The use of enzymes for the asymmetric reduction of activated C=C double bonds can be a viable and straightforward alternative to asymmetric hydrogenation. Traditionally, whole cell microorganisms were used for this purpose but a recent increase in the number of isolated and characterised ENEs means that recombinantly-expressed enzyme preparations are now generally favoured over whole cells, as a number of recent publications demonstrate (1–10).

Double bond ‘activation’ to facilitate ENEs mediated reduction can be achieved in many cases by alpha substituted functional groups including aldehydes, ketones or nitro moieties. Carboxylate derivatives (such as esters, lactones and anhydrides) can also act as activating groups but their ability to sufficiently activate the C=C bond in the absence of other groups is less evident (11, 12). The traditional approach in these cases is to turn to chemocatalytic hydrogenation (see (13–15) for reviews focused on industrial applications). Herein we describe a new approach to activate α,β-unsaturated carboxylic acids for the reduction with ENEs using a substrate engineering approach.

2.1 General

2.2 Enzyme Preparations

2.3 2,2,2-Trifluoroethyl Cinnamate (3a) and 3-Phenyl-Acrylic Acid 2,2,2-Trifluoro-1-Trifluoromethyl-Ethyl Ester (5a)

2.4 3-Phenyl-Acrylic Acid 2,2,3,3,4,4,4-Heptafluoro-Butyl Ester (6a) and (Perfluorophenyl)Methyl Cinnamate (7a)

2.5 1-Cinnamoylpyrrolidin-2-one (9a)

2.6 3-Cinnamoyloxazolidin-2-one (8a)

2.7 (E)-1-(2-Methyl-3-Phenylacryloyl)Pyrrolidin-2-one (10a) and (E)-1-(2,3-Diphenylacryloyl)Pyrrolidin-2-one (11a)

2.8 Small Scale Screening Reactions

2.9 Preparative Scale Screening Reactions

2.10 Analytical Methods

It has been found that a particular ENE in Johnson Matthey’s collection, a homologue from the tobacco ENE reductase fold (16), ENE-105, was capable of reducing methyl ester 2a (Figure 1), albeit in a very low yield of 3% (Entry 2, Table I). By comparison, cinnamic acid 1a was a poor substrate and showed no conversion to the reduced product 1b at pH 7.0 (Entry 1, Table I). The pKa of cinnamic acid 1a is 4.4 and therefore, at pH 7.0, the carboxylic acid should be deprotonated affecting its ability to bind to the enzyme active site. This observation is in line with other literature examples where carboxylates were found to be poor activating groups (17). Encouraged by this initial result, we turned our efforts towards the use of more activated esters. It was envisaged that converting the alkyl chain in the ester moiety to a more EWG could lead to an increase in double bond activation. A similar approach has been reported previously by BASF SE for the lipase-catalysed kinetic resolution of racemic amines and alcohols, where the choice of acylating agent proved critical (18). We chose trifluoroethyl ester 3a as a starting point which was reduced by ENE-105 and *ENE-69 in 6% and 12% conversion respectively (Entry 3, Table I) suggesting that the addition of an EWG had a positive activating-effect on the reduction. To consolidate this theory, ethyl ester 4a was tested with the novel ENEs; only a trace of reduction was observed <0.5% (Entry 4, Table I).

Fig. 1

Other EWGs such as hexafluoroethyl in compound 5a, heptafluorobutyl in 6a and pentafluorobenzyl in 7a could also activate the double bond in the same way, so 5a, 6a and 7a were prepared by reacting cinnamoyl chloride with the corresponding fluorinated alcohols and these substrates were subsequently tested with the ENEs. Hexafluoro 5a was not reduced by ENE-105 or *ENE-69 (Entry 5, Table I), instead, a significant amount of hydrolysis product (cinnamic acid 1a, 10%) was observed. Heptafluorobutyl 6a and pentafluoro 7a were poor activating groups with 6a showing only a trace amount of product 6b (Entry 6, Table I) and 7a giving no conversion (Entry 7, Table I).

With only limited success with the fluorinated activating groups, our efforts turned towards cyclic imides since activated substrates 8a and 9a have been shown to be highly activated towards Michael addition reactions (19, 20, 21) (Figure 2). Compounds 8a and 9a were synthesised and tested with enzymes ENE-105 and *ENE-69. Pleasingly, oxazolidinone 8a was successfully reduced by both ENEs (51% and 39% conversion to 8b, Entry 1, Table II) and pyrrolidinone 9a was reduced to 9b in >95% conversion (Entry 2, Table II), proving to be an excellent activating group. The ¹H NMR shift of the alkene proton alpha to the carbonyl for pyrrolidinone 9a is shifted down field (7.92 ppm) compared to cinnamic acid 1a (6.46 ppm), therefore supporting the electron-withdrawing nature of the activating group.

Fig. 2

The enzymes were then tested for their ability to reduce α-substituted cinnamic acid derivatives such as α-methyl 10a and α-phenyl 11a (Figure 3). Encouragingly, the tri-substituted double bond in 10a was reduced to 10b in >95% conversion by ¹H NMR analysis (Entry 2, Table III). However, bulkier substrate 11a, was not tolerated so well on an analytical scale due to solubility issues causing mass-transfer limitations (Entry 3, Table III). The reaction was repeated on a larger scale with stirring (Entry 4, Table III) and >95% conversion was achieved. 10b and 11b were obtained as racemic mixtures.

Fig. 3

With a successful activating group found, the reaction was repeated on a preparative scale to test reproducibility and scalability (Table IV). Pyrrolidinone 9a was successfully reduced using enzyme ENE-105 at 130 mg scale with the desired product 9b being obtained in 95% conversion by ¹H NMR (Entry 1, Table IV). 72% conversion to 10b was achieved after 20 h (Entry 3, Table IV) on the reduction of pyrrolidinone 10a at 500 mg scale.

Having found enzymes in Johnson Matthey’s collection that could successfully reduce masked carboxylic acids, other commercially available enzymes were tested as a comparison on the reduction of 10a (Table V). Six enzymes from Johnson Matthey collection (Entries 3 to 6, Table V) and seven enzymes purchased from Codexis (Entries 7 to 13, Table V) were compared with ENE-105 and ENE-69* (Entries 1 and 2, Table V). It was found that, despite the extra activation of the C=C double bond, none of the tested enzymes could reduce cinnamic acid derivative 10a, highlighting the unique ability of ENE-105 and *ENE-69 within the focused library (13 enzymes) screened.

In summary, we have shown that cinnamic acid derivatives activated as fluorinated esters or as cyclic imides can be reduced using Johnson Matthey enzymes ENE-105 or *ENE-69. The concept of ‘substrate engineering’ as opposed to ‘enzyme engineering’, offers a complimentary and faster approach to developing a bioprocess, making difficult transformations possible. The reduced products can be subsequently converted to the parent carboxylic acids by LiOH hydrolysis (22, 23) and the potential re-use of these activating groups will be investigated in the future. It is envisaged that the work will lead to further examples of activated acids or esters being reduced by ENEs.

The biocatalysed reduction of the double bond of cinnamic acid derivatives is strongly influenced by the nature of the EWG. While no conversion was observed on the biocatalysed reduction of cinnamic acid 1a, an enzyme in Johnson Matthey’s collection, ENE-105, was capable of reducing methyl ester derivative 2a in low conversion. By replacing the alkyl chain in the ester moiety by a more EWG, such as fluorinated alkanes, and in the presence of enzymes ENE-105 and *ENE-69, we were able to significantly increase conversion to the reduced product. Furthermore, other electronegative derivatives such as cyclic imides proved to be even better activating groups, allowing the reduction of challenging substituted double bonds such as substrates 10a and 11a.

In summary, by ‘masking’ the carboxylic acid moiety into a fluorinated alkyl ester or a cyclic imide, following a straightforward synthetic procedure, and in combination with the right enzyme, it was possible to biocatalytically reduce the conjugated double bond of cinnamic acid and substituted derivatives.

This book is a fascinating account of how nanoparticles and nanotechnology are increasingly employed in a diverse array of applications ranging from plant growth to food packaging, biosensing, enzyme immobilisation and more. The book is divided into 20 chapters, each dealing with a specific application of nanotechnology and written by a different group of eminent academics from Indian universities and research institutes.

Each chapter is a review of its own topic area and the literature citations at the end of each chapter make it easy for the reader to use this book as a reference volume from which further in-depth reading can be pursued by following the cited literature.

Chapter 1 deals with the phytotoxicity of nanoparticles in plants which can lead to both positive and negative outcomes. For example, improvement in germination rate and growth have been reported in seeds of rice exposed to carbon nanotubes; on the other hand, toxicity has also been widely reported, including studies on aluminium oxide and zinc oxide nanoparticles hindering root growth rate.

Chapter 2 considers an entirely separate but equally important area of use of nanomaterials in food packaging. It concludes that “it is essential to perform safety assessment of nanomaterials before their application in food packaging or processing” and provide a citation on how to do this using a “decision tree”.

Chapter 7 introduces how biosensors derived from nanobiotechnology can be used to monitor the environment and gain information relating to its health and the detrimental effects that modernisation and industrialisation have had on the planet. Biosensors need to be specific, rapid, sensitive and cost-effective. The advent of nanotechnology and biosensors has made this possible and the authors of this chapter (Gupta and Kakkar) explore the different types of biosensors that have been developed over recent years. The authors give a brief explanation of how different types of sensors work using a combination of bio-recognition components and different transduction principles. Types include: (a) immunosensors; (b) enzymatic biosensors; (c) whole-cell based sensors; (d) biosensors; (e) genosensors; (f) aptasensors and (g) biomimetic biosensors. The role of the transducer is to convert the biochemical response into an analysable and measurable signal. The outputs can be electrochemical, optical, piezoelectric, thermometric or magnetic.

Chapter 10 tackles the interesting area of enzyme immobilisation and the use of chitosan nanoparticles therein. The biopolymer’s distinct physicochemical properties have been described to offer an excellent microenvironment for enzyme immobilisation through adsorption, covalent binding or cross-linking, to achieve desirable enzymatic activity and stability. On the other hand, nanoparticles as materials of enhanced properties, owing to their high surface to volume ratio, have been introduced as attractive candidates for enzyme immobilisation. The chapter briefly discussed various methods for the preparation of chitosan nanoparticles for enzyme immobilisation including reverse micelle, coprecipitation, ionotropic gelation and ionic or emulsion cross-linking methods. Different methods for enzyme immobilisation such as support binding, cross-linking and entrapment, as well as different materials used as supports have been explained too. This section is then closed by presenting some examples for immobilisation of different enzyme families (for example, α-amylase, β-galactosidase, cellulase, laccase, lipase or protease) through applying chitosan nanoparticles.

Chapter 13 offers an overview of solid lipid nanoparticles (SLN) as pharmaceuticals delivery systems whereas Chapter 19 gives a review of the most commonly used nanocarriers for drug delivery systems, with a focus on vesicular, polymeric and inorganic carriers.

SLN are lipid-based formulations, containing typically non-toxic biodegradable polymers forming a solid hydrophobic core suspended in an aqueous phase, the whole structure being stabilised by surfactants. The therapeutic agent is dissolved or dispersed in the solid lipid core, the SLN being suitable for incorporation and delivery of both hydrophilic and hydrophobic drugs. SLN present significant advantages over conventional drug delivery systems, including but not limited to biocompatibility and bioavailability, reduced drug leakage and increased physical stability of the drugs. In addition, they have been used successfully in various drug delivery techniques.

Novel applications of SLN as drug carriers are described in the field of gene therapy, peptide drug delivery and vaccines. SLN production methods use low mechanical force, allowing successful incorporation and delivery of nucleic acids in gene therapy. Overall, SLN are promising alternatives to traditional drug delivery systems, offering multiple advantages in terms of drug delivery and bioavailability, as well as being economically efficient and easy to produce on scale.

An extensive summary of FDA-approved nanomedicines is included in Table 19.1 (Chapter 19), which also summarises the advantages of these specific formulations. The main types of nanocarriers described in Chapter 19 are vesicular carriers (liposomes and niosomes), polymeric nanoparticles and inorganic carriers (silica, gold and calcium nanoparticles). Liposomes and solid lipid nanoparticles (see also Chapter 13) are suitable for the delivery of drugs by any route, either oral or parenteral and can be used with both hydrophilic and lipophilic drugs. Their main advantages reside in protecting labile drugs, limited toxicity and a sustainable targeted release of the drug.

Inorganic nanocarriers exhibit higher stability and resistance to microbial growth, while having a low toxicity and allowing facile surface modifications. Mesoporous silica nanoparticles allow encapsulation of the therapeutic agent and targeted delivery to tumour cells in cancer therapy. Gold nanoparticles are biocompatible and bio-inert and have been successfully used in covalent conjugation with protein antigens in developing vaccines for cancer immunotherapy. Calcium phosphate nanoparticles are excellent candidates for developing ceramic-based carriers for peptide drugs prone to degradation, such as insulin.

In conclusion, I consider this book to be a positive contribution to the biotechnology literature, although I do not recommend reading this book sequentially from Page 1 as the variety of topics introduced is too great and each individual topic is not explored in depth. It is best used (and deserves recommendation) as a reference source from which each chapter can be used as the starting point to a more in-depth study or review of a particular topic. There are some negative aspects of the presentation of this work which do, unfortunately, detract from its enjoyment. These are exemplified in the poor quality of the diagrams, the grammatical errors and the somewhat odd references of Chapter 7.

Overall, though, this book is a positive addition to the biotechnology reference bookshelf.

In the era of Industry 4.0, with global climate change, increasing population and developing technology, the spread of heavy metal pollutants in aquatic areas is increasing. Bacterial resistance and metal accumulation capability are common phenomena that can be exploited for the bioremediation of the environment, hence these resistant bacteria may be potential candidates for biotechnological applications. Despite the risks caused by antibiotic-resistant bacteria, heavy metal-resistant bacteria can be used in detoxification processes to convert a toxic form to a harmless form of a substance by developing biotransformation mechanisms. Bioremediation studies have been carried out to identify candidate species (1–4).

In recent years, the increase in pollution by toxic compounds and heavy metals in marine areas makes it increasingly important to study the relationships between bacteria and toxic compounds. Studies related to the transformation of compounds into different forms via bacterial metabolic processes for the removal of toxic substances from the environment have gained importance. Detection of bacteria that are resistant to heavy metals in natural environments constitutes the first step to provide data for remediation studies.

Bacteria are some of the most important components in marine ecosystems. Since bacteria adapt to new conditions created by environmental variables around them, knowledge of bacteria provides data in terms of defining environmental factors, public health status and ecosystem function. Marine areas are exposed to domestic and industrial wastes depending on local technology levels and population. Many xenobiotic micro pollutants, antibiotic derivatives and metabolites reach the sea from human activity. This concerning issue is considered an important factor for global health with respect to the evolution and detection of antibiotic resistance in bacterial pathogens (5). Since the spread of antimicrobial resistance is not restricted by phylogenetic, geographic or ecological borders, studies describing regional status of bacterial resistance in natural areas are important.

Antibiotic resistance can spread rapidly among bacterial species (6). It is known that the occurrence of antibiotic-resistant bacteria in natural environments reduces the effectiveness of antibiotics in the treatment of infectious diseases. Due to the increasing global resistance of bacteria against antibiotics, humanity is constantly being forced to develop new antibiotic derivatives. This vicious circle is one of the most important problems of our age and poses a threat for the future. Thus, it is important to know the resistance levels of bacteria and to produce regional antibiotic resistance profiles in natural areas. Aquatic environments constitute a way to disseminate not only antibiotic-resistant bacteria but also the resistant genes in natural bacterial habitats (7). It has been well documented that the aquatic environment is a potential reservoir of antibiotic-resistant bacteria, furthermore the prevalence and persistence of antibiotic resistance in bacterial pathogens is a threat and a source of considerable concern to public health (8–15). It is known that environmental factors such as overpopulation, livestock farming, insufficient drainage and sanitation infrastructure may provide hotspots for environmental antibiotic-resistant bacteria transmission (16).

In aquatic environments, antibiotic-resistant bacteria can be accompanied by heavy metal-resistant bacteria that are often induced by the presence of metal caused by anthropogenic activities and environmental factors (16, 17). Heavy metals are introduced into the marine environment in different ways. Accumulation in sediment can affect aquatic life negatively for a long time. Bacteria that will take part in the transformation of heavy metal salts into harmless forms must be resistant to the heavy metals. Bacteria that cannot adapt to the changes metabolically will be eliminated and therefore various pollution inputs accumulated in the sediment will affect the composition of microbial diversity. Sediments containing harmful, inorganic or organic particles are relatively heterogeneous in terms of physical, chemical and biological properties and are an important source of heavy metal contamination (11). It has been reported that microplastics mediate the spread of metal- and antibiotic-resistant pathogens due to their ability to adsorb various pollutants (18, 19). Bacteria resistant to heavy metals in marine areas have developed various resistance mechanisms to counteract heavy metal stress. Only bacteria that can withstand the current heavy metal concentration can survive in these areas.

Heavy metals accumulate in biota via food chains and are transferred between organisms in marine environments. This cumulative process, named biomagnification, is higher in the sea than in terrestrial environments (15, 20) and this implies significant effects of heavy metal pollution in marine areas. On the other hand, heavy metal-resistant bacteria can play a role in detoxification by converting a toxic form into a harmless form through biotransformation mechanisms that develop in natural environments. These mechanisms include the formation and sequestration of heavy metals in complexes and the reduction of a metal to a less toxic species (21, 22).

Metal-resistant bacteria have developed very efficient and varying mechanisms for tolerating high levels of toxic metals and thus they carry an important potential for controlling heavy metal pollution (23). In many prokaryotes, it has been shown that the mechanism for resisting heavy metals develops over time. This process has been studied in species such as Escherichia coli and Staphylococcus aureus. It is reported that many different species of Pseudomonas, Bacillus, Enterobacter, Providencia and Chryseobacterium are efficient for reducing heavy metals (1–4).

It is known that the occurrence of bacteria resistant to antibiotics and heavy metal salts in the sea is related to the pollutants present in the environment. For the reasons highlighted above, it is important to determine the profile of antibiotic and heavy metal-resistant bacteria in marine environments. Marine areas which have different environmental inputs present novel media for bacterial studies.

For the present study, the Güllük Bay of the Aegean Sea, Turkey, was chosen since it is a dynamic area due to marine transportation, seasonal population growth depending on tourism, aquaculture, recreational and agricultural activities and terrestrial pollution inputs transported from rivers.

Probable faecal source analysis conducted in Güllük Bay showed that the primary source of the detected bacteriological pollution is anthropogenic (24). A significant part of domestic wastewater in the region collects in sealed septic tanks. It is possible for the wastewater to reach the sea by mixing the sedimentary septic tanks with groundwater. Chemical and biological studies (24–33) confirm that regional pollutants have reached Güllük Bay.

It is well known that sewage transported via domestic wastewater carries antibiotics to marine environments. This has an effect on metabolic capabilities of bacteria in marine environments. For example, β-lactam antibiotic derivatives used for human infection treatment may enter marine environments via domestic wastewater. Bacteria may obtain resistance via intercellular contact mostly using a conjugation mechanism (34). The existence of antibiotic-resistant bacteria is an indicator of domestic pollution. Furthermore, antibiotic-resistant bacteria may cause a vicious cycle. This problem has grown in recent years due to systematic use of antibiotics in animal husbandry and overuse of antibiotics (35, 36).

The frequencies of heavy metal-resistant bacteria and antibiotic-resistant bacteria were investigated in seawater and sediment samples collected from Güllük Bay in the period between May 2011 and February 2013.

2.1 Sampling Area

2.2 Sampling

2.3 Bacterial Isolation and Identification

2.4 Bacterial Resistance Against Antibiotics

2.5 Bacterial Resistance Against Heavy Metal Salts

3.1 Bacterial Resistance Against Antibiotics

3.2 Multiple Antibiotic Resistance Indexes

3.3 Bacterial Resistance Against Heavy Metals

Indicator bacteria levels reported in Güllük Bay and the presence of pathogenic bacteria (25, 26) support the relationship between the resistance data detected in the current study with bacteriological pollution levels. In the present study, bioindicator bacteria showing human-induced pollution input isolated from seawater had the highest frequency of resistance against nitrofurantoin (100%) and sulfonamide (95%). Sulfonamides were the first antibiotics developed for clinical use. Sulfonamides have been widely used to treat bacterial and protozoan infections in humans, domestic animals and fish since their introduction to clinical practice in 1935 (45–47). The results of higher resistance against sulfonamide in the present study were similar to the findings of sulfonamide resistance in another study (48). For example, there were significant increases in numbers of bacteria resistant to oxytetracycline, oxolinic acid and florfenicol in sediments from an aquaculture site compared with those from a non-aquaculture control site. Interestingly, in another study a similar number of antibiotic-resistant bacteria were isolated from aquaculture and non-aquaculture sites (49). Gram-negative bacteria (predominantly Plesiomonas shigelloides and Aeromonas hydrophila) were isolated from aquaculture ponds in the south-eastern USA and it was reported that antibiotic resistance to tetracycline, oxytetracycline, chloramphenicol, ampicillin and nitrofurantoin were higher in antibiotic-treated ponds compared to non-treated rivers (50). It was determined that bacteria isolated from Sopot Beach, Poland, were resistant to ampicillin (51). A high percentage of bacteria were reported as resistant to streptomycin (100%), cefazolin (89.8%), ampicillin (83.7%) and trimethoprim-sulfamethoxazole (69.4%), whereas a low percentage of bacteria were resistant to cefepime (12.3%) and meropenem (14.3%) in the aquaculture region of İskenderun Bay, Turkey (52).

In the current study, higher numbers of sulfonamide, rifampicin and ampicillin-resistant bacteria were recorded in the stations around aquaculture areas than other stations. Sphingomonas paucimobilis, Escherichia coli and Enterobacter cloacae isolated from both seawater and sediment at the stations around aquaculture areas had the highest levels of antibiotic resistance. The development of resistant pathogens in aquaculture environments is well documented (53, 54) and evidence of transfer of resistance encoding plasmids between aquaculture environments and humans has been presented recently (55). It has been reported that antibiotic‐resistant bacteria are present in a seafood ecosystem where antibiotics have never been used (56). This is interesting in terms of showing that aquaculture areas may be adversely affected by the presence of environmental antibiotic-resistant bacteria.

In the present study, a high percentage of the bacteria Sphingomonas paucimobilis were isolated, which was especially prevalent in Güllük Bay. The natural habitat of Sphingomonas has not been defined, but it is widely distributed in the natural environment especially in water and soil (57). The second most prevalent species were Escherichia coli and Enterobacter cloacae. Escherichia coli is an indicator of faecal contamination in aquatic environments. Enterobacter cloacae is the most frequent species associated with nosocomial infections along with Klebsiella pneumoniae that is a growing problem in human healthcare. The highest number of Bacillus cereus was isolated from the sediment underneath fish farms. A few Bacilli of marine origin have been reported to produce unusual metabolites different from those isolated from terrestrial bacteria (58). Due to the ubiquity and ability of the Bacillus species to survive under difficult circumstances, Bacillus strains are considered to be species of certain habitats (59, 60). In the current study, Bacillus pumilus, B. thuringiensis, B. mycoides and B. cereus were isolated from the sediment samples of the stations around fish farms.

The high frequency of resistance among bacterial isolates in the present study confirms the earlier reports regarding the role of antimicrobial use that plays a role in selecting antibiotic-resistant bacteria in water and aquatic sediments (46–52). Many previous studies have shown that the increases in antibiotic resistance in human medicine, agriculture and aquaculture are directly related to the amounts of antimicrobials used (61–65).

Infections caused by antibiotic-resistant bacteria are one of the most important public health concerns worldwide. Currently, MARs have been reported in a wide range of human pathogenic or opportunistic bacteria such as Vibrio sp. (66), Klebsiella pneumoniae (67), Salmonella sp. (68), Pseudomonas aeruginosa and also in pathogens (69, 70). Reservoirs of antibiotic resistance can interact between different ecological systems and potential transfer of resistant bacteria or resistant genes from animals to humans may occur through the food chain (70). In the current study, the MAR index of multiple antibiotic-resistant bacteria was found to be 2.6 times greater in the stations around fish farm areas (0.057) than the other stations (0.022).

Marine sediments offer more informative results than seawater about environmental pollution due to the accumulation of various pollutants at the bottom of the sea, therefore analysis of sediments is widely used in tests. The association of microorganisms with sediment particles is one of the primary factors in assessing microbial fate in aquatic systems. In this study, the bacteria isolated from sediment in all samples showed a higher resistance rate than bacteria isolated from seawater. Detection of higher antibiotic resistance in sediment bacteria than bacteria isolated from seawater showed that sediment bacteria were exposed to more antibiotics. Natural ecosystems containing high concentrations of heavy metals are also frequent. Heavy metal resistance genes are commonly found in environmental bacteria (71). The resistance to seven heavy metals has been reported in the order Cu > Mn > Ni > Zn > Pb > Cd > Fe for seawater bacteria isolated from the Golden Horn, Istanbul, Turkey (17). Heavy metal resistance in bacteria found in seawater from the Mediterranean has been reported as Cd > Cu > Cr = Pb > Mn; in Karataş, Turkey Cd > Cu > Cr = Mn > Pb; and İskenderun Bay, Cu > Cd > Mn > Cr > Pb (72).

In the present study, resistance to five different heavy metals (Zn²⁺, Pb²⁺, Cu²⁺, Cr³⁺ and Fe³⁺) were investigated for all isolates. Trends in heavy metal resistance vary depending on the sample sites: Güllük Bay, fish farm water column: Cu > Zn > Pb > Cr > Fe; sediment: Cr > Cu> Zn > Fe > Pb. Frequency of bacteria resistance to heavy metals shows the direct effects of metal pollution. Neisseria animaloris, Aeromonas caviae and Bacillus cereus isolated from sediment samples were the most tolerant of all the heavy metal salts. Chryseobacterium indologenes displayed the highest degree of sensitivity to all metal salts while Lactococcus garvieae showed the highest degree of sensitivity to Zn²⁺, Pb²⁺, Cu²⁺ and Fe³⁺. Kocuria kristinae, Escherichia coli and Acinetobacter lwoffii, which were isolated from the seawater underneath the fish farm, displayed similar sensitivities to all tested heavy metal salts. Resistances to heavy metals for Aeromonas and Pseudomonas isolates were similar to those from İskenderun Bay, with cadmium, 35.0% and 56.5%; copper, 98.3% and 75.4%; chromium, 38.3% and 31.9%; lead, 1.7% and 7.2%; manganese, 43.3% and 44.9%; and zinc 35.0% and 41.3%, respectively (72).

Both Gram-positive and Gram-negative bacteria can resist heavy metals (73). Resistance to toxic metals in bacteria probably reflects the level of environmental contamination with these substances and it may be related to the concentration of bacteria (74). The present project found heavy metal pollution in Güllük Bay sediment samples at all stations. In the sediment samples, the heavy metal contents were reported at varying rates: between 1 μg g⁻¹ and 209 μg g⁻¹ for lead; 10 μg g⁻¹ and 259 μg g⁻¹ for zinc; 1 μg g⁻¹ and 59 μg g⁻¹ for copper; 0.1 μg g⁻¹ and 46 μg g⁻¹ for chromium; <0.01 μg g⁻¹ and 2.8 μg g⁻¹ for cadmium; <0.01 μg g⁻¹ and 0.4 μg g⁻¹ for arsenic; and 0.6% and 5.9% for aluminium, respectively. The region was defined according to cadmium, lead and zinc levels as moderately polluted. Recorded high metal values were evaluated as an indicator of domestic and industrial inputs, carried via Sarıçay Creek, port operations and tourism activities within Güllük Bay (75). In the current study, the high frequencies of heavy metal-resistant bacteria detected in the sediment samples support this data. Bacterial heavy metal resistance detected in the study may depend on many factors. A possible explanation for differences in heavy metal resistance is the proximity of Güllük Bay to iron-steel factories. Additionally, Güllük Harbour is a serious pollution source. It was reported that 2862‐unit ships carried 4.8 million tonnes of ballast water to Güllük Harbour during 2007–2012 (37). Another potential source of increased resistance may be the discharge of thermal power plants located 107 km, 46 km and 39 km away from Güllük Bay. The effects of thermal power plant discharge on the accumulation of heavy metals have been reported in other studies (29, 75).

The association between antibiotic resistance and resistance to heavy metals is quite common in the same organism. The increasing numbers of antibiotic and heavy metal-resistant bacteria could be a result of gene transfer activities demonstrating that industrial pollution most likely selects for antibiotic resistance and vice versa (58). In this study, similarly, the most antibiotic-resistant bacteria such as Sphingomonas paucimobilis, Escherichia coli and Enterobacter cloacae were also resistant to heavy metals. Metal‐resistant isolates from Güllük Bay also showed high resistance to sulfonamide, rifampicin and ampicillin. Bacteria from different sources such as humans, animals and soil can transfer or exchange their resistance genes. At the same time, water contaminated with antibiotics, disinfectants, pesticides and heavy metals might encourage selection and result in antibiotic and heavy metal resistance. Marine environmental conditions are extremely dynamic compared to the terrestrial environment, allowing bacteria to bring resistance mechanisms they have developed together while being adapted to the varying conditions. This makes the isolation of various bacteria useful to assess environmental pollution and provides a pathway to possible solutions to remove pollution from marine environments. For bacteria to take part in the transformation of any heavy metal salt into a harmless form, those bacteria must firstly be resistant to the heavy metal; thus the data related to frequency of metal resistant bacteria can provide knowledge on the continual accumulation or transformation of heavy metals in the marine environment.

The findings of the current study provide data regarding the distribution of heavy metal- and antibiotic-resistant bacteria in seawater and sediment samples of Güllük Bay, Aegean Sea, Turkey. As a result, preliminary data on candidate bacteria will offer opportunities for further studies on the elimination of heavy metal contamination by the detection of heavy metal-resistant bacteria.

Analyses of the presence of antibiotic resistance in bacteria provide knowledge on pollution sources such as septic systems on regional ecosystems. Since antibiotic-resistant bacteria can affect pathogen virulence, these pollution sources can induce pathogens and can create health risks for both humans and the ecosystem. In the present study, bacteria resistant to antibiotics and heavy metals in seawater and sediment were investigated. The bacterial information obtained provides essential data for identifying the regional distribution of resistant bacteria. Levels of resistance against heavy metals and antibiotics in bacteria isolated from seawater and sediments of the Aegean Sea were quantified. Bacteria isolated from Güllük Bay sediment were resistant to all antibiotics tested and exhibited higher resistance than those isolated from seawater. The frequency of antibiotic-resistant bacteria was higher around fish farms and near the exit of Sarıçay Creek. The widespread resistances of indicator bacteria to antibiotics suggest the presence of anthropogenic influences due to domestic waste and maritime transport.

In order for bacteria to take part in the transformation of heavy metal salts into harmless forms, they must initially be resistant to heavy metals. The frequency of resistance thus provides information regarding the continual accumulation or transformation of heavy metal salts in the marine environment. The findings of the present research have shown the existing contamination status of Güllük Bay via heavy metal and antibiotic resistance tests. The study region is under pressure of pollution as stated in previous research (25, 26, 75) and the bacterial resistance data of the current study showed that there is a prevalence of resistant bacteria in the region that may be due to indirect effects of environmental dynamics and pollution.

In this study, the presence of higher levels of resistant bacteria in sediment compared to seawater may indicate the presence of microplastics in the sediment as well as the probability that the sediment is a suitable medium for accumulation of metals and antibiotics. Further studies on this subject will provide detailed data on the spread of antibiotic- and metal-resistant bacteria in marine sediments.

The present study showed bacterial responses to environmental stress and influences in terms of antibiotic and heavy metal resistance both in sediment and seawater samples at Güllük Bay, Turkey. These findings highlight the necessity of holistic assessments with a ‘one health’ approach and the need to control bacteria entering marine areas due to human activities, considering the contributions of resistant bacteria to global distribution. The data may also provide a useful resource to help identify strains of bacteria for environmental remediation applications.

Three soak liquor samples were collected from Istanbul Leather Organized Industrial Zone, Tuzla, Istanbul, Turkey. These samples were immediately placed into sterile sample bags and carried on ice during transportation. Direct and serial dilutions were spread onto nutrient agar plates. The morphologically different colony was picked up to obtain the pure culture of the isolate and was numbered as Isolate 1.

Gram staining, catalase, oxidase, lipase and protease activities were examined. Protease activity of Isolate 1 was examined on gelatin agar medium containing 2% gelatin (w/v). The agar plates were flooded with Frazier solution following 24 h incubation. Clear zones around the colonies were evaluated as positive for protease activity. Lipase activity was tested on Tween^® 80 agar medium containing 1% (w/v) Tween^® 80. After incubation, opaque zones around the colonies were accepted as evidence of lipase activity (29, 30).

Genomic DNA of Isolate 1 which was determined to have protease and lipase activities were extracted by phenol/chloroform extraction and ethanol precipitation. DNA isolation was confirmed by agarose gel electrophoresis. DNA samples were stored at −20°C until use. The 16S rRNA gene was amplified by polymerase chain reaction (PCR) with the universal bacterial primers 27F (5-AGAGTTTGATCMTGGCTCAG) and 1492R (5-TACCTTGTTACGACTT). Negative control was included in PCR amplifications. PCR amplification was carried out by an initial denaturation at 95°C for 4 min, followed by 30 cycles at 95°C for 1 min, 57°C for 1 min and 73°C for 1 min. The reactions were finished by a final extension at 73°C for 7 min. The PCR products were also monitored by agarose gel electrophoresis. These products were purified by GeneJET^TM Gel Extraction Kit (Thermo Scientific^TM, Thermo Fisher Scientific, USA). These purified samples were analysed by Medsantek Ltd Co, Istanbul, Turkey. The 16S rRNA sequence contigs were generated by the software ChromasPro version 2.1.8 (Technelysium Pty Ltd, Australia). Then, consensus sequences were exported in FASTA format for each sample for data analysis. These sequences were compared with sequences in the National Center for Biotechnology Information (NCBI) using the Basic Local Alignment Search Tool (BLAST^®) search program.

The lichen samples belonging to H. tubulosa, H. physodes, E. divaricata, P. furfuracea, P. sulcata and Usnea sp. were collected from fir trees of Kastamonu province in the north-west of Turkey. They were identified through classical taxonomical methods by microscopic examination.

H. tubulosa, H. physodes, E. divaricata, P. furfuracea, P. sulcata and Usnea sp: Turkey, Kastamonu province, Kapaklı Village, 41.24492, 34.18330, G. Çobanoğlu.

The experiment steps included washing, drying in air, weighing, pulverising by liquid nitrogen, adding acetone (ACS, ISO, Reag. Ph. Eur.), keeping in a dark place for 24 h followed by filtration through filter paper. Then, the evaporation of acetone in a rotary evaporator was performed and crude lichen acetone extracts were obtained (27).

The test isolate was grown on Tryptic soy agar media at 37°C for 24 h. The tests were performed in 96-well CELLSTAR^®, F-bottom microplates with lid (Greiner Bio-One GmbH, Austria). Tryptic soy broth was added to each well and nine-fold serial dilutions of the acetone extracts of H. tubulosa, H. physodes, E. divaricata, P. furfuracea, P. sulcata and Usnea sp. were made. Final concentrations of all lichen extracts were 240 μg ml⁻¹, 120 μg ml⁻¹, 60 μg ml⁻¹, 30 μg ml⁻¹, 15 μg ml⁻¹, 7.5 μg ml⁻¹, 3.75 μg ml⁻¹, 1.9 μg ml⁻¹ and 0.9 μg ml⁻¹. Overnight culture of the isolate was added to obtain a total volume of 100 μl with an optical density (OD) 600 nm of 0.01. The experiments included untreated and blank controls. The tests were performed in three replicates. Bacterial growth ratios at an OD 600 nm were measured using Cytation^TM 3 Multi-Mode microplate reader (BioTek Instruments Inc, USA).

Two deteriorated salted sheepskins containing red discolorations were collected from two tanneries in the Istanbul Leather Organized Industrial Zone (40°52′39.7″N,29°20′25.3″E) in Tuzla, Turkey. The samples were immediately placed into sterile sample bags and transported on ice to the laboratory. Then, 20 g of the salt-pack cured sheepskin samples were weighed and separately soaked in flasks containing 180 ml 30% NaCl (Merck KGaA, Germany) solution. The flasks were placed into a shaking incubator at 90 rpm, 24°C for 3 h. The suspension of the skin was diluted with sterile physiological saline water (30% NaCl). An aliquot of 100 μl each of direct and serial skin suspension dilutions was spread onto the surface of modified Brown agar media containing (per litre): 1 g CaCl₂·H₂O, 2 g KCl, 20 g MgSO₄·7H₂O, 3 g trisodium citrate, 250 g NaCl, 5 g yeast extract, 20 g agar, pH 7 (5, 27). The plates were incubated at 39°C for 10 days. Following incubation, red pigmented colonies on the agar media were selected and restreaked several times to obtain pure cultures. A total of 22 isolates were obtained from the sheepskins and then, these strains were examined the proteolytic and lipolytic activities. Proteolytic activity of each strain was detected on gelatin agar medium containing 2% gelatin. After incubation, clear zones around the colonies on the gelatin agar medium indicated protease production (5, 10). Lipolytic activity of each strain was screened on Tween^® 80 agar medium containing 1% Tween^® 80. After growth was obtained, opaque zones around the colonies were interpreted as positive lipase activity (5). In the present study three red pigmented proteolytic and lipolytic strains (AT1, 22T6 and 7T3) were obtained from two salted sheepskins and these strains were used in the present study.

Exponentially growing pure cultures of three strains designated as AT1, 22T6 and 7T3 were used in all experiments. First, the strains’ salt requirement and salt tolerance were examined on Brown agar plates containing different salt concentrations (0%, 0.5%, 3%, 5%, 7.5%, 10%, 12.5%, 15%, 20%, 25% and 30%) (27). After detection of optimum salt concentration for each strain, pH and temperature ranges for growth of each strain (AT1, 22T6, 7T3) were respectively examined at Brown agar plates with different pH values (pH 4, pH 5, pH 6, pH 7, pH 7.5, pH 8, pH 9, pH 10, pH 11 and pH 12) and different temperatures (4°C, 10°C, 15°C, 24°C, 28°C, 35°C, 37°C, 39°C, 45°C, 50°C, 55°C, 60°C) according to the methods described in Proposed Minimal Standards for Description of New Taxa in the Order Halobacteriales (28). Based on the pH, and temperature range of each test strain, the optimal pH and growth temperature of each test strain were determined.

Pigmentation, size, margin, elevation and opacity of colonies of the strains grown on Brown agar media were examined under optimal growth conditions (28). Cell morphology, cell length, cell width and motility of each strain were examined using both light microscopy and electron microscopy. Microscopic observation of each strain was made by using freshly prepared wet mount (28). For SEM observations, 20 ml of each test strain were separately passed through 0.2 μm pore size cellulose nitrate membrane filter placed in the stainless steel funnel via vacuum pump (Sartorius AG, Germany). The archaeal cells of each strain trapped on the membrane filters were observed under SEM (Quanta^TM 450 FEG (FEI, USA)). Gram staining was performed with acetic acid-fixed slides (28–30). Catalase and oxidase activities, indole production, methyl red test, H₂S and NH₃ productions of each strain were investigated according to the procedures described previously (4, 28, 31). Furthermore, each strain’s caseinase activity was determined on the agar medium containing 2% skim milk. After incubation, clear zones around the colonies were evidence of positive caseinase activity (4). Urease production was investigated on Christensen urea agar medium. The tubes were examined for pink or red colour change in the medium after seven days of incubation (28, 31). β-galactosidase activity was screened in test tubes containing ortho-nitrophenyl-β-galactoside (ONPG) discs and 1 ml of sterile saline water (30% NaCl). The yellow colour formation in the test tube was accepted as positive β-galactosidase activity (5, 31). Amino acid utilisation of each strain was examined in the test medium containing 1% amino acid, 0.5% beef extract, 0.5% peptone, 0.05% dextrose, 0.0005% cresol red, 0.001% bromocresol purple, 0.0005% pyridoxal and saline water (30% NaCl). Purple colour formation in the test tube containing archaeal culture was accepted as a positive test after 10 days incubation period at 39°C (31).

Chromosomal DNA was isolated by QIAamp DNA Mini Kit (Qiagen, Germany) and purified by QIAquick PCR Purification Kit (Qiagen, Germany) according to the manufacturer’s directions. The 16S rRNA genes of the strains were amplified by polymerase chain reaction (PCR) using forward primer 21F and reverse primer 1492R (32). The 16S rRNA gene sequences of three strains (AT1, 22T6 and 7T3) were determined by IONTEK Laboratory (Turkey). The sequences of these strains were analysed using ChromasPro v.2.1.8 software (Technelysium, Australia) and then compared with the sequence on the EZBioCloud Database (ChunLab, South Korea) (33).

2.4.1 Preparation of Strains and Cultures Used in Brine Curing Processes

2.4.2 Preparation of Sheepskin Samples for Brine Curing Treatments

To determine total counts of extremely halophilic archaea in the curing solutions before the curing processes, 100 μl of the test medium was removed from the each curing solution and diluted to 10⁻²–10⁻⁴ using sterile 30% NaCl solution. The diluted archaeal suspensions were spread over the Brown agar media. In addition, subsequent to each brine curing process detailed above (T1–T5), the suspensions of cured sheepskin samples were prepared at intervals of 5 days, 16 days, 28 days and 47 days of storage. 2 g of each skin sample were put into a flask containing 18 ml sterile 30% NaCl solution and incubated for 1 h at 24°C and 100 rpm. Direct and serial dilutions of the suspensions were spread onto the surface of Brown agar media. All inoculated Brown media were incubated at 39°C for 10 days and the colonies grown on the test media were counted.

After curing processes, 5 g of the sheepskins were cut and placed into flasks containing 100 ml of sterile distilled water. The flasks were placed in a shaking incubator for 1 h at 100 rpm and then pH was measured with a pH meter. Hairs and dirt on the samples were removed to properly determine the samples’ moisture content. 3 g of the samples were placed into an oven at 102°C for 6 h. The dried samples were weighed, returned to the oven for 1 h, and then were weighed again. The drying procedure was repeated until the first dry weight was equal to the second dry weight. The samples were put into a desiccator for 30 min to cool. Next, we calculated the skins’ moisture contents (20, 21). The dry sheepskins samples were placed in ceramic crucibles and ashed in a muffle furnace at 600°C for 8 h. After cooling, the samples were weighed to determine ash content. Moisture content, ash content and salt saturations of skin samples were calculated according to the aforementioned methods (30, 36). The pH value, ash content, moisture content and salt saturation of all cured sheepskin samples were examined at different storage periods.

All cured sheepskin samples were examined organoleptically (hair slip, deterioration of skins, bad odour, sticky appearance, red heat, hole formation) during different storage periods.

After a 47-day storage period, the sheepskin samples were prepared for SEM observation. The samples were fixed in 4% glutaraldehyde solution prepared in 0.1 M phosphate buffer (pH 7.2) for 30 min. The samples were washed three times with 0.1 M phosphate buffer for 10 min and were treated with 1% OsO₄ prepared in 0.1 M phosphate buffer at room temperature for 1 h. The samples were washed two times in sterile distilled water for 10 min. Then, the water in the sheepskins was gradually removed by 35%, 50%, 75%, 95% and absolute ethanol. The mixtures of ethanol-hexamethyldisilazane (ethanol-HMDS) [1:1 (v/v)] (1 × 30 min), ethanol-HMDS [1:2 (v/v)] (1 × 30 min) and HMDS (2 × 30 min) were used for air drying process. After drying, HMDS was poured from petri dishes and the samples were placed in a desiccator for 12 h. Later, the sheepskin samples were examined under SEM (Quanta^TM 450 FEG) using sample stub with double-sided sticky tape (37).

A total of 22 red coloured strains were isolated from two deteriorated salted sheepskin samples obtained from two tanneries in the Istanbul Leather Organized Industrial Zone in Tuzla, Turkey. While nine, seven and three strains respectively produced protease, lipase, both protease and lipase, three strains did not produce either lipase or protease enzymes. The red coloured three extremely halophilic strains producing both protease and lipase enzymes were selected and used as test strains (AT1, 22T6 and 7T3) in the present study.

Strains AT1, 22T6 and 7T3 grew at 15–30% NaCl, 15–30% NaCl, 20–30% NaCl concentrations, respectively. Optimum salt concentrations of strains AT1, 22T6 and 7T3 were determined as 25% NaCl. Hence, these strains were accepted as extremely halophilic archaea. The pH and temperature ranges for growth of strains AT1, 22T6 and 7T3 were respectively found as pH 6–11 and 20–50°C, pH 6–11 and 15–55°C, pH 5–11 and 15–55°C. All extremely halophilic archaeal strains optimally grew at 39°C and pH 7. The colony pigmentation, size, margin, elevation and opacity of strains AT1, 22T6, 7T3 were respectively observed as: red, 0.6– 2 mm, entire, convex, translucent; red, 1–2 mm, entire, convex, translucent; red, 0.8–1.9 mm, entire, convex, translucent. The cells of strains AT1 (Figure 2(a)) and 7T3 (Figure 2(c)) were non-motile, extremely pleomorphic (triangle, square, irregular disk, short rod). The cells of strains AT1 and 7T3 were approximately 0.4–1.3 μm × 0.4–2.0 μm and 0.3–0.7 μm × 0.3–4 μm, respectively. The cells of strain 22T6 (Figure 2(b)) were motile, pleomorphic rods, approximately 0.5–1.2 μm × 3.2–6.6 μm. All strains were Gram-negative (Table II). While all strains showed positive catalase, oxidase, protease, lipase activities, indole production, the methyl red, caseinase, urease and β-galactosidase reactions of all strains were negative. The strains did not produce H₂S and NH₃ (Table II).

Fig. 2

Our experimental results showed that Haloarcula salaria (AT1), Halobacterium salinarum (22T6), Haloarcula tradensis (7T3) strains have protease activities which can breakdown proteins in corium of sheepskin causing loss of skin substance. When the protein structure of salted skins is broken down by proteolytic extremely halophilic archaea, these microorganisms can utilise some amino acids as a source of carbon, nitrogen and energy. Haloarcula salaria AT1 and Halobacterium salinarum 22T6 utilised most of the amino acids examined. While Haloarcula salaria AT1, Halobacterium salinarum 22T6 utilised 17 amino acids, Haloarcula tradensis 7T3 used only three amino acids (Table III). In another study, the liquid test media containing calfskin samples, 30% NaCl and proteolytic red and pink strains of the extremely halophilic archaea were separately prepared to show disintegration of the skin proteins. After an incubation period, decomposition of the skin samples in the media was detected by visual observation. While contents of asparagine, threonine, serine, glutamine, proline, glycine, alanine, valine, isoleucine, leucine, phenylalanine, lysine and arginine in the test tubes were detected at high levels, contents of methionine, tyrosine and histidine were low (10).

Phenotypic features of extremely halophilic AT1, 7T3 and 22T6 strains detected in this study were fairly similar to phenotypic features of Haloarcula salaria, Haloarcula tradensis and Halobacterium salinarum isolated by other researchers (15, 38, 39).

The phylogenetic analysis revealed that three strains shared highly similar identities with their closest phylogenetic relatives. Strains AT1, 22T6, 7T3 were respectively assigned to Haloarcula salaria (98.36%-1344 base pairs), Halobacterium salinarum (99.78%-1345 base pairs), Haloarcula tradensis (98.37%-1355 base pairs). The gene sequence data of the strains AT1, 22T6, 7T3 were respectively deposited in GenBank^® (National Center for Biotechnology Information, USA) under accession numbers as MN585896, MN585803, MN585804.

In our previous study, extremely halophilic archaeal strains were isolated from Tuz Lake and its salterns (5). In Turkish leather industry, curing salt is mostly obtained from Tuz Lake and its salterns. Hence, we suspect that contaminations of our sheepskin samples with Haloarcula salaria AT1, Halobacterium salinarum 22T6 and Haloarcula tradensis 7T3 were due to the curing salt obtained from Tuz Lake and its salterns.

In the study carried out with 25 salted sheepskin samples (Australia, Bulgaria, Dubai, Greece, Israel, Kuwait, South Africa, Turkey, USA) and 25 salted goat skin samples (Australia, Turkey, Bulgaria, Israel, South Africa, Russia, China, France), proteolytic extremely halophilic archaea and lipolytic extremely halophilic archaea were detected as 10²–10⁵ CFU g⁻¹; 10²–10⁶ CFU g⁻¹ and 10²–10⁶ CFU g⁻¹; 10²–10⁶ CFU g⁻¹ on salted sheepskins and goat skins, respectively (40). The highest number of proteolytic and lipolytic extremely halophilic archaea on the salted skins was found as 10⁶ CFU g⁻¹ (40). Therefore, the archaeal cell numbers of test strains in the brine curing solutions were adjusted to 10⁶ CFU ml⁻¹. Before the curing processes of sheepskins, while the archaeal cell numbers in the brine solutions of Treatments 1, 3 and 4 were detected as 2.1 × 10⁶ CFU ml⁻¹, the archaeal cell numbers in the brine solution of Treatment 2 was detected as 2.2 × 10⁶ CFU ml⁻¹.

The archaeal cell numbers in the mixed culture was detected as 2.1 × 10⁶ CFU ml⁻¹ in the electrolysis cell before 1.5 A DC application. While the archaeal cell numbers in the mixed culture were reduced from 2.1 × 10⁶ CFU ml⁻¹ to 3.2 × 10⁵ CFU ml⁻¹ after 1 min of DC treatment, the cell numbers of 1.24 × 10² CFU ml⁻¹ was detected after 4 min of DC treatment. All archaeal cells in the mixed culture were completely killed in 7 min of DC treatment. In the present study, log₁₀ value of the mixed culture of extremely halophilic archaea in the brine solution before the DC treatment was 6.32. After 1 min, 4 min and 7 min of 1.5 A DC treatment; 0.82, 4.23 and 6.32 log₁₀ reduction values (CFU ml⁻¹) of the mixed culture in the brine were detected, respectively.

Temperature and pH of the electrolysis cell were respectively measured as 31°C and pH 6 prior to the electric current treatment. After treating the brine solution with the electric current, the temperature of the brine was adjusted to 24°C for using in curing process of sheepskin in the Treatment 5. While the temperature and pH of the test medium respectively increased from 31°C to 41°C and from pH 6 to pH 8.5 during the electric current treatment, voltage values slightly decreased from 4.7 V to 4.3 V.

We also demonstrated the inactivation of extremely halophilic strains via DC and alternating electric current (AC) in our previous studies (35, 41, 42). A 0.5 A DC was applied for 30 min to several strains of extremely halophilic archaea (10⁷ CFU ml⁻¹) isolated from Tuz Lake, Kaldırım and Kayacık salterns (35). While the mixed culture of extremely halophilic archaea was exterminated in 10 min, protease producing extremely halophilic archaea were killed in 5 min. However, lipase or lipase and protease producing extremely halophilic archaea were exterminated in 20 min (35). In another experiment, lipase and protease producing extremely halophilic strains (10⁵–10⁶ CFU ml⁻¹), separately grown in liquid Brown media, were inactivated by a 10 min treatment with 0.5 A DC (41). It was also detected that 1 min of 2 A AC treatment was enough to kill extremely halophilic archaea found in brine solution (10²–10⁴ CFU ml⁻¹). When 2 A AC was applied to lipolytic extremely halophilic archaea, proteolytic extremely halophilic archaea, both proteolytic and lipolytic extremely halophilic archaea, and a mixed culture of these strains (10⁶ CFU ml⁻¹), all test microorganisms found in 25% NaCl solution were exterminated in 5 min (42).

After the curing processes of sheepskins, we did not detect any extremely halophilic archaea on the sheepskin sample cured with the sterile brine solution (Control) and the sheepskin sample cured with the brine solution containing electrically inactivated mixed culture (T5) during the all storage periods.

While extremely halophilic archaeal numbers on both skin samples cured with each strain and the skin sample cured with mixed cultures of the strains slowly increased from 10⁶ CFU ml⁻¹ to 10⁷ CFU during five days and 16 days storage periods, the numbers of these strains slowly decreased 28 days and 47 days storage periods due to attachment of these cells to sheepskins (Table IV).

After the curing processes of skins, pH values of the sheepskin samples were measured as pH 7.35 for Control; pH 6.89 for T1; pH 7.09 for T2; pH 7.05 for T3; pH 7.16 for T4; pH 8.05 for T5. While salt saturation values of all cured sheepskins were higher than 100% during all storage periods, pH, ash content and moisture content values changed during different storage periods. pH, ash content and moisture content values of the cured skins were detected between pH 6.52–7.81, 20–44%, 24–57%, respectively (Table IV).

Moisture, minimum and maximum ash contents, salt saturation values of adequately cured salted hides were suggested as 40–48%, 14–48%, higher than 85%, respectively (36). Due to detection of high moisture content in all samples (between 50–57%) after five days storage, sterile salt was added to all sheepskins to reduce their moisture contents according to curing procedure described in the previous study (43). While all skin samples reached the suggested moisture content values (39–46%) after 28 days, the suggested saturation values were detected after five days. The samples’ lowest moisture content values were detected after 47 days. Ash contents of all skins (20–44%) were close to suggested values (36). While the skins’ pH values changed during storage periods, all values were found sufficient to support the growth of extremely halophilic strains (Table IV). The pH, moisture content, ash content and salt saturation values detected in this study were also consistent with pH range (pH 6.53–8.01), moisture content (32–68%), ash content (12–30%) and salt saturation (58–100%) values of 25 salted sheepskin samples determined in the previous experiment (40).

While hair slip and bad odour were detected on the sheepskin samples cured with each strain and the mixed culture after five days at 33°C, sticky appearance and red heat were observed on the cured sheepskin samples after 16 days (T1–T4, Figure 3). In addition to the aforementioned organoleptic properties, hole formations were observed on these sheepskin samples after 28 days. However, we did not detect any organoleptic properties on sheepskin samples cured with sterile brine and the brine treated with 1.5 A DC (Control and T5, Figure 3).

Fig. 3

In another study, the commercially cured hides stored one year in the USA were also examined for proteolytic activity of extremely halophilic archaea. Experimental results of that study showed that the flesh side of hides containing extremely halophilic archaea had pink discolorations called red heat. When these hides were incubated at 35°C–40°C, bad odour, hair slip and severe grain damage were detected. Damaged grain surfaces were observed on leather made from these hides (10). In another experiment researchers emphasised that temperatures of the brines and hides should be maintained below 20°C to prevent growth of extremely halophilic archaea (44).

Figure 4 shows extremely halophilic archaeal cells of the mixed culture on 0.2 μm pore-size cellulose nitrate membrane filter in pleomorphic shapes such as triangle, square, irregular disk and rod. As seen in the SEM micrograph, 1.5 A DC treatment significantly debilitated structural integrity of the cells in the mixed culture trapped on the filter (Figure 5). The SEM images clearly showed that electric current application damaged cell structures of each strain in the mixed culture (Figure 5). As seen in Figure 6, the sterile brine curing process protected the sheepskin against microbial damage during 47 days of storage.

Fig. 4

Fig. 5

Fig. 6

Attachment of Haloarcula salaria AT1, Halobacterium salinarum 22T6 and Haloarcula tradensis 7T3 to corium fibres and the consequent destructive effects on sheepskins are seen in Figures 7–10. Haloarcula salaria AT1, Halobacterium salinarum 22T6 and the mixed culture of the strains caused fibres in the corium to split and weaken (Figures 7, 8 and 10). In contrast with the skin samples treated with Haloarcula salaria AT1, Halobacterium salinarum 22T6, skin sample treated with Haloarcula tradensis 7T3 had compact appearance, although the shredding of the fibres was still present in corium (Figure 9). That damage was due to the proteolytic activities of these microorganisms.

Fig. 7

Fig. 8

Fig. 9

Fig. 10

Figure 11 clearly shows that the curing process of sheepskin with the brine containing mixed culture treated with DC prevented extremely halophilic archaea from contaminating the sheepskin and furthermore protected the skin very well against microbial damage during a long storage period.

Fig. 11

The present study proved that organoleptic changes detected in the sheepskins were closely related to proteolytic and lipolytic activities of extremely halophilic archaeal strains on the skin. Electron micrographs also showed that each test isolate and a mixed culture of extremely halophilic strains destroyed the skins’ collagen fibres. We did not detect any difference when assessing the efficacy of sterile brine and electrically treated brine curing processes of sheepskin samples throughout 47 days. We did not observe any damage to the compactness of sheepskin structure cured with both the sterile brine and electrically treated brine containing the mixed culture. Both methods were found very effective for preventing archaeal growth and damage on the brine cured sheepskins.

Our results were consistent with those of other experimental studies on the extremely halophilic strains and culture collection strains of extremely halophilic archaea (25, 26). In our previous experiment, SEM images showed that hides cured with proteolytic extremely halophilic archaeal strains had red heat and severe grain damage after 49 days of storage at 41°C (25). In another study, the cured hides with extremely halophilic Haloferax gibbonsii (ATCC^® 33959^TM) exhibited hair loss, thinner and flaccid structure; these consequences of deterioration and loss of hide substance. The open fibre structure was also detected in the corium of the hide inoculated with Haloferax gibbonsii (27). The SEM images showed that the fibre structures of hide were broken down into the smaller fibres after 43 days (27).