupled with SERS technique was an effective tool for the quantification of algae and bacterial biofilms on titanium surfaces in order to improve the control measures of biofouling of condenser materials in power plants.
[15] [16] [17] [18] [19] [20] [21] [22] [23] [24] [25] [26] [27] [28] [29] [30] [31] [32] [33] [34] [35] [36] [37] [38] [39] [40] [41] [42] [43]
Acknowledgements [44]
The authors wish to acknowledge the support and encouragement extended by Dr. P.R. Vasudeva Rao, former Director, Metallurgy and Materials Group, Dr. T. Jayakumar, Director, Metal-
[45] [46]
W.G. Characklis, K.E. Cooksey, Adv. Appl. Microbiol. 29 (1983) 93–138. B. Little, P. Wagner, F. Mansfeld, Mater. Rev. 36 (1991) 253–258. R.P. George, D. Marshall, R.C. Newman, Corros. Sci. 45 (2003) 1999–2015. X. Shi, R. Avci, M. Geiser, Z. Lewandowski, Corros. Sci. 45 (2003) 2577–2595. K. Pedersen, Water Res. 24 (1990) 239–243. O.M. Vaisanen, E.L. Nurmiaho-Lassila, S.A. Marmo, M.S. Salkinoja-Salonen, Appl. Environ. Microbiol. 60 (1994) 641–653. R.P. George, P. Muraleedharan, N. Parvathavarthini, H.S. Khatak, T. Rao, Mater. Corr. 51 (2000) 213–218. N.J. Kline, P.J. Treado, J. Raman. Spectrosc. 28 (1997) 119–124. D. Naumann, H. Helm, Labischinski, Nature 351 (1991) 81–82. D. Naumann, S. Keller, D. Helm, C. Schultz, B. Schader, J. Mol. Struct. 347 (1995) 399–405. G.J. Puppels, F.F.M. Demul, C. Otto, J. Greve, M. Robertnicoud, D.J. Arndtjovin, T.M. Jovin, Nature 347 (1990) 301–303. M.F. Escoriza, J.M. VanBriesen, S. Stewart, J. Maier, P.J. Treado, J. Microbiol. Methods 66 (2006) 63–72. L.-P. Choo-Smith, K. Maquelin, T.V. Vreesuijk, H.A. Bruining, G.J. Puppels, N.A. Ngo Thi, C. Kirsciner, D. Naumann, D. Ami, A.M. Villa, F. Orsini, S.M. Doglia, H. Lamfarrej, G.D. Sockalingam, M. Manfait, P. Allouch, H.P. Endltz, Appl. Environ. Microbiol. 67 (2001) 1461–1469. K. Maquelin, L.-P. Choo-Smith, T.V. Vreeswijk, H. Endtz, B. Smith, R. Bennet, H. Bruining, G. Pupples, Anal. Chem. 72 (2000) 12–19. A.C. Williams, H.G.M. Edwards, J. Raman. Spectrosc. 25 (1994) 673–677. K.C. Schuster, I. Reese, E. Orlaub, J.R. Gapes, B. Lendi, Anal. Chem. 72 (2000) 5529–5534. K. Kneipp, Y. Wang, H. Kneipp, L. Perlman, I. Itzkan, Phys. Rev. Lett. 78 (1997) 1667–1670. S. Nie, S.R. Emory, Science 275 (1997) 1102–1106. C.D. Keating, K.K. Kovaleski, M.J. Natan, J. Phys. Chem. B 102 (1998) 9414–9425. S. Lefrant, J.P. Buisson, J. Schreiber, O. Chauvet, M. Baibarac, I. Baltog, Synth. Met. 139 (2003) 783–785. L. Zeiri, S. Efrima, J. Raman. Spectrosc. 36 (2005) 667–675. B. Alberts, D. Bray, J. Lewis, M. Raff, K. Roberts, J.D. Watson, Molecular Biology of the Cell, 2nd ed., Garland Publishing, New York, 1989. J. Gopal, B.V.R. Tata, R.P. George, P. Muralidharan, R.K. Dayal, Surf. Eng. 24 (2008) 447–451. J. Gopal, R.P. George, P. Muraleedharan, H.S. Khatak, Biofouling 20 (2004) 167–173. G.D. Sockalingum, H. Lamfarraj, P. Pina, A. Beljebbar, P. Allouch, M. Manfait, Spectrosc. Biol. Mol. New Dir. 8 (1999) 599–600. R. Keir, D. Sadler, W.E. Smith, Appl. Spectrosc. 56 (2002) 551–559. S. Farqharson, A.D. Gift, P. Maksymiuk, F.E. Inscore, Appl. Spectrosc. 58 (2004) 351–354. L.A. Gearheart, H.J. Ploehn, C.J. Murphy, J. Phys. Chem. B 105 (2001) 12609–12615. E. Podstawka, Y. Ozaki, L.M. Proniewicz, Appl. Spectrosc. 58 (2004) 570–580. E. Podstawka, Y. Ozaki, L.M. Proniewicz, Appl. Spectrosc. 58 (2004) 581–590. E. Podstawka, Y. Ozaki, L.M. Proniewicz, Appl. Spectrosc. 58 (2004) 1147–1156. L. Zeiri, B.V. Bronk, Y. Shabtai, J. Czege, S. Efrima, Colloids Surf. 208 (2002) 357–362. R.M. Jarvis, A. Brooker, R. Goodacre, Anal. Chem. 76 (2004) 5198–5202. A. Shamsaie, J. Heim, A.A. Yanik, J. Irudhayaraj, Chem. Phys. Lett. 461 (2008) 131–135. R. Patzold, M. Keuntje, A.A. Von Anlften, Anal. Bioanl. Chem. 386 (2006) 286–292. P. Rosch, M. Schmitt, W. Kiefer, J. Popp, J. Mol. Struct. 661 (2003) 363–369. K.C. Schuster, E. Urlaub, J.R. Gapes, J. Microbiol. Methods 42 (2000) 29–38. D. Zhang, Y. Xie, M. Mrozek, C. Ortiz, J. Davisson, D. Ben-Amotz, Anal. Chem. 75 (2003) 5703–5709. P.C. Lee, D. Meisel, J. Phys. Chem. 86 (1982) 3391–3395. W. Yu, H. Xie, L. Chen, Y. Li, C. Zhang, Nanoscale Res. Lett. 4 (2009) 465–470. Y. Wang, P. Chen, M. Liu, Nanotechnology 17 (2006) 6000–6006. P. Yuan, H.P. He, D.Q. Wu, D.Q. Wang, L.J. Chen, Spectrochim. Acta Part A 60 (2004) 2941–2945. Ji. Zhen-Gang, Water quality and eutrophication, in: Hydrodynamics and Water Quality: Modeling Rivers, Lakes and Estuaries, John Wiley & Sons, Inc., Hoboken, NJ, 2008, p.289. K. Maquelin, C. Kirschner, L.-P. Choo-Smith, N. van den Braak, H.P. Endtz, D. Naumann, G.J. Puppels, J. Microbiol. Methods 51 (2002) 255–271. K. De Gussem, P. Vandenabeele, A. Verbeken, L. Moens, Spectrochim. Acta Part A, 61 A (2005) 2896–2908. J.T. Pelton, L.R. McLean, Anal. Biochem. 277 (2000) 167–176.
S. Ramya et al. / Applied Surface Science 256 (2010) 5108–5115 [47] A. Synytsya, J. Copikova, P. Matejka, V. Machovic, Carbohydr. Polym. 54 (2003) 97–106. [48] D. Lin-Vien, N.B. Colthup, W.G. Fateley, J.G. Grassetti, The Handbook of Infrared and Raman Characteristics Frequencies of Organic Molecules, Academic, Boston, 1991. [49] M. Harz, P. Rösch, K.D. Peschke, O. Ronneberger, H. Burkhardt, J. Popp, Analyst 130 (2005) 1543–1550. [50] U. Neugebauer, U. Schmid, K. Baumann, W. Ziebuhr, S. Kozitskaya, V. Deckert, M. Schmitt, J. Popp, ChemPhysChem 8 (2007) 124–137. [51] M.L. Laucks, A. Sengupta, K. Junge, E.J. Davis, B.D. Swanson, Appl. Spectrosc. 59 (2005) 1222–1228. [52] H. Zhao, B. Yuan, X. Dou, J. Opt. A: Pure Appl. Opt. 6 (2004) 900–905.
5115
[53] N.P. Ivleva, M. Wagner, H. Horn, R. Niessner, C. Haisch, Anal. Chem. 80 (2008) 8538–8544. [54] A. Sengupta, M.L. Laucks, N. Dildine, E. Darpala, E.J. Davis, J. Aerosol. Sci. 36 (2005) 651–664. [55] R.M. Jarvis, R. Goodacre, Anal. Chem. 76 (2004) 40–47. [56] N.P. Ivleva, M. Wagner, H. Horn, R. Niessner, C. Haish, Anal. Bioanal. Chem. 393 (2009) 197–206. [57] N. Gierlinger, M. Schwanninger, Plant Physiol. 140 (2006) 1246–1254. [58] H. Schulz, M. Baranska, R. Baranski, Biopolymers 77 (2005) 212–221. [59] I. Notingher, S. Verrier, S. Haque, J.M. Polak, L.L. Hench, Biopolymers 72 (2003) 230–240.
Contents lists available at ScienceDirect
Applied Surface Science journal homepage: www.elsevier.com/locate/apsusc
Detection of algae and bacterial biofilms formed on titanium surfaces using micro-Raman analysis S. Ramya ∗ , R.P. George, R.V. Subba Rao, R.K. Dayal Corrosion Science and Technology Division, Indira Gandhi Centre for Atomic Research, Kalpakkam 603102, India
a r t i c l e
i n f o
Article history: Received 21 December 2009 Received in revised form 17 March 2010 Accepted 17 March 2010 Available online 25 March 2010 Keywords: Raman spectroscopy Colloids Biofouling Biofilms Characterization
a b s t r a c t Biofouling is one of the major impediments in the use of titanium in sea-water cooled condensers of power plants, which is otherwise an excellent material with respect to corrosion resistance. Raman microscopic experiments were carried out on biofilms grown on titanium surfaces to find out the chemical composition of complex extracellular polymeric substances (EPS) in the biofilm. Though the spectral resolution of normal Raman experiments on these systems was very poor, it was improved when microSERS experiments were carried out using mono and bimetallic Ag and Cu colloids. It was observed that spatial distribution of polysaccharides was higher than that of proteins in algae biofilms formed on titanium matrix. Similar experiments were performed on laboratory cultured bacterial films of Pseudomonas aeruginosa. It was evidenced that algal and bacterial biofilms on titanium can be clearly distinguished with the help of Raman mapping coupled with SERS technique using bimetallic Ag/Cu colloids. © 2010 Elsevier B.V. All rights reserved.
1. Introduction Microorganisms in the aquatic systems are susceptible to form biofilms on metallic surfaces. The interaction of these films with metal surface facilitates biofouling [1–7], which is thought to be one of the major problems in cooling water systems of many industries. For instance, titanium suffers serious biofouling, which is otherwise an excellent material for corrosion resistance. The surface coverage and thickness of these films form on titanium are seldom uniform. Since biofouling checking measures involve biofilm characterization, proper monitoring of biofilm formation and the quantification of its extracellular polymeric substances (EPS) are essential to impact the mechanical cleaning and water treatment programs. In this regard, the present work concerns about the detection and the semi quantification of algae and bacterial biofilms on titanium surfaces using Raman microscopy. The application of Raman spectroscopy to complex biological systems is of great importance [8–12] due to highly specific vibrational spectra. However, biological systems suffer overlapping of various spectral signatures and lesser intensity because of their large fluorescence background. Quenching of the fluorescence signals was desirable to enhance and resolve the vibrational spectra of EPS components, and was achieved normally by surface enhanced Raman spectroscopy (SERS) [13–16]. Generally, SERS is a strong tool for the detection of materials present in lower concentration. The
∗ Corresponding author. Tel.: +91 44 27480121; fax: +91 44 27480121. E-mail addresses: [email protected] (S. Ramya), [email protected] (R.P. George). 0169-4332/$ – see front matter © 2010 Elsevier B.V. All rights reserved. doi:10.1016/j.apsusc.2010.03.079
vibrational spectra of the analyte molecules in the proximity of substrate metal colloids can be obtained in SERS experiments where local optical fields of metallic colloids yield enhanced Raman signals of the surrounding environment [17–21]. Generally, natural biofilms can be algae, bacteria or fungi and their cell wall is made up of microfibrillar elements that provide rigidity and strength. The most common components of the microfibrils are polysaccharides, proteins, glycoproteins, lipids and minerals [22]. As far as the algae (plant) cell wall is concerned, the foremost portion constitutes polysaccharides. In the case of bacteria, the cell wall is peptidoglycan or lipoproteinaceous, made from polysaccharides or lipids cross-linked by unusual peptides containing d-amino acids. In other words, polysaccharides are present as major components in algae cell wall whereas proteins are the chief component in bacterial cell wall. The factor that creates the distinction between algae and bacterial biofilms is their composition of the cell wall wherein the polysaccharides to protein ratio varies distinctly. Our earlier study based on chlorophyll estimation indicated higher algal density in the marine water biofilm formed on the titanium matrices [23,24]. However, fast and effective method for monitoring the biofouling was needed. Hence, the single analysis through Raman spectroscopy with simple sample preparation was considered for analyzing the molecular composition in biofilm components. In this regard, Raman imaging was adapted to find out the distribution ratio of protein to polysaccharide counterparts so as to confirm the algal dominance on the titanium surfaces. There are many reports on SERS studies for the characterization of various single cells, proteins, amino acids and bacteria [25–38]. However, the application of bimetallic Ag/Cu colloids as SERS substrate for
S. Ramya et al. / Applied Surface Science 256 (2010) 5108–5115
biofilm detection using laser Raman microscopy was not reported. The reason for selecting bimetallic Ag/Cu colloids over individual Ag and Cu colloids as SERS substrates for biofilm detection was explored in the present task. For the first time, a promising approach to investigate biofouling on titanium surfaces with SERS using mono and bimetallic Ag and Cu colloids was attempted and it was intended to identify the difference between algal and bacterial biofilms based on Raman mapping experiments coupled with SERS technique. Generally, a possible way to measure the relationship between SERS spectra and the nature of the colloid is to perform the measurements for a given simple molecule in presence of different colloids. However, the performance of colloids depends on various parameters such as laser wavelength, particle size and analyte molecules. Proper substrate colloid and excitation wavelength must be chosen for any simple analyte molecule. An extensive study was done by Zeiri and Efrima [21] to study the effect of excitation wavelength and the colloid milieu and they reported that the observation of spectral signatures from different biochemical components of the microorganisms will be facilitated by altering the type of colloid and the laser excitation wavelength. They also stated that the normal Raman protocol can be switched to resonance or surface enhanced Raman by changing the relative spectral contributions of the various cell components and different types of information can be obtained depending on the types of colloid. In this respect, we made an attempt to find out a suitable substrate colloid for biofouling quantification and suggested a use of Ag/Cu bimetallic colloid as a better substrate than individual Ag and Cu systems for biofouling studies on titanium surfaces. Moreover, all our Raman experiments were carried out using red excitation since both the substrate and analyte were shown better stability throughout the analysis under this excitation. 2. Materials and methods 2.1. Algae and bacterial biofilm samples Two different kinds of biofilms on titanium surfaces i.e., algae and laboratory cultured bacterial biofilms were considered for Raman analysis. Commercially pure titanium Grade-2 samples were used in this study. The Ti metal was bought in the form of sheets and the coupons of size 30 mm × 20 mm × 1 mm were cut from it. These specimens were pickled in an acid bath (HNO3 , 400 g/L + HF, 40 g/L + water) to remove any surface scales present. The pickled specimens were then cleaned ultrasonically in a detergent solution, washed in running tap water and finally rinsed in distilled water and air-dried. One set of titanium specimens were suspended in the outfall water from Madras Atomic Power Corporation (MAPS), Kalpakkam. The specimens were withdrawn after 1 week and it was clearly seen that there was a considerable biofilm growth on the titanium surfaces. Based on epifluorescence microscopy and chlorophyll estimation, the algal dominance in the natural biofilm was confirmed (results are not shown here). This algal biofilms were kept for further analysis by Raman microscopy. Another set of titanium specimens were exposed to the culture of Pseudomonas aeruginosa bacteria in 10% nutrient broth for 72 h and withdrawn for biofilm characterization. 2.2. Colloidal milieu The silver and copper colloidal suspensions of 4 × 10−4 M concentration were prepared using the citrate and ascorbic acid reduction methods [39–41], respectively. For the preparation of bimetallic (1:4 molar ratio) Ag/Cu colloidal particles, the copper seed colloidal particles were initially prepared by reducing the
5109
aqueous cupric nitrate using ascorbic acid in presence of poly vinylpyrrolidone (PVP) as a capping agent. The reaction time for the preparation of copper colloid was about 7–8 h [40], followed by the addition of silver nitrate and the reaction was continued for another 2 h to get a reddish brown colloidal suspension. The added ascorbic acid served as a reducing agent for both copper and silver nitrates. Also, the synthesis for copper and Ag/Cu colloids was carried out in an inert atmosphere. The obtained mono and bimetallic colloidal particles were characterized in terms of their respective surface plasmon bands (figure not shown here) using UV–visible absorption spectroscopy. The individual silver and copper colloids have characteristic surface plasmon bands at 420 and 590 nm, respectively. In the present study, the synthesized bimetallic alloy colloid formation was confirmed by the presence bands at 450 and 583 nm due to surface plasmonic oscillations of silver and copper colloids. The shift in the position of the bands when compared to pure colloids was due to the formation of bimetallic alloy colloid of both the silver and copper systems. 2.3. Raman protocol for biofilms The titanium coupons with the biofilms were immersed in the colloidal suspensions and their analysis in Raman microscopic scanning mode were carried out with an HR 800 Jobin Yvon Raman spectrometer equipped with 1800 grooves/mm holographic grating. He–Ne laser of 633 nm was used as an excitation source. The system consists of an Olympus optical microscope mounted at the entrance of the Raman spectrograph and the 10× long distance objective was used to focus the beam on the sample. The laser spot size focused on the surface was approximately 3 m in diameter and laser power was 5 mW at the sample. The slit width of the monochromator was 300 m, which corresponds to a resolution of 4 cm−1 . The back scattered Raman spectra were recorded using super cooled (<−110 ◦ C) 1024 × 256 pixels CCD detector over the range 80–4000 cm−1 . Raman spectra were collected at various points (at least 4) of each sample, and were observed to be reproducible. All the spectra were collected with 4s integration and 20 accumulations. Both integration time and number of accumulations were kept constant for all the experiments. The obtained Raman spectra were further baseline corrected for better comparison. The micro-SERS spectra taken in different colloidal systems showed some minor differences in their peak positions due to slight changes in sampling, morphology and the heterogeneity of the biofilms on titanium surfaces. The spectral maps were obtained by collecting spectra over 200 m × 200 m area with 4 s exposure time and 20 CCD accumulations. Data acquisition was done by a software package (Kaiser-Optical System). 3. Results and discussion 3.1. Normal Raman experiments on algae biofilms on titanium surfaces The main reason for choosing the He–Ne laser with 633 nm as an excitation source was to avoid the degradation and strong fluorescence background of most of the cellular components. Normal Raman experiments (NR) on these biofilms on titanium surfaces did not give much information with respect to EPS of the biofilm. However, some medium and lower intensity peaks were detected (Fig. 1a) in the region below 800 cm−1 and these peaks were assigned to deformations, tetrahedral, ring breathing and stretching vibrations from diatomaceous silicate components in the cell wall [42]. The observed silicate vibrations were supported by the fact that the silicon is the main component of the cell wall of groups of algae [43]. For instance, the cell wall of the some brown and green
5110
S. Ramya et al. / Applied Surface Science 256 (2010) 5108–5115
well resolved Raman spectra. When SERS experiments were carried out using silver colloids, the Raman spectra gained from surface plasmon enhancement. However, at longer excitation wavelengths (here 633 nm) such enhancements were reduced and weaker spectra were obtained owing to the decrease in the effectiveness of silver colloids on some of the components in the cell wall. When considering the chemisorptions of silver with proteins, the bonding can be achieved through Ag–N. In this regard, peaks at lower wave numbers were expected. In the present case, we did not observe lower wavenumber peaks around 200 cm−1 due to the use of longer excitation wavelength and citrate silver colloids [21]. Also, these features were strongly dependent on metal-adsorbate interactions [21]. The micro-SERS experiments using silver colloid gave enhancement of polysaccharide frequencies of algae biofilm. However, Raman signatures from the proteins were not enhanced. Hence, Raman mapping experiments using silver colloids were not carried out to find out the protein and polysaccharide distributions.
Fig. 1. Micro-Raman spectra of the algae biofilms on titanium surfaces; (a) represents the Raman spectra taken in the absence of colloid, (b) represents the Raman spectra taken in presence of Ag colloid, (c) represents the Raman spectra taken in presence of Cu colloid. Laser excitation was 633 nm using He–Ne laser, laser power = 5 mW, exposure time = 4 s, CCD accumulations = 20.
algae is enriched in silicon. Algae diatoms generally take silicon up as silicates [43]. 3.2. Micro-SERS experiments on algae biofilms on titanium surfaces in presence of Ag colloid Normal Raman experiments (NR) on these biofilms on titanium surfaces gave vibrations corresponding to silicates and it was difficult to resolve other spectral signatures with regard to organic components, even by increasing the exposure time and the number of CCD accumulations. Hence, the experiments were carried out in presence of silver colloids to trace out some of the major components. The intensity of micro-SERS spectrum was higher by many orders in magnitude than that of NR experiments. A tentative band assignment was performed by referring to Raman and SERS assignments for biological samples from the literature [25–38,44–59]. A strong peak at 1398 cm−1 must be due to COO− symmetric stretching vibrations from glycoproteins or polysaccharides [48]. The SERS spectra (Fig. 1b) with Ag colloid showed the presence of polysaccharides owing to the presence of some smaller intensity bands 300–600 cm−1 [16,45,56]. Also, the peaks around 970–998 cm−1 were due to the presence of O–CH3 stretching from polysaccharide units [47,57]. In addition to that, a small peak at 1068 cm−1 was seen and attributed to C–O and C–C stretching frequencies of polysaccharides [44,45]. A small intensity peak at 889 cm−1 was assigned to C–C and C–O–C glycosidic linkages [46,49,50]. A strong band was identified around 2900 cm−1 due to C–H stretching vibrations from both polysaccharides and proteins [12,47]. In addition to that, medium intensity peaks were observed at 1504 [56,58] and 1595 cm−1 [44,50,56] due to C C stretching vibrations and aromatic ring stretching frequencies, respectively. The peak at 1660 cm−1 [57] can be assigned to ring conjugated C C stretching vibrations. Though red excitation (633 nm) was used, the micro-SERS spectrum from silver colloid was superimposed on a background, due to the presence of high fluorescence nature of various components in the algae biofilm. Also, features from the protein components were not clearly observed. In addition to that, the silicate peaks observed in NR experiments were not detected in micro-SERS due to the overlapping of silicates and polysaccharide frequencies. It was expected that quenching of fluorescence to tolerable levels by silver colloids will allow the acquisition of
3.3. Micro-SERS experiments on algae biofilms on titanium surfaces in presence of Cu colloid One of the principal features that we wanted to examine was the effect of types of colloids on the SERS spectra. Replacing silver by copper changed the optical properties (copper plasmons are red shifted to 170 nm compared to silver particles) of analyte and substrate. Very strong and informative spectra were obtained with 633 nm excitation using copper colloids. Possible reason that, the excitation wavelength was closely in resonance with copper colloids than silver colloids. Fig. 1c depicts the micro-SERS spectra of algae biofilm in presence of copper colloid, which shows similar features as that of silver colloids in terms of band positions. However, major differences were observed below 800 cm−1 and the Raman signals were about 10 times higher than those with silver colloids and show many additional features with regard to proteins. It was clearly evidenced that copper colloid was found to be a better candidate in resolving the signal of both polysaccharides and proteins of algae biofilm when compared to silver colloid. However, the major difficulty in dealing with the copper colloid was its stability. Especially, when Raman mapping experiments were carried out, copper colloids readily got oxidized due to longer laser irradiation time, and no Raman signals were observed after sometime. Hence, the idea of getting spatial distribution of particular component in the biofilm was merely lost. Though copper colloid gave larger enhancement and better resolution, it suffered greater instability compared to silver colloid. It was concluded that copper colloidal dispersion is a good substrate for the detection of cellular components but not for their quantification. 3.4. Micro-SERS experiments on algae biofilms on titanium surfaces in presence of Ag/Cu bimetallic colloid In the present investigation, we wanted to distinguish between algal and bacterial biofilms based on their polysaccharide and protein distributions. Though micro-SERS with copper colloids gave better results, in order to get quantification in terms of spatial distribution using Raman mapping, the colloidal milieu was modified from Cu to Ag/Cu bimetallic colloids of better stability and the resolving power of both the individual silver and copper colloids was also retained. Fig. 2a shows the micro-SERS spectra of algae biofilm using bimetallic Ag/Cu colloids and the corresponding frequencies were mentioned in Table 1. Even though the intensity of the SERS spectra with this bimetallic milieu was slightly lesser than that of pure copper colloid, the spectral resolution with regard to proteins and polysaccharides was significantly improved. Fig. 3a shows the average micro-SERS spectra taken from the algae biofilm, wherein the presence of dominant polysaccha-
S. Ramya et al. / Applied Surface Science 256 (2010) 5108–5115
5111
Table 1 Assignments of the peaks (cm−1 ) for the micro-SERS spectra of natural algae biofilm in various types of colloids using 633 nm excitation. Peak position (cm−1 ) Ag colloid
Peak position (cm−1 ) Cu colloid
417
356 489 555
619
636 716 755 801
Peak position (cm−1 ) Ag/Cu colloid
560 670
804
889 970 998
910 973 1012
1067
1082
1160
1174
1271
1256 1332 1375
1137
1356
1504
1448; 1452
1595 1660
1580
1570
2901 Other small
2930 peaks
2976
Tentative peak assignments Carbohydrates [16,45,56] Carbohydrates [16,45,56] C–O–C glycosidic ring deformation of polysaccharides and COO− wagging and C–C skeletal [16,44] COO− bending [16,51] CH2 ; Adenine ring stretching [44] Ring breath of proteins [44,56,59] Ring breath of proteins and O–P–O stretching from nucleic acids [44]; Different C–N stretching from proteins [44,58,59] C–C stretching and C–O–C glycosidic linkage [44,59] Different C–N stretching from proteins [44] O–CH3 stretching [47] Carbohydrates [57] C–CH3 deformation from pigments [58,56] C–O; C–C str; C–OH deformation from carbohydrates [44,46] C–C stretching; C–O ring breathing and asymmetric stretching [45,56] CH2 twisting and C–H in-plane deformation [56]; Amide III [44,56] ı(CH) of proteins and carbohydrates [56] COO− symmetric stretching [48,56] COO− symmetric stretching; ␦ (CH2 ) scissoring and CH2 deformation [48] ı(CH2 ) scissoring and CH2 deformation of proteins, lipids and carbohydrates [48,58] C C stretching from pigments [56,58] G-A ring stretching [44,50,56] Ring conjugated C C stretching [57] Ring conjugated C C stretching; C O stretching from lipids or carbohydrates [47,56] Amide I [44,49,56] CH and CH2 stretching [12,47] Unassigned
tribution of polysaccharides than proteins in algae biofilm. The Raman map of polysaccharides was taken from 990–1100 cm−1 (C–O and C–C stretching of polysaccharides). Similarly the map of proteins was taken from 770 to 850 cm−1 (ring breathing and different C–N stretching vibrations of proteins). The mapping experiments were repeated for other spectral regions of polysaccharides and proteins. The observed results were reproducible and in good agreement with the earlier findings. 3.5. Micro-SERS experiments on Pseudomonas aeruginosa bacterial films on titanium surfaces in presence of bimetallic Ag/Cu colloid
Fig. 2. (a) represents the micro-SERS spectra extracted from the Raman map of algae biofilm on titanium surface in presence of bimetallic Ag/Cu colloid. (b) represents the micro-SERS spectra extracted from the Raman map of bacterial biofilm on titanium surface in presence of bimetallic Ag/Cu colloid. Laser excitation was 633 nm using He–Ne laser, laser power = 5 mW, exposure time = 4 s, CCD accumulations = 20.
ride, glycoproteins and lipid frequencies around 1000–1500 cm−1 was identified. Fig. 4a depicts the light microscopic image of the algae biofilm formed on the titanium surface. Raman mapping experiments were performed on the selected region of the algae biofilm. The Raman spectral maps for proteins and polysaccharides (Fig. 4b–d) were obtained by collecting the spectra with 4 s exposure time. The mapping experiments showed the greater dis-
Due to the good performance of bimetallic Ag/Cu colloid on algae biofilm analysis, the colloidal milieu was expected to show similar trend for bacterial biofilm. In this regard, NR and micro-SERS experiments were carried out on laboratory cultured P. aeruginosa bacterial film. The resolution of NR spectrum of P. aeruginosa was very poor and the identification of components in the biofilm was impossible. However, micro-SERS of the P. aeruginosa film in presence of bimetallic Ag/Cu colloid was distinct and well resolved (Fig. 2b). The obtained frequencies for the P. aeruginosa film (Table 2) were in good agreement with the literature findings [51]. Table 2 depicts the tentative assignments of the peaks seen in the micro-SERS spectra of P. aeruginosa. The Raman peaks around 500–1300 cm−1 region were due to various groups including glycosides, nucleic acids, proteins, polysaccharides and unsaturated fatty acids that are present in bacterial cell [51]. The peak at 1655 cm−1 was due to amide I [16]. We did not get specific phenylalanine peak at 1004 cm−1 as reported for many bacterial cultures due to the overlapping of other spectral frequencies. Another pos-
5112
S. Ramya et al. / Applied Surface Science 256 (2010) 5108–5115
Fig. 3. (a) represents the average Raman spectra taken from the algal biofilm on Ti surface. (b) represents the average Raman spectra taken from the bacterial biofilm on Ti surface. Laser excitation was 633 nm using He–Ne laser, laser power = 5 mW, exposure time = 4 s, CCD accumulations = 20. The average spectra were collected over 200 m × 200 m area.
Fig. 4. Distribution of polysaccharides and proteins in algae biofilm formed on titanium matrix. (a) represents the light microscopic image of the algae biofilm obtained by Olympus Raman microscope using 10× long distance objective. The square box inside (a) represents the selected region of interest where Raman mapping was carried out. In (a) dark region corresponds to algae biofilm whereas the bright region corresponds to Ti surface. (b) represents the Raman map of polysaccharides and the mapping region covers 990–1100 cm−1 with a central peak position at 1082 cm−1 . (c) Represents the Raman map of polysaccharides and the mapping region covers 600–700 cm−1 with a central peak position at 670 cm−1 . (d) represents the Raman map of proteins and the mapping region covers 770–850 cm−1 with a central peak position at 809 cm−1 . Table 2 Tentative peak assignments [48] (cm−1 ) for micro-SERS spectra of P. aeruginosa in bimetallic Ag/Cu colloid using 633 nm excitation. Band (cm−1 )
Assignments
360 487 639 767
Carbohydrate [16,45,51,56]. Carbohydrate [16,45,51,56]. C–O–C glycosidic ring deformation of polysaccharides; COO− wagging and C–C skeletal [20,53]. Ring breath of proteins [44,56,59]. Different C–N stretching from proteins [51]. Polysaccharides and C–C, C–O, C–O–H deformation [51]. N–H, C–N, amide III (protein) [51,56]. CH2 bending from proteins and lipids [52,56]. G–A ring stretching [44,50,56]. Amide I [16]. C–H stretching [47,12]. Unassigned.
876 1052 1280 1473 1571 1655 2900, 2953 Other small peaks
sibility is that chemical effects may control the production of SERS-active centers in specific sites within the cellular components [21]. Amides III bands at 1220–1320 cm−1 are absent in the SERS spectra of algae film due to the broad overlapping of other spectral signatures in that region (Fig. 2a). However, it appeared as a weak band in the bacterial SERS spectrum (Fig. 2b). The Raman mapping experiments were carried out on P. aeruginosa bacterial films in bimetallic Ag/Cu colloid to see the spatial distribution of polysaccharides and proteins. Experimental parameters for Raman mapping experiments were kept as same as it was done for algae biofilms. The choice of the frequencies for mapping should be the same for both algae and bacterial biofilms. However, in the present Raman investigation of bacterial biofilms, we did not trace out proper Raman signals for proteins in the region 770–850 cm−1 (chosen for algae mapping). But, distinct signal was obtained at 876 cm−1 and was accounted for proteins based on the literature. Since our aim was to see the differences in the protein ratios in algal and bacterial biofilms, the variations in the spectral positions (due to different Raman active modes) of proteins were considered less
S. Ramya et al. / Applied Surface Science 256 (2010) 5108–5115
5113
Fig. 5. Distribution of polysaccharides and proteins in bacterial biofilm formed on titanium matrix. (a) represents the light microscopic image of the bacterial biofilm obtained by Olympus Raman microscope using 10× long distance objective. The square box inside (a) represents the selected region of interest where Raman mapping was carried out. (b) represents the Raman map of proteins and the mapping was carried out with respect to 1280 cm−1 corresponding to amide III band. (c) represents Raman map of proteins and the mapping was carried out with respect to 876 cm−1 corresponding to different C–N stretching of proteins. (d) represents Raman map of polysaccharides and the mapping was carried out from 990 to 1100 cm−1 corresponding to C–O and C–C stretching of polysaccharides. (e) represents Raman map of polysaccharides and the mapping was carried out from 600 to 670 cm−1 corresponding to C–O–C vibrations of glycosidic ring deformations.
important. In order to overcome this disadvantage, Raman mapping of proteins was repeated at 1280 cm−1 (amide III) for verifying the mapping results. For bacterial biofilms, the features corresponding to proteins and polysaccharides were clearly seen. Hence, the mapping was carried out with respect to those frequencies without any uncertainty. However, the mapping experiments on algal biofilms led to a small confusion in the selection of proper protein and polysaccharide vibrations. Because, most of the protein vibrations including amide bands were absent due to the broad overlapping of other spectral signatures. Hence, the region 770–850 cm−1 was chosen for mapping the protein signatures of algal biofilm. The strong and prominent peaks at 1356 and 1570 cm−1 were not considered for algal mapping. Because, the region 1350–1400 cm−1 was attributed to the presence of COO− symmetric stretching; ı(CH2 ) scissoring and CH2 deformation from proteins, lipids and carbohydrates [48,56,58] and the peaks around 1500 cm−1 were due to aromatic ring stretching frequencies [56,58]. It was observed from the bacterial mapping experiments (Fig. 5) that the spatial distribution of polysaccharides was less compared to that of proteins, which is a characteristic feature of bacterial cell wall. Similar mapping was carried out for other polysaccharides and protein spectral regions and the observed distribution of polysaccharides and proteins ratios were consistent and good agreement with the previous results. The bacterial spectrum was in less coin-
cidence with the algae spectrum. In the Raman spectra of the algae biofilm, signals due to proteins and amino acids were not prominent. Rather, strong signals with comparatively higher background were obtained around 1350–1400 and 1580–1600 cm−1 . The spectral region 1350–1400 cm−1 was attributed to the presence of COO− symmetric stretching; ı(CH2 ) scissoring and CH2 deformation from lipids and carbohydrates [48,56,58]. Some medium intensity peaks were observed around 1500 cm−1 due to C C stretching vibrations and aromatic ring stretching frequencies [56,58]. The Raman mapping experiments on the algae biofilms showed more polysaccharide distribution whereas the bacterial biofilm showed greater protein distributions. The differences between these two biofilms were clearly observed from the Raman mapping experiments. Though the algal and bacterial biofilms showed the presence of basic proteins and polysaccharide signatures as expected, an interesting feature was observed with regard to C–H stretching frequencies around 2900 cm−1. In the Raman spectra of algal biofilm, single peak was observed around 2900 cm−1 . In the case of bacteria, a strong doublet was detected in the same wave number region due to C–H and CH3 stretching vibrations from proteins and lipids. Since Gram-negative P. aeruginosa bacteriam has a characteristic lipoproteinaceous cell wall, these vibrations were probably aroused from the proteins and the lipids of the cell wall. As per literature, algae cell wall consists more of polysaccharides whereas bacterial mem-
5114
S. Ramya et al. / Applied Surface Science 256 (2010) 5108–5115
brane contains more protein (peptidoglycan) units. This is the basis for the present study to detect the micro levels of algae and bacteria on the titanium condensers using SERS coupled with Raman microscopy. Generally, Raman map gives the intensity distributions of one particular spectral range from which the changes in the composition of a specific component can be determined. In our present study, we selected lesser number of data points (36 points) and hence, the image obtained was coarse and not exactly matching with the light microscopic image. Also, the Raman map represented the image distribution of one particular wave number range (correspond to one component) whereas the light microscopic image depicted the distribution of all the components. It was assumed that different protein to polysaccharide ratio will be reflected in the Raman mapping. Since colloids enhance the Raman signals of molecular components in their close proximity even when it is not the major constituent, the titanium coupons with the biofilms were immersed in the colloidal suspensions throughout the Raman analysis. When the Raman experiments were carried out for pure P. aeruginosa bacterial culture, it was observed that both polysaccharide and protein frequencies were enhanced and the idea of doing mapping experiments to find out the protein to polysaccharide distribution was achieved. Hence, the Raman microscopy coupled with SERS using bimetallic Ag/Cu colloid was identified as an effective tool for differentiating algae and bacterial biofilms in order to see the impact of biofouling on Ti surfaces.
lurgy and Materials Group, Dr. A.K. Arora and Dr. M. Rajalakshmi, Materials Science Division, and Dr. Baldev Raj, Director, Indira Gandhi Centre for Atomic Research, Kalpakkam, during the course of this work. The authors thank Dr. M.P. Srinivasan and Mr. P. Chandra Mohan, BARCF, Kalpakkam for their timely help and suggestions. The support of K.S. Krishnan research fellowship is also acknowledged. References [1] [2] [3] [4] [5] [6] [7] [8] [9] [10] [11] [12] [13]
[14]
4. Conclusions The Raman microscopic experiments were carried out on algae and bacterial biofilms grown on titanium surfaces in order to have a proper understanding of the chemical composition of the complex extracellular polymer substances (EPS) in the biofilm. The salient conclusions are as follows: (a) normal Raman experiments on algae biofilms gave information only about silicates and the spectral details corresponding to organic components in the biofilm were not identified due to high complexity of the biofilm. (b) Micro-SERS experiments using silver colloids on these biofilms clearly indicated the contributions of polysaccharides. However, the significance of protein was not identified distinctly. (c) Resolution with regard to both polysaccharides and proteins was remarkably increased when similar micro-SERS experiments were performed using copper colloids. Conversely, it was observed that copper colloid was good for identification of the components but not for their quantification due to their lack of stability while performing mapping experiments. (d) Similar experiments using bimetallic Ag/Cu colloids showed good results and hence, it was easy to resolve the signals of most of the algae biofilm components. The Ag/Cu colloid showed better stability than individual silver and copper colloids and the Raman mapping experiments were conducted to explore the spatial distribution of biofilm components. (e) Raman mapping experiments using bimetallic Ag/Cu colloids on algae biofilm showed greater the distribution of polysaccharides than proteins. (f) The effectiveness of bimetallic Ag/Cu colloid was checked even for the laboratory cultured P. aeruginosa bacterial films. It was concluded that Raman microscopy coupled with SERS technique was an effective tool for the quantification of algae and bacterial biofilms on titanium surfaces in order to improve the control measures of biofouling of condenser materials in power plants.
[15] [16] [17] [18] [19] [20] [21] [22] [23] [24] [25] [26] [27] [28] [29] [30] [31] [32] [33] [34] [35] [36] [37] [38] [39] [40] [41] [42] [43]
Acknowledgements [44]
The authors wish to acknowledge the support and encouragement extended by Dr. P.R. Vasudeva Rao, former Director, Metallurgy and Materials Group, Dr. T. Jayakumar, Director, Metal-
[45] [46]
W.G. Characklis, K.E. Cooksey, Adv. Appl. Microbiol. 29 (1983) 93–138. B. Little, P. Wagner, F. Mansfeld, Mater. Rev. 36 (1991) 253–258. R.P. George, D. Marshall, R.C. Newman, Corros. Sci. 45 (2003) 1999–2015. X. Shi, R. Avci, M. Geiser, Z. Lewandowski, Corros. Sci. 45 (2003) 2577–2595. K. Pedersen, Water Res. 24 (1990) 239–243. O.M. Vaisanen, E.L. Nurmiaho-Lassila, S.A. Marmo, M.S. Salkinoja-Salonen, Appl. Environ. Microbiol. 60 (1994) 641–653. R.P. George, P. Muraleedharan, N. Parvathavarthini, H.S. Khatak, T. Rao, Mater. Corr. 51 (2000) 213–218. N.J. Kline, P.J. Treado, J. Raman. Spectrosc. 28 (1997) 119–124. D. Naumann, H. Helm, Labischinski, Nature 351 (1991) 81–82. D. Naumann, S. Keller, D. Helm, C. Schultz, B. Schader, J. Mol. Struct. 347 (1995) 399–405. G.J. Puppels, F.F.M. Demul, C. Otto, J. Greve, M. Robertnicoud, D.J. Arndtjovin, T.M. Jovin, Nature 347 (1990) 301–303. M.F. Escoriza, J.M. VanBriesen, S. Stewart, J. Maier, P.J. Treado, J. Microbiol. Methods 66 (2006) 63–72. L.-P. Choo-Smith, K. Maquelin, T.V. Vreesuijk, H.A. Bruining, G.J. Puppels, N.A. Ngo Thi, C. Kirsciner, D. Naumann, D. Ami, A.M. Villa, F. Orsini, S.M. Doglia, H. Lamfarrej, G.D. Sockalingam, M. Manfait, P. Allouch, H.P. Endltz, Appl. Environ. Microbiol. 67 (2001) 1461–1469. K. Maquelin, L.-P. Choo-Smith, T.V. Vreeswijk, H. Endtz, B. Smith, R. Bennet, H. Bruining, G. Pupples, Anal. Chem. 72 (2000) 12–19. A.C. Williams, H.G.M. Edwards, J. Raman. Spectrosc. 25 (1994) 673–677. K.C. Schuster, I. Reese, E. Orlaub, J.R. Gapes, B. Lendi, Anal. Chem. 72 (2000) 5529–5534. K. Kneipp, Y. Wang, H. Kneipp, L. Perlman, I. Itzkan, Phys. Rev. Lett. 78 (1997) 1667–1670. S. Nie, S.R. Emory, Science 275 (1997) 1102–1106. C.D. Keating, K.K. Kovaleski, M.J. Natan, J. Phys. Chem. B 102 (1998) 9414–9425. S. Lefrant, J.P. Buisson, J. Schreiber, O. Chauvet, M. Baibarac, I. Baltog, Synth. Met. 139 (2003) 783–785. L. Zeiri, S. Efrima, J. Raman. Spectrosc. 36 (2005) 667–675. B. Alberts, D. Bray, J. Lewis, M. Raff, K. Roberts, J.D. Watson, Molecular Biology of the Cell, 2nd ed., Garland Publishing, New York, 1989. J. Gopal, B.V.R. Tata, R.P. George, P. Muralidharan, R.K. Dayal, Surf. Eng. 24 (2008) 447–451. J. Gopal, R.P. George, P. Muraleedharan, H.S. Khatak, Biofouling 20 (2004) 167–173. G.D. Sockalingum, H. Lamfarraj, P. Pina, A. Beljebbar, P. Allouch, M. Manfait, Spectrosc. Biol. Mol. New Dir. 8 (1999) 599–600. R. Keir, D. Sadler, W.E. Smith, Appl. Spectrosc. 56 (2002) 551–559. S. Farqharson, A.D. Gift, P. Maksymiuk, F.E. Inscore, Appl. Spectrosc. 58 (2004) 351–354. L.A. Gearheart, H.J. Ploehn, C.J. Murphy, J. Phys. Chem. B 105 (2001) 12609–12615. E. Podstawka, Y. Ozaki, L.M. Proniewicz, Appl. Spectrosc. 58 (2004) 570–580. E. Podstawka, Y. Ozaki, L.M. Proniewicz, Appl. Spectrosc. 58 (2004) 581–590. E. Podstawka, Y. Ozaki, L.M. Proniewicz, Appl. Spectrosc. 58 (2004) 1147–1156. L. Zeiri, B.V. Bronk, Y. Shabtai, J. Czege, S. Efrima, Colloids Surf. 208 (2002) 357–362. R.M. Jarvis, A. Brooker, R. Goodacre, Anal. Chem. 76 (2004) 5198–5202. A. Shamsaie, J. Heim, A.A. Yanik, J. Irudhayaraj, Chem. Phys. Lett. 461 (2008) 131–135. R. Patzold, M. Keuntje, A.A. Von Anlften, Anal. Bioanl. Chem. 386 (2006) 286–292. P. Rosch, M. Schmitt, W. Kiefer, J. Popp, J. Mol. Struct. 661 (2003) 363–369. K.C. Schuster, E. Urlaub, J.R. Gapes, J. Microbiol. Methods 42 (2000) 29–38. D. Zhang, Y. Xie, M. Mrozek, C. Ortiz, J. Davisson, D. Ben-Amotz, Anal. Chem. 75 (2003) 5703–5709. P.C. Lee, D. Meisel, J. Phys. Chem. 86 (1982) 3391–3395. W. Yu, H. Xie, L. Chen, Y. Li, C. Zhang, Nanoscale Res. Lett. 4 (2009) 465–470. Y. Wang, P. Chen, M. Liu, Nanotechnology 17 (2006) 6000–6006. P. Yuan, H.P. He, D.Q. Wu, D.Q. Wang, L.J. Chen, Spectrochim. Acta Part A 60 (2004) 2941–2945. Ji. Zhen-Gang, Water quality and eutrophication, in: Hydrodynamics and Water Quality: Modeling Rivers, Lakes and Estuaries, John Wiley & Sons, Inc., Hoboken, NJ, 2008, p.289. K. Maquelin, C. Kirschner, L.-P. Choo-Smith, N. van den Braak, H.P. Endtz, D. Naumann, G.J. Puppels, J. Microbiol. Methods 51 (2002) 255–271. K. De Gussem, P. Vandenabeele, A. Verbeken, L. Moens, Spectrochim. Acta Part A, 61 A (2005) 2896–2908. J.T. Pelton, L.R. McLean, Anal. Biochem. 277 (2000) 167–176.
S. Ramya et al. / Applied Surface Science 256 (2010) 5108–5115 [47] A. Synytsya, J. Copikova, P. Matejka, V. Machovic, Carbohydr. Polym. 54 (2003) 97–106. [48] D. Lin-Vien, N.B. Colthup, W.G. Fateley, J.G. Grassetti, The Handbook of Infrared and Raman Characteristics Frequencies of Organic Molecules, Academic, Boston, 1991. [49] M. Harz, P. Rösch, K.D. Peschke, O. Ronneberger, H. Burkhardt, J. Popp, Analyst 130 (2005) 1543–1550. [50] U. Neugebauer, U. Schmid, K. Baumann, W. Ziebuhr, S. Kozitskaya, V. Deckert, M. Schmitt, J. Popp, ChemPhysChem 8 (2007) 124–137. [51] M.L. Laucks, A. Sengupta, K. Junge, E.J. Davis, B.D. Swanson, Appl. Spectrosc. 59 (2005) 1222–1228. [52] H. Zhao, B. Yuan, X. Dou, J. Opt. A: Pure Appl. Opt. 6 (2004) 900–905.
5115
[53] N.P. Ivleva, M. Wagner, H. Horn, R. Niessner, C. Haisch, Anal. Chem. 80 (2008) 8538–8544. [54] A. Sengupta, M.L. Laucks, N. Dildine, E. Darpala, E.J. Davis, J. Aerosol. Sci. 36 (2005) 651–664. [55] R.M. Jarvis, R. Goodacre, Anal. Chem. 76 (2004) 40–47. [56] N.P. Ivleva, M. Wagner, H. Horn, R. Niessner, C. Haish, Anal. Bioanal. Chem. 393 (2009) 197–206. [57] N. Gierlinger, M. Schwanninger, Plant Physiol. 140 (2006) 1246–1254. [58] H. Schulz, M. Baranska, R. Baranski, Biopolymers 77 (2005) 212–221. [59] I. Notingher, S. Verrier, S. Haque, J.M. Polak, L.L. Hench, Biopolymers 72 (2003) 230–240.