Fast and reliable inactivation of bacterial endospores can be achieved with a great variety of different antiseptics and disinfectants. Most of the sporicidal agents provoke lysis and leakage of intracellular constituents by degrading most of the permeability barriers, which play different roles in the endospore's resistance. The various layers of proteinaceous spore coats, which surround the spore cortex, protect the spore from attacks by a large number of chemicals, particularly oxidizing agents such as hydrogen peroxide, sodium hypochlorite, chlorine dioxide, or ozone (52
). Smaller hydrophobic and hydrophilic molecules are hindered on their way to the core by the extremely low permeability of the spore's inner membrane (36
Most of the oxidizing agents, strong acids, and ethanol kill spores by causing some type of damage to the spore's permeability barriers, such that when the treated spores germinate, these damaged membranes rupture, resulting in spore death (36
). For some of the oxidizing agents, the location of damage could be pinpointed to the endospore's inner membrane (5
); for others, only a general disruption of spore permeability barriers is reported. Interestingly, Cortezzo et al. found out that spore inactivation via a variety of oxidizing agents is not accompanied by loss of DPA (5
), whereas acid-inactivated spores extrude DPA and other fibrillar material, including DNA (51
). In addition, pretreatment with oxidizing agents makes the surviving spores more sensitive to inactivation by normally nonlethal heat and osmotic stresses in the presence of high salt concentrations in plating media.
These findings are in good agreement with the results we obtained by using PAA as a disinfectant. Among other peroxides, peracetic acid is frequently used in common disinfection procedures and is considered to be a more potent biocide than hydrogen peroxide. As with hydrogen peroxide, the hydroxyl radical appears to be of prime importance, and its inactivation mechanism is based not on DNA damage but rather on a weakening of the endospore's inner membrane by disrupting sulfhydryl (—SH) and sulfur (S—S) bonds of proteins (32
). As shown in Fig. and in the dendrogram in Fig. , this internal damage only partially forces the spores to extrude core material, since most of the endospores remained intact and retained intracellular DPA after a short treatment (15 min) with highly diluted PAA (1%). The leakage is more pronounced at higher PAA concentrations (2%) and with longer exposure times (>1 h). Furthermore, the probability of measuring not only intact or harmed endospores but also metabolites of the vegetative cells increased with the inactivation time and PAA concentration, as shown in Fig. . According to these results, not a microbial cell but a residue of formerly intracellular granules of PHB has been measured. When PHB-producing cells die, PHB is released into the environment, where it is transformed into a denatured semicrystalline state (21
In case of the chlorine-releasing product Danchlorix, the complete spore lysis we observed was also achieved in other studies with sodium hypochlorite (28
). In those studies, loss of refractivity, separation of the spore coats from the cortex, extensive discharge of Ca2+
, dipicolinic acid, and DNA/RNA, and finally lysis occurred.
Inactivation of endospores by wet heat had the same depleting effect. Here again, not the DNA but proteins are assumed to be the target (36
), explaining the shift of the amide III band in the observed Raman spectra of Fig. . This is confirmed by previous work (38
), where alterations of amide I and amide II bands due to autoclaving could be monitored by means of Fourier transform infrared spectroscopy (FT-IR), and where the absorption band at 1,570 cm−1
, a diagnostic band for DPA, was lost, as shown in Fig. . Since the major part of this salt is deposited in the core of endospores, these ruptures due to autoclaving-induced cleavage of endospores seem to reach into the inner part of the cell, where most spore enzymes, as well as DNA, ribosomes, and tRNA, are located. Thus, not only denaturation of macromolecules and subcellular structures, including proteins, cytoplasmic membranes, and nucleic acids, occurs during autoclaving, but also a major loss of those biomolecules, as well as the ubiquitous DPA salt, takes place.
This method of extracting soluble microbial proteins can be beneficial for some analysis methods that rely on those proteins as biomarkers, e.g., for the matrix-assisted laser desorption-time-of-flight (MALDI-TOF)/intact-cell mass spectrometry (ICMS) methodology (29
). But unlike these approaches, Raman spectroscopy does not analyze bulk samples or fractions thereof but intact single cells.
That is why all the types of chemical agents discussed above are inappropriate for the purpose of identifying bacterial spores by means of Raman spectroscopy, since retaining the structural integrity and most of the biochemical composition of single cells is a necessity for micro-Raman-based identification of inactivated pathogenic endospores. The considerable alterations of spore integrity by PAA, Danchlorix, and autoclaving obviously have a nonnegligible impact on the Raman spectra of the endospores, which coincide with a loss of information in the Raman spectra and thus with decreased identification accuracy. Additionally, the homogeneity of the inactivation treatment has to be guaranteed. The achievement of a maximum of uniformity among the sterilized cells is another important objective, since chemotaxonomic classification relies mainly on constantly recurring spectral patterns among an ensemble of single-cell spectra. If the inactivation treatment alters cells of the same species differently and thus decreases the uniformity of the respective spectra, these Raman data are less useful for building up a database for supervised identification routines. Alternatively, the database might comprise all the inactivation-induced spectral variances, but then it would probably gain a dimension of enormous extent.
Formaldehyde is also employed for a variety of decontamination processes, since it is sporicidal (49
). Loshon et al. achieved a 99% killing rate for B. subtilis
endospores in 40 min at 30°C with 25 g/liter formaldehyde (30
). Endospores are inactivated by formaldehyde due to some unique features of this molecule. This small molecule can pass through all the protective layers to advance directly into the core of the endospore. There the genotoxic properties of formaldehyde take effect, causing spore inactivation at least in part by DNA damage; protein-DNA cross-linking is proposed to be one mutagenic mechanism of formaldehyde (30
). However, the precise nature of the DNA damage is as yet unknown. The major resistance mechanisms of spores against formaldehyde are the saturation of spore DNA with α/β-SASPs and the recA
-encoded repair pathway, which can repair at least some of the formaldehyde-induced lesions in DNA (36
). The interaction of formaldehyde with proteins might also give rise to degradation of the spore integuments, but apparently not to such an extent that spore core material is released.
For the purposes of Raman spectroscopy, the use of formaldehyde to inactivate endospores is superior to any of the other treatments analyzed in this work. The inactivation experiments suggest the usage of this agent as a sporicide, since reliable, strain-independent, and fast inactivation of the Bacillus strains analyzed was achieved. It was shown by Raman spectroscopy that the formaldehyde inactivation technique is suitable for obtaining reproducible Raman spectra.
Possible reasons for competing intraspecies varieties in the Raman spectra are manifold, e.g., the preparation/inactivation process might induce spectral variances on the single-cell level, as can be seen in the case of PAA-inactivated endospores; cultivation parameters, such as growth time, temperature, and nutritional conditions, which have an impact on whole batches, are also factors to be reckoned with (14
). According to our results shown in Tables and , at least the formaldehyde-induced inhomogeneities can be neglected, since all four different Bacillus
strains were chemotaxonomically identified with the help of other, independently cultivated batches of the same strains grown under the same cultivation conditions. The interspecies distinctions are obviously still high enough to obtain satisfactory identification rates for different Bacillus
species, which serve as B. anthracis
models. This is noteworthy insofar as only a very limited model database was used, comprising just four different Bacillus
species. The comparison between the accuracies of the cross-validation and the hold-out techniques for the bacterial database shows that the main problem of our current evaluation method is overfitting. Accuracy decreases if a trained classifier is adopted to an independent data set. This can be avoided by building a database up out of more than one batch and more data. But even with a larger and more diverse database, preprocessing and, most importantly, dimension reduction is necessary.
For anthrax detection, evaluation of as many different B. anthracis strains as possible, as well as genetic near neighbors, is mandatory. Currently, we are elaborating a micro-Raman-based procedure for the identification of B. anthracis strains inactivated with 20% formaldehyde according to the procedure described here.
Taken together, the results of this study showed that detection of the anthrax agent, isolated from real-world samples, via micro-Raman spectroscopy can be a reliable alternative for fast point-of-care testing. On-site diagnosis is ensured by inactivating the samples with the formaldehyde treatment described in this study, which is compatible with the micro-Raman identification approach. Further investigations should aim at analyzing possible interference due to, e.g., the influence of the native surrounding matrices or the isolation procedure. Additionally, efforts should be focused on the question of whether the inactivation efficacy of formaldehyde is reduced due to matrix effects.