Studies of infrared (IR) spectra of various types of microorganisms (bacteria, yeast, fungi, microalgae, viruses) commenced as early as in the middle of the 20th century. Note that in a review on this topic, Norris (1959) [1] pointed to the main difficulties which, in addition to the high cost of the equipment and strong absorption of water traces contained in microbiological samples, generally comprised the impractical and laborious methodology, especially for the purpose of identifying microorganisms. For the IR spectroscopic study of viruses, their separation from the accompanying biological material was indicated as the main difficulty [1]. It should be noted that the IR spectroscopic equipment of that period had a low sensitivity and resolution, while later, from the 80s of the XX century, the situation was improved by the advent of Fourier transform IR (FTIR) spectrometers. The latter greatly contributed to the development of this research field [2, 3], which is still ongoing.

One of the most valuable, professional and highly informative guides for both specialists and beginners who apply the FTIR spectroscopic technique in microbiology is, in our opinion, up to now still the classic encyclopaedic review chapter by D. Naumann [3] first published in 2000. It contains a large number of examples, experimental data, as well as an informative discussion of the essence of IR spectroscopy, its capabilities and the main characteristics observed using this technique. Among the enormous and annually increasing number of microbiological studies using FTIR spectroscopy within recent years, a series of review papers can be pointed out. Some of them, when considering various instrumental techniques for studying particular microbiological objects, in most cases include fairly generalised and usually limited information about the IR spectroscopy technique (in different modes of measurement) [4–11]. Information of a different kind, most often more detailed, is presented both by reviews on various aspects of the application of FTIR spectroscopy in microbiology [12–17] and general reviews on various aspects of the application of the technique to biological objects (see, e.g., [18, 19]). Of particular interest are reviews of data on the FTIR spectroscopic study of individual types of bio(macro)molecules that make up a significant part of any microbiological systems, primarily proteins [20–24], sugars, oligo- and polysaccharides [25], lipids [24, 26]. These review articles, together with individual professionally made experimental reports, constitute a valuable “database” for researchers interpreting IR spectra of their own microbiological samples.

From the viewpoint of the FTIR spectroscopic methodology, review-type publications describing specialised measurement protocols for biological systems can be considered to some extent close to the scope of this article (see, e.g., [27–31], as well as chapters 9, 11, 15, 16 in the monograph [32]). Note that, although some of these protocols include IR spectroscopic analysis of biological tissues [29, 31], some essential methodological details of biospectroscopic analysis are largely common, especially for such complex supramolecular objects as microbial cells and biofilms. The very fact of the regular appearance of a number of such guides indicates the complexity and diversity of methodological approaches used in FTIR spectroscopy when dealing with similar biological samples as well as the relevance of this topic.

In this work, we did not aim to consider the main reports published even in recent years on the use of FTIR spectroscopy in microbiological studies—their number is too large, and the level of the presented materials, with regard to both methodology and the interpretation of spectroscopic data, varies greatly. Instead, some of the most significant, from the viewpoint of the authors, specific features of the FTIR spectroscopic methodology and the interpretation of experimentally obtained spectroscopic data, as applied to the analysis of microbiological objects, are discussed using typical examples from the results published in the specialised scientific literature, including the authors' publications. Also considered are some typical inaccuracies and errors that occur even in articles published by fairly high-rank specialised international journals. The information presented supplements the reviews discussed above available in the literature and will be useful to researchers who use the FTIR spectroscopic technique in microbiological analyses.

1 SPECIFICITIES OF THE FTIR SPECTROSCOPIC METHODOLOGY IN MICROBIOLOGICAL ANALYSES

One of the main factors that can hinder studying the relative content and structure of biomacromolecules in microbial cells (their general composition is given in Table 1) by FTIR spectroscopy is a significant content of water in samples, which is common for native biological materials, as water molecules strongly absorb in the corresponding IR spectral regions [33, 34]. In particular, the most significant is a noticeably broadenedFootnote 1 rather intense absorption band in the region of bending (scissoring) vibrations δ(H–O–H) at about 1640 cm–1, overlapping with the amide I region of vibrations of peptide bonds of proteins with different types of secondary structure (bands in the region of ~1620–1690 cm–1) [20, 35, 36]. The exclusion of the contribution from water in this case is a well-known problem and requires the use of special approaches, strict standardisation of sample parameters and measurement modes [35–40]. In the case of microbiological samples, this is not always easily achievable, also when using special equipment and methodology, for example, microfluidics and synchrotron FTIR spectromicroscopy [41]. While using the IR spectroscopic methodology of attenuated total reflectance (ATR) [35–37, 40], in studying aqueous suspensions of bacterial biomass or biofilms (as in the case of studying, e.g., aqueous solutions of proteins), one uses subtraction of the contribution of water obtained by a control measurement of the IR spectrum of the water layer. In this case, the accuracy of measurements can decrease owing to a significantly higher intensity of water absorption compared to the intensity of the bands of the biomaterial studied [35].

Table 1.   General biomacromolecular composition (in % of dry biomass) of microbial cells – prokaryotes (bacteria) and eukaryotes (yeast) (according to data from [3])
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Another approach to reduce the contribution of water contained in the sample is to dry the biomass of cells (or biofilms) separated from the culture medium [30, 42–45]. The applicability of this method is determined by the fact that, as shown by numerous studies, the native structure and functional activity of biomacromolecules and supramolecular structures may not change significantly under certain conditions; in this case, microbial cells reversibly transfer into a physiologically inactive (dormant) form [46–49]. As is known, dried microbial preparations (including those obtained by lyophilisation) can be stored for a sufficiently long time, and the cells in them remain alive, which is widely used in various fields of biotechnology [50–52]. However, it should be noted that upon drying, for some microorganisms, redox processes can occur, including those which are destructive, especially during storage of dry preparations [53]. In such cases, FTIR spectroscopy (along with other instrumental nondestructive techniques) can be very informative for a comparative analysis of the total macromolecular composition of dry cells and the consequences of these processes [45, 47, 54–56].

Dried samples of microbial biomass can be studied by FTIR spectroscopy both in the ATR mode and in the transmission or diffuse reflection (the commonly accepted term is “diffuse reflection infrared Fourier transform”; DRIFT) modes [3, 30, 42–45, 54, 56]. The DRIFT methodology, although it requires a special accessory to the spectrometer, allows for using dry biomass practically without any additional sample preparation and in a small amount (including that without using a halide matrix (KBr) for pressing sample pellets). In this case, a comparative analysis of the intensity and position of the bands (generally coinciding with those of the absorption bands owing to the fact that IR radiation, diffusely reflected from the surface of a powdered or surface-roughened material and “collected” by a special concave mirror, contains information about its absorption in the surface layer) gives largely the same structural and quantitative information as for the traditional absorption mode [2, 43, 54, 57].

It has been experimentally documented that for studying dry microbial biomasses by FTIR spectroscopy, lyophilisation [30, 50] is not necessary. To perform a sufficiently informative analysis, it is quite acceptable to dry the biomass, washed from the components of the culture medium, in air (at room temperature, and possibly in a desiccator with a water-absorbing agent to speed up the process, or in a drying chamber with a slight heating, which should not exceed ca. 45…50°C to exclude the processes of thermal denaturation of cellular proteins) [3, 27, 30, 42–48, 54, 57, 58]. It should be noted that the stage of washing the cell biomass or biofilm from the components contained in the nutrient medium (in most cases these are carbon sources—carboxylic acid salts or yeast extract, as well as inorganic salts (including phosphates) of potassium, sodium, ammonium and a small amount of trace elements—iron, calcium, magnesium, molybdenum, etc.) is significant. As shown in [59], to eliminate the contribution of the bands of these components to the IR spectrum of dry biomass, it is necessary to use more than one stage of its washing with physiological saline (the latter should rather be used instead of water to avoid osmotic stress for cells). In addition, the duration of drying (depending on its conditions) should be sufficient (for not too small amounts of biomass, drying “to a constant weight” is commonly used).

Figure 1 illustrates the degree of reproducibility of IR spectra of a bacterial biomass (several milligrams were used) which was dried in air at 45°C for 1.5 h (which leads to a visually practically dry sample) and for 23 h (according to the data reported in [60]; to compare the peak intensities, all spectra obtained in parallel measurements have been normalised to the intensity of the amide I band of cellular proteins with a maximum at ~1650 cm–1). The results show that the variants of the samples dried for 1.5 h (Fig. 1a) differ noticeably (the maximum differences in band intensities reach 20 to 39% in different regions of the spectrum), which indicates the presence of various residual traces of water in these samples and its unequal influence on the absorption bands of different functional groups, including the region of their overlap. As is known [30, 33, 34, 37–40, 58], water in the condensed phase has a strong absorption in the region of stretching vibrations of hydrogen-bonded O–H groups (a broad asymmetric absorption region at ~3500–2500 cm–1; in the same region, all OH groups of biomacromolecules, primarily polysaccharides, also absorb) and below 1000 cm–1 (a broad intensive band with a maximum in the region of 600–800 cm–1; librational vibrations of H2O). We also mentioned above the role of a moderately intensive absorption band in the region of bending (scissoring) vibrations δ(H–O–H) at about 1640 cm–1, overlapping with the amide I region of vibrations of peptide bonds in proteins, which in this case introduces additional differences in spectra when studying an insufficiently dried sample. (Note, however, that with a much thinner layer of biomass samples, complete drying sufficient to obtain a high-quality IR spectrum can evidently be achieved faster, in a time of about 1 to 2 h [27, 58], even at room temperature [45].)

Fig. 1.
figure 1

FTIR spectra of biomass samples of the bacterium Azospirillum brasilense Sp7 dried at 45ºC for (a) 1.5 h and (b) 23 h (three parallel measurements in each case with superposition of the three spectra normalised by the intensity of the amide I band of cellular proteins). The spectra were measured in the transmission mode in the form of thin films on ZnSe discs (the Figure was prepared by the authors using the data presented in [60]).

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Drying for 23 h (see Fig. 1b) led to a satisfactory agreement between the spectra in parallel measurements. Thus, this drying mode is quite suitable for preparing not too small amounts of cell biomass or biofilms for comparative IR spectroscopic analyses, especially when additional chemical analyses of their composition are required [59, 60].

Note also that Fig. 1b presents a typical example of a spectrum of samples of dry bacterial cell biomass, in which, against the background of a wide range of stretching vibrations of OH groups ν(O–H) in polysaccharides, bands of stretching vibrations of amides ν(N–H) and a characteristic region of different stretching vibrations ν(C–H) of aliphatic groups are observed (primarily symmetric (νs) and antisymmetric (νas) vibrations of –CH3 and –CH2– groups, representing a series of less intensive bands within the region of ~3000–2800 cm–1) [59, 60]. The most informative are: the amide I and amide II bands typical of all proteins [3, 20, 23, 35, 36, 43, 61–63]; a highly characteristic band of stretching vibrations of the carbonyl group ν(C=O), in this case (about 1740 cm–1) of low intensity, corresponding mainly to the ester group (cellular lipids, as well as reserve substances of the polyester class synthesised and accumulated under certain conditions—polyhydroxyalkanoates, PHA) [3, 30, 42–44, 54, 57–60, 64–69]; the region of different vibrations of polysaccharides and polysaccharide-containing biomacromolecular complexes (~1200–950 cm–1) [3, 25, 30, 59, 60].

The use of a thin film of bacterial biomass in measuring FTIR spectra in the ATR mode during spontaneous drying makes it possible to follow not only the change in the spectra during the removal of water (Fig. 2a), but also to reveal differences in the behaviour of the intracellular reserve biopolymer, poly-3-hydroxybutyrate (PHB) (Figs. 2b–2c). Figure 2 shows FTIR-ATR spectra for the bacterium Cupriavidus necator, which is an important PHB producer (accumulating PHB up to 90% of dry biomass [69]), in the process of spontaneous water removal (according to the data reported in [58]). From the data in Fig. 2b (the expanded part of Fig. 2a with spectra measured in the time range from 60 to 150 min) it can be seen that after 60 min of spontaneous drying up to 2.5 h, the changes observed in the most informative part of the spectrum (1800–1000 cm–1) are insignificant; in particular, the absorption bands have virtually the same positions. Nevertheless, a similar spectrum for the same biomass preliminarily subjected to a thermal stress (80°C for 90 min in a phosphate buffer, pH 7.4) shows that the PHB bands undergo changes during the removal of water (Fig. 2c). The indicated shift of the ν(C=O) band (from 1734 to 1719 cm–1) and the corresponding changes in the range of C–C–O vibrations of the PHB ester group (1300–1200 cm–1) provide evidence for spontaneous crystallisation of the biopolymer [58]. Basing on the observation of morphological changes in C. necator cells during the indicated hydrothermal treatment followed by drying, the authors of [58] related these changes in the degree of crystallinity to the coagulation of biopolymer granules and the removal of traces of intragranular water which plays the role of a “plasticiser”. (We also note that a comparison of Figs. 2b and 2c shows a distinct change in the shape of the bands of cellular proteins, especially of the amide I band at about 1650 cm–1, evidently related to protein denaturation upon thermal treatment (see Fig. 2c).)

Fig. 2.
figure 2

FTIR spectra in the ATR mode (single-reflection diamond ATR crystal) of a thin film of the biomass of the bacterium Cupriavidus necator H16. (a) Measurements in the course of drying after periods of time from 0 (the spectrum with the highest intensities of the bands marked with arrows), with intervals of 10 min (intermediate spectra), to 160 min (the spectrum with the lowest intensities of the bands marked with arrows); arrows mark the characteristic bands of water vibrations. (b) Measurements in the range of 1800–1000 cm–1 in the course of drying after 60 to 150 min (intermediate spectra with intervals of 10 min) of the same biomass (control). (c) Measurements in the range of 1800–1000 cm–1 in the course of drying after 60 to 150 min (intermediate spectra with intervals of 10 min) of biomass preliminarily subjected to a thermal stress (80°C for 90 min in phosphate buffer, pH 7.4); the main changes in the spectra associated with crystallisation of the reserve substance (PHB) are indicated by arrows. (The Figure was prepared by the authors using the data presented in [58].)

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In IR spectroscopic studies of small amounts of biomass, pressing of dry samples ground and mixed with KBr powder into special pellets is in some cases still used for measurements in the transmission mode [30, 39, 45]. Mixing with KBr is also used without pressing for the DRIFT methodology (see [66] and references cited therein). However, a special study on examples of various biological samples, including dried microbial biomass, has shown [66] that grinding a biomass sample with KBr (even without pressing) leads to shifts in the maxima of the main bands of several polar functional groups.

Figure 3a shows FTIR spectra obtained in the DRIFT mode for a powder of dry biomass of the bacterium Azospirillum baldaniorum Sp245 (grown under conditions corresponding to the intracellular accumulation of a reserve material, PHB [66], a homopolymer characteristic of azospirilla [42, 43, 54, 57]) after grinding without and with KBr. As can be seen from the spectroscopic data, the main bands of PHB, ν(C=O) (at about 1740 cm–1) and C–C–O vibrations (at about 1300 cm–1) of the ester fragment, after grinding in the presence of KBr shifted from 1746 to 1728 cm–1 and from 1300 to 1285 cm–1, respectively, i.e. by 18 and 15 cm–1. Such a significant shift of the maxima for bands which are known to be sensitive to the degree of PHB crystallinity [43, 58, 60, 67] provides evidence for the process of crystallisation of cellular PHB, which is initially present in cells in the form of amorphous granules [66, 68], induced by this mechanical action (in the presence of the polar KBr matrix).

Fig. 3.
figure 3

FTIR spectra of dried biomass samples of the bacterium Azospirillum baldaniorum Sp245. The measurements were carried out (a) in the diffuse reflectance mode (1) in the form of a powder without using KBr and (2) after grinding with KBr [66] and (b) in the transmission mode in the form of a film on the surface of a ZnSe disc (1) without grinding of the dry biomass and (2) after its grinding [60]. The bands for which the largest frequency shifts of the maxima were observed are highlighted with shading (the Figure was prepared by the authors using the data presented in [60, 66]).

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It is important to note that for the dry biomass of A. baldaniorum Sp245 grown under different conditions (with a lower accumulation of PHB), the grinding process in the absence of KBr resulted in a shift of the indicated bands similar in direction, although smaller in magnitude (8 cm–1 for the ν(C=O) band at about 1740 cm–1 [60]) (Fig. 3b). For the other bands in the spectra in Fig. 3, the positions of the maxima practically did not change within the instrumental error (±2…4 cm–1), although in Fig. 3a (in the case of a stronger influence of the polar KBr matrix), some changes in the shape of the amide I and amide II bands can also be noticed.

Thus, for the in situ analysis of native intracellular biopolymers (without their isolation) by FTIR spectroscopy, possible effects of such treatments during sample preparation [60] should be taken into account. At the same time, in the course of crystallisation of PHB (and other PHAs), the shape and width of the ν(C=O) band, which is most convenient for the in-situ quantitative analysis of this biopolymer (and other polyhydroxyalkanoates [3, 67]), can change significantly (see, for example, Fig. 2c). In this case, as shown in [68], a more accurate determination and comparison of the relative PHB content in cell biomass should be performed by taking into account not the intensity (height) of the ν(C=O) peak but its area.

2 SPECIFICITIES OF INTERPRETING FTIR SPECTROSCOPIC DATA IN MICROBIOLOGICAL ANALYSES

One of the most important properties of FTIR spectroscopy is its sensitivity not only to the structure and quantitative content of particular compounds (in biological samples, this corresponds to the presence of particular functional groups in biomacromolecules and supramolecular bioorganic complexes), but also to various intra- and intermolecular interactions [3, 19]. The latter are known to play a vitally important role in maintaining the native structure of biomacromolecules and their functional activity. This sensitivity of FTIR spectroscopy, on the one hand, can significantly complicate the interpretation of spectroscopic data, since under the influence of intra- and intermolecular interactions (primarily hydrogen bonds), the energies of molecular vibrations and, accordingly, the frequencies of the bands of particular functional groups can change; additional bands may also appear in the spectrum. On the other hand, this is a significant advantage of the technique which allows these interactions to be detected and studied nondestructively in situ and in vivo [3, 19, 30]. The above-mentioned physicochemical regularities of FTIR spectroscopy, as well as possible overlap of the frequency ranges characteristic of various functional groups, are the main difficulty that requires significant experience and qualification of the researcher. This explains the fact that the FTIR spectroscopy technique, which has long been used in the field of materials science as a routine tool, has not yet become routine in the life sciences, including microbiology, despite the constantly growing number of reports for which it was used.

In addition to the analysis of characteristic intensive bands, primarily those that do not overlap with other vibrational bands (such as the above-discussed ν(C=O) band at about 1740 cm–1, which is characteristic of polyesters of the PHA class [3, 43, 57–60, 66–68]), weaker bands in the IR spectra, e.g., in the range of different ν(C–H) vibrations of aliphatic groups (~3000–2800 cm–1), can also provide valuable information, especially in combination with the data of other techniques or results of chemical analysis. As an example, let us consider FTIR spectra of samples of two fractions of the biofilm matrix (BM) formed by the bacterium Azospirillum baldaniorum Sp245, that were obtained by its chromatographic separation (Fig. 4) and are characterised by different distributions of the molecular masses of their macrocomponents (80–40 kDa for BM1 and 35–20 kDa for BM2; Table 2) [59].

Fig. 4.
figure 4

FTIR spectra of two fractions of the matrix isolated from a biofilm formed by the bacterium Azospirillum baldaniorum Sp245 that were obtained by chromatographic separation of the matrix: (1) BM1 and (2) BM2 (see Table 2; according to the data from [59]). The characteristic bands of symmetric (νs) and antisymmetric (νas) stretching vibrations of –CH3 and –CH2– groups, ν(C=O) of the ester fragment, as well as the characteristic vibration range of polysaccharides (PS) are marked with arrows. The inset shows the structural formula of lipid A of the lipopolysaccharide typical of azospirilla [70] (with regions of the amide bond (C=O)–HN marked with an oval). (The Figure was prepared by the authors using the data presented in [59, 70].)

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Table 2.   Chemical composition (content of components in wt %) of lipopolysaccharide (LPS) isolated from cells, biofilm matrix (BM) and its fractions (BM1 and BM2) obtained by chromatographic separation of the matrix, as well as of the polysaccharide fraction from the biofilm matrix (BM3) obtained by its mild acidic hydrolysis, from the bacterium Azospirillum baldaniorum Sp245 (according to the data from [59])
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First of all, from the spectra in Fig. 4 it follows that the absorption in the polysaccharide vibration region (~1200–950 cm–1) for the BM1 fraction is much stronger than that for BM2. This is in good agreement with the chemical analysis data (see Table 2) showing that the total sugar content in BM1 is 4 times higher than in MB2. Since the carbohydrate part of these fractions is represented by lipopolysaccharide [59, 70], this is quite consistent with a more noticeable shoulder in the spectrum of the BM1 fraction in the range of ν(C=O) vibrations at about 1740 cm–1.

It has to be specially noted that in polyesters of the PHA class (and, in particular, in PHB), each monomer unit contains a carbonyl group, and when a PHA accumulates in noticeable amounts, the ν(C=O) band in IR spectra becomes noticeable even in microbial biomass [3, 57, 66–68]. In contrast, the molar fraction of the ester fragment [–(C=O)–O–] in the composition of lipids is relatively small (see, e.g., the inset in Fig. 4); in addition, the content of lipids in a bacterial cell is usually around 10% (see Table 1). As a result, in the absence of PHA accumulation, the ν(C=O) band of lipid components at about 1740 cm–1 appears in IR spectra of cell biomass as a very weak (often hardly noticeable) shoulder at the left wing of the amide I band of cellular proteins [3, 68].

From Fig. 4 it can also be seen that the region of generally weaker ν(C–H) vibrations of aliphatic groups (3000–2800 cm–1) for the BM1 fraction is noticeably more intensive as well (a contribution to this region is made by aliphatic groups not only of lipids but also of side chains of amino acid residues in proteins and of other biomolecules). Moreover, a comparison of this region for the BM1 and BM2 fractions shows that for BM1, the relative intensities of both bands of the stretching vibrations of methylene groups, νas(CH2) and νs(CH2), compared to the corresponding vibrations of terminal methyl groups νas(CH3) and νs(CH3), are significantly higher than those for BM2. This is in full agreement with the increased content of sugars (presented as components in LPS) in BM1 as compared to that in MB2 (see Table 2), since the lipid part of LPS contains long aliphatic chains of fatty acid residues (see the inset in Fig. 4).

Thus, a thorough analysis of different regions of an IR spectrum (even for such complex and inhomogeneous objects as microbial cells, biofilms and their macroconstituents), the measurement of which is much less laborious than chemical analyses, can provide valuable information on the relative macromolecular composition of the samples.

In analysing the composition of a biofilm matrix, another important point should be emphasised. For the polysaccharide fraction of the matrix obtained by its mild acidic hydrolysis [59], the IR spectrum of this fraction (BM3; see also Table 2) contains characteristic bands in the region of amide I and amide II at 1638 and 1545 cm–1, respectively (Fig. 5). However, according to both the data of chemical analysis (Table 2) and the results of polyacrylamide gel electrophoresis (SDS-PAGE), there is no protein in this fraction, and the polysaccharide is present in the composition of LPS [59]. (In the left wing of the broad band related to amide I in Fig. 5, above 1700 cm–1, a contribution from the ν(C=O) vibrations of the [–(C=O)–O–] ester fragments can be noticed corresponding to LPS; see the inset in Fig. 4). In this case, the appearance of amide I and amide II bands in the absence of proteinaceous macrocomponents is explained by the presence of amide bonds in lipid A in the composition of LPS (see the inset in Fig. 4; amide bonds [–NH–(C=O)–] are marked with an oval). ). This fact must be taken into account when analysing spectroscopic data of complex (micro)biological samples.

Fig. 5.
figure 5

A FTIR spectrum of the polysaccharide fraction from the biofilm matrix of the bacterium Azospirillum baldaniorum Sp245 obtained by mild acidic hydrolysis of the matrix (BM3; see Table 2; according to the data from [59]). The amide I and amide II bands, as well as the characteristic region of polysaccharide (PS) vibrations are marked with arrows. (The Figure was prepared by the authors using the data presented in [59]).

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Note also that the amide I band in Fig. 5 has a maximum at 1638 cm–1. This may have contributed to the splitting of the amide I band in the spectrum of the BM1 fraction (see Fig. 4, spectrum 1) which, in addition to the protein (with the corresponding commonly observed amide I band showing a maximum at 1654 cm–1 characteristic of α-helices [3, 20, 23, 35, 36]), also contains a significant proportion of LPS (see Table 2 for the content of sugars in LPS) resulting in the appearance of its corresponding “amide-type” peak with a maximum at 1635 cm–1.

Finally it is worth mentioning that, along with a growing number of publications in which FTIR spectroscopy in different variants has been successfully used to study microorganisms and other biological objects, in some reported results various inaccuracies can repeatedly be observed. They are both of a methodological nature (which can usually be easily distinguished by the spectral shape which is not typical for the studied objects and (or) by unusual relative intensities of typical peaks [71], the presence of a strong CO2 doublet at about 2350 cm–1 (see [72]), e.g., in [73], indicating an insufficient purging of the spectrometer internal volume for removing CO2 and possible accompanying water vapours [27], etc.) and arising in the interpretation of spectroscopic data. For example, the authors of [74] assigned to “isocyanate” a similar weak CO2 doublet at 2352 cm–1 in the IR spectrum of a biofilm formed by the bacterium Acinetobacter baumannii.

In the IR spectra of samples of PHB and its copolymer with poly-3-hydroxyvalerate isolated by the authors of [75], along with typical PHA bands (in par-ticular, the most intensive ν(C=O) band at 1729 cm–1), there are also strong amide I and amide II bands at ca. 1648 and 1547 cm–1, as well as a broad intensive band in the ν(O–H) region (~3500–3000 cm–1). These data indicate an evidently insufficient purification of the biopolymers from cell biomass and (or) its macrocomponents. A similar mistake was made by the authors of [76] where in the IR spectrum of the PHB sample isolated (as indicated in the caption to the Figure) from biomass of the bacterium Bacillus cereus, there are strong bands characteristic of proteins (including amide I and amide II at ca. 1634 and 1533 cm–1). At the same time, the ν(C=O) band in the region of ~1740 cm–1 is practically absent, which makes the presence of the biopolymer, “isolated” by the authors of [76], in the sample rather doubtful.

The authors of a remarkably informative comparative study of different modes of measuring FTIR spectra of samples of the bacterium Aquabacterium commune [45], when interpreting their data, described an asymmetric ν(C=O) absorption band in the region of 1739–1725 cm–1 with a significant intensity comparable with those of the bands of proteins (amide I and amide II) as corresponding to membrane lipids and fatty acids. Such an interpretation, in our opinion, does not correspond to the physiological composition of cells (in particular, to the content of lipids), since the ν(C=O) band of such an intensity in bacterial cells is observed only when significant amounts of reserve polyesters (PHA) are accumulated [3, 66–68]. In the article with the first description of this bacterium [77], it was mentioned that it contained PHA as reserve biopolymers, and the micrographs of this bacterium and of the other described Aquabacterium species published in [77] show rather large multiple PHA granules. (Note that the authors of [45] cultivated A. commune on the same medium and under conditions similar to those indicated in [77], so the general composition and morphology of the cell biomass obtained in [45] is expected to have been similar.) Comparing the IR spectra of dry biomass (biofilm and planktonic culture) of the bacterium A. commune [45] and a pure PHB sample isolated from cells of the bacterium Azospirillum baldaniorum (previously known as A. brasilense) [66] (Figs. 6a–6b) shows that all the main bands characteristic of PHB (and of other PHAs [65, 67]) are present in the spectra of bacterial biomasses of A. commune. In addition, large PHA granules are visible in micrographs of bacterial cells [77] (Fig. 6c). It is also important to emphasise that the main changes in the shape of the bands in Fig. 6a (spectra 1 and 2 of A. commune biomass) correspond to two main vibration regions of PHB that are sensitive to the degree of its crystallinity (marked in Fig. 6a by larger dark arrows).

Fig. 6.
figure 6

FTIR spectra of (a) dry biomass of the bacterium Aquabacterium commune (1) in the form of a biofilm formed on the surface of stainless steel after 100 h, and (2) in the form of a planktonic culture isolated from the same cultivation medium (measurements were carried out by reflectance micro-FTIR spectroscopy [45]); (b) a thin film of poly-3-hydroxybutyrate (PHB) isolated from biomass of the bacterium Azospirillum baldaniorum Sp245 [66] (measurements were carried out in transmission mode), as well as (c) electron micrographs of Aquabacterium commune cells (scale bar 1 μm) [77]. In Fig. (a), arrows show the positions of bands corresponding to those of PHB in Fig. (b); two larger dark arrows indicate the regions of the main PHB bands corresponding to the vibrations of the C=O (about 1740 cm–1) and C–O–C (about 1290 cm–1) bonds of the ester fragment which are sensitive to the degree of PHB crystallinity [66–68]. In Fig. (c), cell images show polyphosphate inclusions (dark) and large PHA granules (white) [77]. (The Figure was prepared by the authors using the data presented in [45, 66, 77]).

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Regretfully, in some publications there were also more significant errors. Thus, in [78, 79] the FTIR spectra of biogenic selenium nanoparticles of microbial origin, that had been measured in the absorption mode (with peak maxima directed upwards), were presented in the transmission scale on the ordinate axis (hence considering minima in the spectra instead of peaks) and, as a consequence, with an incorrect interpretation. However, it should be noted that relevant corrections have later been published in both cases (see [80, 81], respectively).

From the information mentioned above concerning the common mistakes found in the literature, that had been made during measurements and interpretation of the data obtained, the following main conclusions can be drawn. Since the FTIR spectroscopy technique, especially with modern instruments and accessories, often does not require any complex sample preparation even for microbiological specimens, this apparent “simplicity” may distract from the need for strict adherence to the main stages of preparing the equipment and sample treatment noted in the relevant manuals [3, 27, 30]. The technique itself, however, can be somewhat ‘misleading’ in the sense that, while performing incorrect measurements and (or) improper preparation of samples, an IR spectrum can always be obtained, but it may not reflect the real structure and composition of the system under study. Nevertheless, these methodological details are quite easy to take into account. The most important part of the study after obtaining adequate experimental IR spectra is definitely their interpretation, which requires considerable skill, experience and attentiveness. The complexity and high informativity of IR spectra of biological samples have already been noted above, being associated both with possible overlapping of the regions (energy values) related to vibrations of various functional groups of biomolecules and with their sensitivity to intermolecular and intramolecular interactions. Thus, when interpreting the data, it is important to take into account the potential ambiguity of the information contained in the spectra. This also necessarily includes taking into account the presence of different vibration types (stretching, various types of deformational or bending, symmetric and asymmetric vibrations, etc.), which is especially important for polyatomic functional groups. These “fingerprints” should be specially noted in the analysed spectra, considering their relative intensity, which can vary within a significant range, in particular, depending on intermolecular interactions, the state of the sample, the conditions of its preparation, etc.

3 CONCLUSIONS

In this mini-review, using examples selected by the authors from the literature mainly over the last 10–15 years, the importance of the main stages in sample preparation of microbiological objects, and primarily of cell biomass and biofilms, is considered. Possibilities of changing the native state of the system under study during sample preparation (drying, grinding, etc.) is specially emphasised. Thus, while FTIR spectroscopy is non-destructive (does not change the composition of the sample) and largely non-invasive (does not affect its chemical and morphological structure), it is important to observe this rule also when collecting and preparing samples. Some important features of the interpretation of spectrochemical data and possibilities of comparative quantitative estimates of the contents of macrocomponents are considered when analysing the relative intensities of characteristic vibrations. Examples are presented of some typical methodological and analytical inaccuracies and errors observed in the literature on FTIR spectroscopic studies of microbiological objects; their causes are analysed; recommendations are given to ensure the correct preparation and implementation of an IR spectroscopic bioanalysis.