Microbial collagenases: challenges and prospects in production and potential applications in food and nutrition

Abstract

Microbial collagenases are promising enzymes in view of their extensive industrial and biological applications. The emerging evidence of potential food applications and health benefits have revealed that microbial collagenase shows significant promise as a main component for bioactive functional ingredients and the preparation of peptides. Collagenases are important virulence factors which play a crucial role in the global degradation of the extracellular matrices of animals, due to their collagen degradation ability. There is a lack of scientific consensus on a well-defined and proper screening of collagenase-producing microorganisms. A vast controversy can be found in the literature regarding the correct identification of microbial collagenase. This review summarizes the current technologies and strategies used to improve the screening, production, and purification of microbial collagenase with a comprehensive insight, especially focusing on the classification, structures and collagen-degrading mechanisms of M9 family representative collagenases. It also highlights the potential of microbial collagenase in the development of a process for meat tenderization and bioactive “true” collagen peptides, or the preparation of hydrolysates. In addition, the critical challenges and various strategies for potential applications of collagenase in food, nutrition, biotechnological and medical sectors are highlighted.

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Gaurav Kumar Pal

Gaurav Kumar Pal is a PhD scholar in the Department of Meat and Marine Sciences at the CSIR-Central Food Technological Research Institute, Mysuru, India under the guidance of Dr Suresh PV. He received his Master's degree in Applied Microbiology from Chaudhary Charan Singh University, Uttar Pradesh, India. He has authored/co-authored a number of research articles in SCI journals and a book chapter (CRC Press, USA). He is a recognized reviewer of Elsevier, Wiley, Springer, and other journals. His research is focused on microbial enzymes with specialization in collagenase and its application in seafood collagen-based ingredients for functional food/beverage products.

Suresh P. V.

Dr Suresh PV is a Principal Scientist and Associate Professor (AcSIR) in the Meat and Marine Sciences Department, CSIR-Central Food Technological Research Institute, Mysuru, India. He received his PhD in biotechnology from Cochin University of Science and Technology, Kerala, India in 1997. His research covers multidisciplinary areas, including microbial enzymes, solid state fermentation, food microbiology, and functional evaluation of marine biopolymers/molecules. He has more than 25 years of research/academic experience with over 50 peer-reviewed articles, posters/public presentations, book chapters and patents. Currently, his research team focuses on microbial enzymes and marine derived functional molecules such as chitin, chitosan and collagen.

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1. Introduction

Collagens are the major structural proteins of connective tissues such as skin, bone, cartilage, tendon, and blood vessels in mammals. Collagen is composed of three α-chains of predominantly repeating Gly-X-Y triplet sequences (X and Y are often proline and hydroxyproline), which induce each α chain to maintain a left-handed helix with remarkable primary, secondary, tertiary and quaternary structures.^1–4 There are 29 distinct collagens, and each collagen differs considerably in its amino acid sequence, structure, and function.^1–3,5,6 The proteolysis of collagen is essential for numerous physiological functions, including tissue remodeling, morphogenesis, and wound healing. Collagen has been recognized as an essential contributing factor in numerous pathologies, such as tumor cell spreading, arthritis, tissue ulceration, cardiovascular disease, neurodegenerative disease, and periodontal disease.^1–3,6 The commercial applications of collagen and collagen derived products are increasing at a rapid rate as ingredients in beverages, foods, drugs, cosmetics, and a variety of health care products.^4,7–10 Due to the decrease in collagen synthesis within the body, the demand for collagen in skin, hair and bone tissues increases with aging. It has been reported that collagen peptides, a product of collagen degradation, possess several biological activities of industrial, food, nutritional and biomedical interest.^8,29,30 Furthermore, collagen degradation products reportedly have various activities of medical and pharmaceutical interest as agents for osteoporosis, gastric ulceration, hypertension, skin moisturizers, and preservatives. Collagen degradation products can be added to food and beverage products to improve their functional and nutritional properties without causing any production problems due to their low viscosity and high solubility in water.⁷

Type I collagen is the most abundant and commercially important type of collagen; it is used widely in the food, biomedical, pharmaceutical and cosmetic industries due to its excellent cell adhesion properties, high biocompatibility, biodegradability, and weak antigenicity.^4,7–12 Collagens and their fibrils with other polymers can also be used to prepare porous scaffolds. These scaffolds have been extensively used in tissue engineering and biomedical sectors.^13–19 At present, the skin and bones of land-based animals (bovine and porcine) are the chief sources of commercial collagen, although epidemics of bovine spongiform encephalopathy (BSE), transmissible spongiform encephalopathy (TSE) and foot-and-mouth disease (FMD) have caused anxiety among consumers of collagen and collagen-derived products from these land-based animals.⁴ Furthermore, collagens and collagen-derived products from bovine and porcine animals are forbidden in some religions.⁴ Consequently, the demand for realistic collagens from substitute sources, particularly of aquatic/marine origin, has been increasing in recent years.⁴

Enzymes have been used an integral part of various industries for a long time. Enzymatic methods constitute an essential and important part of production processes due to their highly specific nature and high activity at very low concentrations under mild conditions of pH and temperature, which may also result in fewer unwanted side-effects and by-products.²⁰ Microbial collagenases are secreted by anaerobic/aerobic pathogenic and non-pathogenic microorganisms to utilize collagen as a source of nutrients.²¹ The ‘native’ collagens are resistant to most common proteases due to their triple-helical structure but are readily cleaved at a specific site by collagenases. In literature, many enzymes that were originally described as collagenases were shown later to be either proteases or peptidases of broad or different specificity. The collagenases are found in animals (EC 3.4.24.7) and microorganisms (EC 3.4.24.3), and differ in their substrate specificity. Animal collagenase splits/digests native triple-helical collagen at a single peptide bond.³⁷ However, microbial collagenase is unique and capable of hydrolyzing native collagen under physiological conditions. Collagenases possess broad substrate specificities to degrade both water-insoluble ‘native’ collagens and water-soluble denatured collagens.²⁷ Due to this unique activity, microbial collagenases play important roles in embryo development, morphogenesis, tissue remodeling, wound healing, and human diseases, such as arthritis, cancer, and atherosclerosis.^6,28 Therefore, microbial ‘true’ collagenases have gained much attention for their numerous industrial, biotechnological, pharmacological, medicinal and food applications.^3,8–10,21

Collagenases are used in various sectors, such as food, tanneries, fur, fish processing, brewing, meat processing, clarification and stabilization of beer, medicine, cosmetics, fish silage, fish sauce, fish meal, and in the processing of animal feedstuffs as well as in scientific and analytical research.^21,31–34 Hence, screening of high yielding enzyme-productive microorganisms and development of low-cost cultivation media, as well as different cultivation methods for economic production of collagenolytic enzymes with novel properties, are being intensively pursued by the scientific community.^9,21,36 In fact, this review is intended to highlight pertinent information related to an overview of microbial collagenases and the trends and prospects for the future, with special emphasis on the outlook for the potential applications of these collagenases in food industries. This review also discusses new strategies that can be used for the development of novel functional food and beverage products using collagen hydrolysate.

2. Collagenases: occurrence and distribution of collagenase

Collagenases have been usually recognized as enzymes that specifically attack ‘native’ and water-soluble denatured collagens.^21,27 The term ‘true collagenase’ was coined on the basis of their capacity to hydrolyze/digest native bovine achilles tendon collagen.²¹ The collagenases are mainly found in animals, microorganisms and plants (Fig. 1). The occurrence of collagenases in plants also has been reported.^22,23,36

Fig. 1 Major collagenases and different sources of microbial collagenases.

Collagenase is one of the most widely used enzymes in several applications. In the past, much attention has been given to the isolation and extraction of collagenases from animal tissues. Recently, emphasis has shifted to obtaining microbial collagenases, which are advantageous in comparison with animal collagenases.^{8–10,21,45,46} At the industrial scale, this enzyme is obtained from the pathogenic microorganism Clostridium sp. Weinberg and Randin reported the first evidence for bacterial collagenase production in 1932.²¹ In 1937, Maschmann suggested the name collagenase for an enzyme from C. perfringens that can digest both ‘gelatin’ and ‘collagen’.²¹

2.1. Plant collagenase

Collagenases are mostly found in animals and microorganisms. Collagenase has also been reported from a few plants.^22,23,36 The plant collagenases preferentially cleave the native form of collagen at a specific site. These enzymes have not been shown to be involved in biological processes; however, they may be related to a biological function, such as defense against pests such as nematodes.²² The collagenolytic activity of plants may play a significant role as a line of defense against environmental changes, and their activation can be triggered by applying various types of stress.²² The isolation and characterization of collagenase from the ginger rhizome (Zingiber officinale)²³ and fig (Ficus carica var. Brown Turkey) latex³⁶ has been reported.

2.2. Animal collagenase

Animal collagenase (EC 3.4.24.7, interstitial collagenase) splits/digests collagen at a single peptide bond across the three α chains organized in the native triple-helical structure. Animal collagenase can cleave the triple helix of collagen at about three-quarters of the length of the molecule from the X–Gly bond. Collagenases are widely distributed in vertebrate animals. In animal collagenases, the degradation of ‘native triple helical collagen’ or ‘water-insoluble native collagen’ significantly depends on the species of origin and the collagen type.^21,31,32,49 Collagenases of animal origin were frequently extracted and purified from fish processing byproducts or the viscera of fish^47,48 and other animals.^21,32,44 A review on collagenases from fish processing byproducts has been published.³²

2.3. Microbial collagenase

Microbial collagenases (EC 3.4.24.3) digest native collagen in the triple helical region at X–Gly bonds.^10,31,32 Microbial collagenases possess broad substrate specificities and degrade both ‘water-soluble/insoluble native’ and ‘denatured’ collagens in their triple helical conformation.^27,50 Microbial collagenases can degrade each polypeptide chain of collagen at multiple sites. Production of microbial collagenases has been reported in some pathogenic bacteria and fungi (Fig. 1). Bacterial sources have been widely used for the production of microbial collagenases.^{9,10,27,34,46,51,52} The first identified and characterized microbial collagenases were produced by Clostridium sp. The main source of our knowledge on microbial collagenases comes from extensive studies of C. histolyticum collagenases.^2,28,53–56 Seven different polypeptides with collagenolytic activity have been demonstrated in C. histolyticum and are divided into two groups on the basis of substrate specificity.⁵⁵

Apart from the well-studied collagenase-producing microorganism C. histolyticum, some other clostridia can also produce collagenase; however, little information can be found in the literature regarding their biochemical properties.⁵⁵ C. perfringens and C. tetani are the other collagenase-producing clostridia. They are extensively distributed in nature, particularly in soil and water contaminated with feces. They also live in the intestinal tracts of humans and animals.⁵⁵ Sometimes, C. perfringens causes infection, representing the most common pathogen of clostridial histotoxic infection.⁵⁵ It was reported that pathogenic strains such as C. histolyticum and C. tetani use collagenase to facilitate host invasion, colonization, and toxin diffusion during anaerobic infections.² Furthermore, another well-investigated bacterial collagenase is Vibrio alginolyticus collagenase. The collagenase activity of V. alginolyticus collagenase was found to be higher than that of any other bacterial collagenase.^71,72,75 In recent years, much emphasis has been shifted to microbial collagenases compared to animal collagenases.^{8–10,21,45,46} To date, microbial collagenases have been purified from a few species, and their genes have been cloned and sequenced.⁵⁷ However, many microbial collagenases have not yet been enzymatically and structurally characterized.⁵⁷

3. Commercial availability of microbial collagenase

The first commercially available microbial collagenase was isolated from C. histolyticum; for a long period, it was the only commercially available microbial collagenase. These clostridial collagenases were well studied and their properties were characterized. Currently, they are the only reference enzymes for comparison with newly discovered collagenolytic enzymes.²¹ These enzymes are important virulence factors in a variety of pathogenic microorganisms. Clostridial collagenases are large multi-modular zinc-metalloproteinases of approximately 115 kDa, consisting of four to six domains. Two to four accessory domains of approximately 10 kDa each form C-terminal collagen recruitment units of variable composition, providing important exosites for native collagenolysis, responsible for collagen binding and swelling.² Although most clostridial strains possess one collagenase, C. histolyticum encodes for two collagenases with complementary characteristics. Collagenase G (Col G) exhibits high collagenolytic and low peptidolytic activity; however, its homologue collagenase H (Col H) shows low collagenolytic and high peptidolytic activity.²

4. Isolation and screening of collagenase-producing microorganisms

Collagenase producing microorganisms have been found in diverse habitats such as soil,³³ thermal regions,^39–41 gut debris of earthworms,³⁴ food materials, fish sauces,⁴³ fish byproducts/waste (skin, bone, and fins) material,³² soil and sewage samples from slaughterhouses, various pathological sources, and leather houses/industries.^21,44

Several methods have been reported by various research groups to screen collagenase-producing microorganisms. However, a vast scientific controversy is found in the literature regarding the appropriate and well-defined screening of collagenase-producing microorganisms. This controversy is due to the existence of similar enzymes, such as collagenolytic protease, gelatinase and other proteases.^{29,34,45,46,51,52,54,58} There is a lack of a well-defined standard method for the isolation and screening of collagenase-producing microorganisms on the basis of qualitative and quantitative properties. Thus, more research should be focused on the standardization of a rapid, authentic and cost effective qualitative screening of collagenase-producing microorganisms. Here, we discuss a number of methods reported by different research teams for the screening of collagenase-producing microorganisms.

4.1. Approach based on gelatin hydrolysis

Gelatin is a class of protein fractions that do not exist in nature but are derived from the parent protein collagen by denaturation. Gelatin hydrolysis can enable the qualitative determination of collagenase activity. Several researchers have reported the use of 1 to 3% gelatin as a supplement in different medium compositions, such as trypticase soy agar, selective medium, potato dextrose agar, and nutrient agar, to screen extracellular collagenase-producing microorganisms.^9,10,45,46 Collagenase production is evident as a clear or clear halo zone around the colonies.^{29,34,44–46,51,52,58} However, the clarity of the hydrolyzed zone may not be very sharp on the media plate. Therefore, the clarity of the hydrolyzed zone around the microbial colony can be improved by precipitating the proteins with trichloroacetic acid (TCA).⁵⁹ It was reported that application of 30% TCA produced a very sharp and clear zone of gelatin hydrolysis.⁵⁹ An agar cup method was also reported to screen collagenase-producing microorganisms.⁴⁶ Cell-free supernatant was placed into cup-plates (8 mm) containing 1 to 2% (w/v) gelatin. The ability to digest gelatin is demonstrated by the formation of transparent halos on the cup-plates and is expressed in mm.⁴⁶

4.2. Approach based on hydrolysis of collagen and collagen hydrolysate

The collagenolytic activity of microorganisms can be detected using insoluble/soluble collagen or collagen hydrolysate products.⁶⁰ Most collagens have low denaturation temperatures (30 to 37 °C). To prevent collagen denaturation, collagen supplemented media cannot be sterilized by autoclaving. Therefore, the sterilization of growth medium supplemented with collagen is a challenging task for the scientific community.⁶⁰ The aseptic addition of collagen to sterilized growth medium may be problematic due to the non-sterility of the collagen. Furthermore, the acidic collagen solution (collagen is usually dissolved in dilute acid, e.g., 0.5 M acetic acid) may alter the pH of the medium. To overcome these problems, various other sterilization techniques have been used to disinfect and/or sterilize resuspended collagen, such as propylene oxide, ethylene oxide, gas plasma sterilization, gamma radiation, chloroform, glutaraldehyde, formaldehyde, acidic pH, electron beam sterilization, peracetic acid sterilization, or combinations of these.^61–63 These sterilization techniques do not adversely affect the structure and bio-tropic properties of the source material/collagen.^62,63

5. Measurement of collagenase activity

Microbial collagenase can act on both water-insoluble ‘native’ collagens and water-soluble denatured collagens (i.e., gelatin). Collagenase activity is influenced by various factors, such as solubility, viscosity, the concentration of the collagen solution, the type of acid used for dissolving the collagen, the amount of enzyme, collagen degradation products such as hydroxyproline, amino acids, peptides, and other reaction conditions (pH, temperature, and agitation). Water insoluble and soluble collagen are mostly used as substrates for collagenase assays.^40–42 In these methods, quantitative estimation of amino acids produced from the collagen substrates is commonly used to determine collagenase activity.^64,138 Rosen's modified colorimetric ninhydrin method for the quantitative estimation of amino acids is usually used to measure the collagenase activity.¹³⁸ In this method, a collagen substrate (usually ‘native’ collagen) is incubated with enzymes; collagenase activity is determined by measuring the free amino groups released from the collagen substrate. The collagenase activity is usually expressed as micro moles (μmol) of amino acids released per min per mL under standard assay conditions, using L-leucine/glycine as the reference standard. However, some research groups have also reported the determination of collagenase activity using gelatin as an assay substrate.^52,60

A modified ninhydrin-based method incorporating polyethylene glycol (PEG) for the assay of collagenase activity by spectrophotometry was demonstrated.⁶⁴ PEG is a series of polyether compounds that can stabilize Ruhemann's purple efficiently during reactions involving ninhydrin, and the coexistence of PEG makes ninhydrin-based reactions suitable for collagenase assays. The absorbance increases with increasing digestion time. Most collagenase assays are time-consuming (3 to 18 h).⁶⁴ If the collagenolytic assay is conducted at temperatures higher than 37 °C (above the denaturation temperature of collagen), the gelatinolytic activity is most likely being measured instead of the collagenolytic activity.²¹ At high temperature, collagen is hydrolyzed and collagen hydrolysate or gelatin is certainly obtained. Therefore, various synthetic peptides such as 4-phenylazobenzyloxycarbonyl-L-Pro-L-Leu-Gly-L-Pro-D-Arg (Pz-peptide), N-[3-(2-furylacryloyl)]-L-leucyl-glycyl-L-prolyl-L-alanine (FALGPA), and azo dye-impregnated collagen were developed as substrates for efficient quantification of collagenase activity.^{8,9,41,42,45,65}

In the azocoll assay, CaCl₂ is used as a cofactor, and the reaction time is 3 to 5 h at a maximum temperature of about 37 °C. In this assay, one collagenase activity unit (U) is defined as the amount of enzyme per mL that produces an increase in the optical density of 0.1 at 520 nm, due to the formation of azo dye linked soluble peptides.^9,10 Radiolabeling with ¹⁴C, ²H, ³H or fluorescence-labeled collagens was also developed as a substrate for quantitative estimation of collagenase activity.⁶⁴ These methods are considered to be more convenient and sensitive assays for collagenase activity. However, these methods require expensive devices and lead to the generation of radioactive waste. Fluorescent collagens and synthetic peptides seem more attractive as substrates; however, labeling with fluorescein isothiocyanate (FITC) or biotin is a time-consuming and expensive process.⁶⁴

A sensitive, fast and specific electrophoresis method for the determination of collagenase activity is also described.⁶⁷ In this zymography method, collagen is used as the substrate. Collagen is incorporated in the polyacrylamide gel and non-denaturing electrophoresis is carried out. This method demonstrates the hydrolysis of collagen by the enzyme, which is detected as clear zones/bands surrounded by the dark background of the stained protein (collagen). This method can detect collagenase activity up to nanogram levels. The collagenase activity band can be visualized by staining the gel with Coomassie blue dye. Although a number of collagenase assays exist, the need for a simple, reliable and economical assay to determine collagenase activity is, surprisingly, still unaddressed.

6. Classification of microbial collagenases

True microbial collagenases, especially bacterial collagenases, are described as enzymes that cleave the helical regions of ‘native’ fibrillar collagen under physiological conditions.^21,41,42,70 However, it is frequently difficult to distinguish true microbial collagenases from gelatinases and other bacterial proteases.²¹ The terms collagenolytic protease, gelatinase, collagenase, and collagenolytic enzymes have caused considerable mystification among the scientific community. This leads to controversy and inaccuracy in both the classification and nomenclature of microbial collagenases. To avoid confusion regarding their use, these terms should be defined.^34,68 Moreover, the frontiers should be very clear for the differentiation of vertebrate collagenases and gelatinases from microbial collagenases.

Animal collagenases split collagen by hydrolyzing a single peptide bond across the three α-chains organized in its native triple-helical conformation. After this primary fragmentation of collagen, attack on the α-chains is very limited, and the resulting fragments tend to uncoil into collagen polypeptides, which are more susceptible to other proteases, such as gelatinases.^21,35 Contrastingly, almost all collagen types are likely to be attacked by microbial collagenases at various specific sites of the α-chains.^21,50 It is essential to stress that a large number of microbial proteases have the capacity to hydrolyze single-stranded and denatured collagen polypeptides. These collagenolytic bacterial proteases should not be confused with the true microbial collagenases, which can degrade the triple-helical native collagen.²¹

Matrix metalloproteinases (MMPs) are a family of zinc-dependent endopeptidases (including interstitial collagenases, stromelysins, gelatinases, and membrane-type metalloproteinases) that possess collagenolytic activity.^21,35,72 Members of the family of MMPs are key enzymes in normal and pathological tissue remodelling. In MEROPS, the peptidase database, all MMPs belong to the M10 family; however, microbial collagenase belongs to the M9 family, which has entirely different biochemical and structural properties.⁷² Presently, MMPs belong to the sub clan MA (M) of the zinc metallopeptidase (M10 family). Previously, MMPs were included in clan MB, which contains only HEXXH metallopeptidases.²¹ In 1999, when the three dimensional structures became available, the clan MB was merged with clan MA in the MEROPS peptidase database. Presently, MMPs (M10 family) have a longer zinc binding consensus sequence (HEXXHXXGXXH) compared to the microbial collagenase (M9 family) zinc binding consensus sequence (HEXXH).^21,71–73 Details of the MMPs are presented in Table 1.

Table 1 Details of matrix metalloproteinases (MMPs) with MEROPS peptidase database IDs (source: Duarte;²¹ Rawlings^71–73)^a

MMP	Other name	EC number	Substrate	MEROPS ID
a NA = not assigned.
Collagenases
MMP-1	Collagenase 1, interstitial collagenase, matrix metalloproteinase 1, vertebrate collagenase, fibroblast collagenase	3.4.24.7	Type I, II, III, VII, VIII, IX and X fibrillar collagens, gelatin, aggrecan, fibronectin	M10.001
MMP-8	Collagenase-2, neutrophil collagenase	3.4.24.34		M10.002
MMP-13	Collagenase-3, AgMMP 3, rat collagenase	NA		M10.013
MMP-18	Xenopus collagenase, collagenase-4, xCol4	NA		M10.018
Gelatinases
MMP-2	Gelatinase A, 72 kDa gelatinase, type IV collagenase, 3/4 collagenase, matrix metalloproteinase 5, tissue gelatinase	3.4.24.24	Nonfibrillar collagens, gelatin, type IV, V, VII, X, XI and XIV collagen, elastin, aggrecan, fibronectin, galectin-3	M10.003
MMP-9	Gelatinase B, type IV collagenase, type V collagenase, 92 kDa gelatinase, macrophage gelatinase, neutrophil gelatinase	3.4.24.35		M10.004
Stromelysins
MMP-3	Stromelysin-1, transin, proteoglycanase, collagenase activating protein, matrix metalloproteinase 6, procollagenase activator	3.4.24.17	Proteoglycans, collagen telopeptides, fibronectin, laminin, casein, gelatin, type III, IV, V, VII, IX, X and XI collagens collagen, elastin, fibrinogen, laminin, elastin	M10.005
MMP-10	Stromelysin-2, transin-2	3.4.24.22		M10.006
MMP-11	Stromelysin-3	NA		M10.007
MMP-12	Metalloelastase, macrophage elastase	3.4.24.65		M10.009
Matrilysins
MMP-7	Matrilysin-1, putative metalloprotease, putative metalloproteinase-1 (PUMP-1), uterine metalloendopeptidase	3.4.24.23	Fibronectin, laminin, type IV collagen, proteoglycans, gelatin, elastin, fibrinogen, laminin, aggrecan, pro-MMP-1, pro-MMP-2, pro-MMP-7, pro-MMP-8, and pro-MMP-9	M10.008
MMP-26	Matrilysin-2, endometase	NA		M10.029
Membrane-type
MMP-14	MT1-MMP, MT-MMP-1, membrane-type matrix metalloproteinase 1, matrix metalloproteinase, membrane-type 1	3.4.24.80	Large tenascin-C, fibronectin, laminin	M10.014
MMP-15	MT2-MMP, MT-MMP-2, membrane-type matrix metalloproteinase 2, matrix metalloproteinase, membrane-type 2, SMCP-2	NA		M10.015
MMP-16	MT3-MMP, MT-MMP-3, membrane-type matrix metalloproteinase 3, matrix metalloproteinase, membrane-type 3, ovary metalloproteinase, membrane-type matrix metalloproteinase (Gallus domesticus)	NA		M10.016
MMP-17	MT4-MMP, MT-MMP-4, membrane-type matrix metalloproteinase 4, matrix metalloproteinase, membrane-type 4	NA		M10.017
MMP-24	MT5-MMP, MT-MMP-5, membrane-type matrix metalloproteinase 5	NA		M10.023
MMP-25	MT6-MMP, leukolysin, MT-MMP-6, membrane-type matrix metalloproteinase 6, MT-6 MMP	NA		M10.024
Other MMPs
MMP-19	Matrix metalloproteinase 19, RASI-1, RASI-6	NA	Type IV, V collagens, fibrin, elastin, gelatin, fibronectin, casein, laminin, aggrecan, amelogenin, fibrinogen, α1-antitrypsin	M10.021
MMP-20	Enamelysin	NA		M10.019
MMP-21	XMMP (Xenopus)	NA		M10.026
MMP-23	Cysteine-array matrix matalloproteinase, CA-MMP, femalysin	NA		M10.022
MMP-27	CMMP (Gallus domesticus), matrix metalloproteinase 22, MMP-22 (Gallus domesticus), MMP-27 (Homo sapiens), matrix metallopeptidase 27	NA		M10.027
MMP-28	Epilysin	NA		M10.030
—	Microbial collagenase	3.4.24.3	Native collagen	M09.001
				M09.002
				M09.003
				M09.004

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In some cases, the collagenase name was given to different enzymes, even though those enzymes were from different families.^21,85 Geobacillus collagenovorans MO-1, a thermophilic and collagen-degrading species, was claimed to produce an array of collagenolytic enzymes.^21,84 This strain produces three enzymes that are associated with collagen breakdown. A collagenolytic protease produced by this strain was able to hydrolyze collagen and two Pz-peptidases (metallopeptidases from peptidase family M3). It can also hydrolyze 4-phenylazobenzyloxycarbonyl-Pro-Leu-Gly-Pro-D-Arg (a synthetic peptide). This collagenolytic protease is a serine protease that belongs to the peptidase family; the two metallopeptidases are Pz-peptidases, which only act on small oligopeptides, not on fibrillar collagens. The term collagenase is indiscriminately assigned to both metallopeptidases and serine proteases. In recent years, remarkable progress has been made to identify new microbial collagenases and their collagen degradation mechanisms. Hence, before further discussing microbial collagenases, it is very necessary to discuss the structure and classification of microbial collagenases to prevent confusion. A strict standard of collagenase classification should be followed in publications to prevent controversy among the scientific community.⁶⁹ However, it is also suggested that collagenolytic bacterial proteases can include all proteases that are able to degrade at least one type of collagen.⁶⁹ The classification of microbial collagenases is described in Table 2 and Fig. 2.

Table 2 Characteristics of representative microbial collagenases according to the MEROPS peptidase database⁷²

Peptidases and homologues	MEROPS ID
(A) Peptidase family M9A
Bacterial collagenase V	M09.001
VMC peptidase	M09.004
Subfamily M9A non-peptidase homologues	Non-peptidase homologue
Subfamily M9A unassigned peptidases	Unassigned
(B) Peptidase family M9B
Bacterial collagenase G/A	M09.002
Bacterial collagenase H	M09.003
Subfamily M9B non-peptidase homologues	Non-peptidase homologue
Subfamily M9B unassigned peptidases	Unassigned

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Fig. 2 Schematic classification and domain organization of microbial collagenases. The domain structures of collagenase from Vibrio parahaemolyticus (a); V. mimicus (b); V. alginolyticus (c); Col G from Clostridium histolyticum (d) and Col H from C. histolyticum (e). AD = activator domain; CD = catalytic domain (peptidase domain); PKD = polycystic kidney disease-like domain; CBD = collagen binding domain; PPC = pre-peptidase C terminal domain.

6.1. Metallocollagenases

Metallocollagenases are zinc-containing enzymes that usually require calcium ions for optimum activity and stability. These metallocollagenases are found mostly in vertebrates and bacteria. C. histolyticum and Vibrio sp. metallocollagenases have been widely studied.^71–73 The Nomenclature Committee of the International Union of Biochemistry and Molecular Biology (NC-IUBMB) assigned these microbial collagenases (endopeptidases) to the EC 3.4.24.3 group of the metalloendopeptidase sub-class (EC 3.4.24).²¹ These microbial collagenases (Clostridium and Vibrio collagenase) belong to the MEROPS peptidase family M9.⁷² The MEROPS M9 peptidase family is divided into subfamilies, M9A and M9B.⁷²

6.1.1. M9A family: Vibrio collagenases. The collagenases from Vibrio are the most widely studied after Clostridium collagenase. The subfamily M9A has four related groups of peptidases whose holotypes are bacterial collagenase; Vibrio VMC peptidase (from V. alginolyticus and V. mimicus); non-peptidase homologues; and unassigned peptidases (Fig. 2, respectively).^{21,71,72,74,75} In the first step of collagen degradation, Vibrio collagenase cleaves the triple-helical collagen at a point three-quarter of the way from the N-terminus by preferentially hydrolyzing the peptide bond X–Gly.²⁶ Vibrio collagenase cleaves native collagen to a higher degree than vertebrate metalloproteinase. Vibrio collagenase hydrolyzes the Pz-peptide (Pz-Pro-Leu-Gly-Pro-D-Arg) by cleaving the peptide bond Leu–Gly, which is also degraded by Clostridium collagenase.

According to substrate specificity and structural similarity to other zinc proteases, the metalloproteinases from Vibrio were divided into two classes: class I (cannot digest collagens) and II (can hydrolyze collagens).^21,76 Based on their amino acid sequences, enzyme molecular masses, substrate specificities, and function, these two classes of Vibrio extracellular metalloproteinases were re-organized into three categories.⁷⁶ The molecular weight of class I enzymes is approximately 36 to 38 kDa, and they cannot digest collagen. The class I enzymes are members of the M4 family (thermolysin family). Class II enzymes are about 62 to 71 kDa in molecular mass. These enzymes cannot hydrolyze casein but can digest collagen. Class III has ∼89 kDa molecular weight. These enzymes can digest caseins, gelatin, and collagen.⁷⁶ Class II and Class III enzymes are members of the subfamily M9A in the MEROPS database. However, Class II and Class III enzymes have significant differences in their structure and function. Especially, class II enzymes have a zinc-binding motif and contain no C-terminal domain. Meanwhile, the class III proteases have a zinc-binding motif, contain polycystic kidney disease (PKD) like domains and have a pre-peptidase C-terminal (PPC) domain at their carboxyl terminus.^21,69,76,77 Although class II enzymes do not contain a C-terminal extension, it is reported that the motif in the carboxyl terminus of V. mimicus collagenase is involved in binding to collagen.⁷⁵ In contrast, neither the PPC domain nor the PKD-like domain of class III enzymes has yet been demonstrated to function as a collagen binding domain.⁶⁹ Only Vibrio metalloproteinases from class III (MEROPS M09.001) and class II (MEROPS M09.004) should be considered as collagenases.²¹ To best of our knowledge, the crystal structure of Vibrio collagenase or its domain has not been resolved. Therefore, it is still unclear how Vibrio collagenase recognizes and cleaves native collagen.

6.1.2. M9B family: Clostridium collagenases. Clostridium collagenase was the first bacterial collagenase discovered and reported by Maschmann in 1937. Followed by C. histolyticum, collagenases have been extensively studied by various research groups in basic science laboratories for medical treatment and other purposes. M9B subfamily Clostridium collagenases have been further classified into two well-recognized classes, viz. Class I and class II collagenases. The class I and II collagenases were differentiated on the basis of their kinetic and structural characteristics, which includes their relative activities towards different substrates (collagen, gelatin, and synthetic substrates), freeze–thawing stability, and primary, secondary and tertiary structures.²¹ The class I microbial collagenases are highly active on native collagen and moderately active on synthetic collagen peptides. However, class II collagenase shows moderate activity towards native collagen and high activity towards synthetic peptides. Matsushita's research group and his collaborators showed the presence of two collagenase genes, colG, and colH, in the C. histolyticum chromosome. Class I collagenases are encoded by the colG gene, and class II collagenases are encoded by the colH gene.^{55,69,78–80} In addition to colG and colH, collagenase genes from other Clostridium species are also reported, such as colA from C. perfringens and colT from C. tetani. In the MEROPS database, all Clostridium collagenases are members of the M9 family.^69,71,72 More investigations on microbial collagenases and their encoding genes are necessary to draw a conclusive evolutionary picture for these enzymes.

A Clostridium collagenase molecule (unit) is composed of the N-terminal activator (collagenase) domain and the C-terminal recruitment (peptidase) domains, which contain a conserved zinc-binding motif and function as catalytic domains. The recruitment domains usually contain one or two collagen binding domains (CBDs) and one or two polycystic kidney disease (PKD)-like domain(s). The recruitment domains are not required for the degradation of triple helical collagen, but are most likely needed for larger collagen entities such as fibrils. Mostly, the activator domain and catalytic domain in the collagenase module remain closed during native collagen cleavage but relax to the open ground state once the native collagen is degraded.³⁸ The Clostridium collagenases hydrolyze native collagens into a mixture of smaller fragments or peptides (Fig. 3). The distinct hyper-reactive sites for class I and II Clostridium collagenases are Y–Gly bonds in the repeating Gly–X–Y collagen sequence. The two class of collagenases bind to different portions of collagen and have different specificities to cut native collagens, i.e. they show synergy in collagen degradation.^{69,80,81,83,84}

Fig. 3 A schematic model for the degradation of native collagen by a microbial collagenase.

6.2. Serine collagenolytic proteases and serine collagenases

Several S8 proteases with collagenolytic activity have been reported from bacteria, archaea, crab, crustaceans, fish and human pathogens.^36,69,88,89 Some S8 proteases have been reported in recent years to be collagenolytic proteases. Proteases of this family (S8 family) are characterized by an Asp/His/Ser catalytic triad and an alpha/beta fold catalytic center containing a seven-stranded parallel beta-sheet.⁶⁹ Most proteases in the S8 family have no collagenolytic activity. Serine collagenolytic proteases are sometimes referred to as serine collagenases. The serine collagenases reported to date possess broad proteolytic activities.^{21,69,71–73} The thermostable serine collagenolytic protease from Geobacillus collagenovorans MO-1 is the first reported S8 collagenolytic protease. This enzyme has a C-terminal collagen-binding domain, and its cleavage sites on collagen are various but specific.^86,87 Recently, two S8 proteases from marine bacteria, Pseudoalteromonas sp., were characterized as collagenolytic proteases, and their actions toward collagen degradation were also revealed.^69,88–93 However, there is no evidence that serine proteases can hydrolyze triple-helical native collagen. Therefore, it is suggested that these peptidases could be considered to be serine gelatinases, not serine collagenases.²¹

7. Fermentative production of collagenase

A limited number of reports are available on the production of microbial collagenases. Most of the microbial collagenases reported to date have been produced by the submerged fermentation (SmF) process.^8–10,45,46 Various types of collagen supplements (native collagen, gelatin/collagen hydrolysate) have been used in culture media for the production of extracellular collagenase enzymes. Various researchers have used malt extract with 1% gelatin and soybean flour as a supplement for culture media, which can induce collagenase enzyme production.^8–10,46,58 Substrates such as insoluble collagen^34,51 and fish collagen²⁷ have also been reported as inducers in collagenase production. Crude substrates such as fish scale powder and fish skin, as well as mammalian, shrimp and crab by-products, are also used as sole carbon/nitrogen sources in culture media for collagenase production by some research groups.^27,29,34,51 A maximum collagenase activity of 240 U per mL from Nocardiopsis dassonvillei in SmF using shrimp and crab by-products as the sole nitrogen and carbon source has been reported.²⁹ The extracellular collagenase produced by Bacillus pumilus showed a maximum 129.5 U of collagenase activity.⁴⁴ Collagenase production from B. cereus (23.07 U per mL) and Klebsiella pneumoniae (10.84 U per mL) has also been reported.⁵² Tran and Nagano reported a maximum activity of 3.07 U per mL for a collagenase from B. subtilis.⁶⁰ A maximum collagenolytic activity of 6.8 ± 0.4 U was reported for Candida albicans collagenase.⁴⁶ The potential uses of microbial collagenases and their high demand is attracting the interest of the scientific community to find new microbial species/strains that are able to produce high titers of extracellular collagenases with novel properties.

8. Strategies used to improve microbial collagenase production

In the last decade, collagenase research has gained momentum because collagenases have therapeutic, industrial and biotechnological applications in addition to those of conventional proteases.^94,95 The development of low-cost industrial media formulations for the production of microbial collagenases can play a significant role in the cost, concentration, yield, and productivity of the products. Media composition is one of the most significant factors in the industrial production of collagenase because 30 to 40% of the production costs are due to the high cost of the growth media.^9,10 Optimizing the cost/benefit ratio of such processes is, therefore, a leading concern. The cost of collagenase production is a major hurdle in the successful industrial application of collagenases. An inexpensive medium can be used for economically feasible industrial collagenase production with higher yield. The optimization of physiochemical conditions (pH, temperature, carbon source, and gelatin) for collagenase production from B. cereus CNA1 and K. pneumoniae CNL3 has been reported.⁵²

8.1. Statistical design methods

To meet commercial requirements, the optimization of different process parameters and media compositions are well-known methods for the overproduction of enzymes in large quantities.^94–98 Culture media formulations and physicochemical bioprocess conditions are reported as significant factors to enhance microbial collagenase production.⁹⁶ However, designing a medium is a laborious, expensive and often time-consuming process involving several experiments. Statistical tools such as response surface methodology (RSM) play an important role to optimize the level of various bioprocess variables for enhanced production of extracellular microbial collagenases.^9,10,94,95 Statistical optimizations of microbial collagenase production by the SmF process has been reported.^9,10,46 Using factorial design and RSM approaches, the optimization of culture conditions (pH, time, temperature, inoculum size, orbital agitation speed and substrate concentration) for extracellular collagenase production from C. albicans URM3622 was studied using 2^6-2 factionary factorial and 2³ full factorial designs.⁴⁶ A full two-level factorial design with three variables (medium pH, temperature and soybean flour concentration) followed by a central composite design (CCD) was reported for the optimization of collagenase production from Penicillium aurantiogriseum URM4622.^9,10 They observed a 7.06 U improvement in collagenolytic activity from C. albicans URM3622 (ref. 46) and a 5-fold increase in activity from P. aurantiogriseum URM4622 (ref. 9 and 10) after statistical experimental designs. However, an improvement in microbial collagenase production requires a comprehensive understanding of microbial strain, metabolic functions through metabolic model construction, and subsequent in silico experimentation using systems biology methods.¹³² These in silico experiments can suggest cell manipulations that can also be applied using in vitro synthetic biology techniques, leading to increased microbial collagenase production.¹³²

9. Gene cloning of collagenase

Microbial collagenases have been purified from a limited number of bacterial species, and their corresponding genes have been cloned and sequenced.^{24,38,50,55–57,65,83,102–104} The expression of Burkholderia pseudomallei putative collagenase in Escherichia coli was reported.¹⁰⁵ The collagenase gene was cloned, and the protein was expressed as a glutathione S-transferase (GST) fusion and purified. However, there is a lack of information about the domain composition and features of B. pseudomallei collagenase. Collagenase activity with FALGPA peptide was weak; however, this could be related to the glutathione S-transferase fusion.²¹ High gelatinase activity was detected, which led the authors to postulate that B. pseudomallei express an active collagenase/gelatinase; however, no class assignment was possible.¹⁰⁵ The catalytic domain of collagenase G from C. histolyticum has been cloned,³⁸ recombined and expressed in E. coli, and purified using the affinity and size-exclusion column chromatographic methods.⁸³ Ducka et al. established an E. coli expression system for a range of constructs of collagenase T from C. tetani and collagenase G and H from C. histolyticum.^54,101 The collagenase gene was cloned from Grimontia (Vibrio) hollisae, and its complete nucleotide sequence was determined.⁵⁷ The Brevibacillus expression system produced the recombinant collagenase. Collagenases, or genes encoding collagenases, have been identified in many Clostridium species: C. histolyticum, C. perfringens, C. botulinum, C. tetani and C. difficile.^21,55,69 The cloning of the collagenase G and H genes simplified the development of microbial collagenase expression systems. The Cl. perfringens expression system alleviated translational problems in expressing ColH.²⁸ Mutagenization and nucleotide sequence analysis of a cloned gene would provide insights into some structure–function relationships for the enzyme. Matsushita et al. cloned and sequenced the col gene encoding a 120 kDa collagenase from C. perfringens.^55,56 They also described the cloning and sequencing of the colH gene encoding 116 kDa collagenase from C. histolyticum. It is evident that a number of microbial collagenases are awaiting deep structure studies, classification and characterization.¹⁰⁵

10. Purification of microbial collagenases

The purification of microbial collagenase is complicated due to the presence of multiple forms and other proteases with similar physical and chemical characteristics.⁵⁵ The purified form of collagenase is required to study its biochemical properties, enzyme structure, catalytic mechanism, structure–function relationships and biotechnological and medical applications.⁹⁶ However, the commercial applications of collagenases in the food and leather industries do not require highly purified enzymes. Purification of microbial collagenase by various techniques resulted in the identification of two to seven different microbial collagenase fractions.^55,56 Microbial collagenases have been purified by some researchers.^{36,41–44,48,55,65,66,77,83,88,99,100} The purification of microbial collagenase from C. histolyticum,³⁸ B. pumilus,⁴⁴ B. licheniformis²⁷ and Rhizoctonia solani⁵¹ has been reported. Various procedures (ammonium sulfate precipitation, ultrafiltration, immobilized metal affinity chromatography, amylose affinity chromatography, gel filtration chromatography, ion exchange chromatography, size exclusion chromatography and N-terminal tag removal) have been applied for the purification of microbial collagenases.^{2,38,40,50,51,81–83} Although several research groups have performed extensive studies, there is no consensus on the number of enzymes and their properties. This may be due to the difference in procedures used for the enzyme preparation and strains used for collagenase production. However, most reports on microbial collagenase purification schemes were carried out on a small/laboratory scale. Furthermore, excessive purification is very expensive and also reduces the overall recovery of the microbial collagenase. 101-fold purification of a thermostable alkaline collagenase from Thermoactinomyces sp. was reported.⁴⁰ The purified B. licheniformis and B. pumilus collagenases showed 26.3-fold and 31.53-fold increases in specific activity, respectively.^27,44 Purification of recombinant collagenase from Grimontia (Vibrio) hollisae,⁵⁷ C. histolyticum (ColH) 116 kDa collagenase,¹³³ and C. perfringens 120 kDa collagenase⁵⁵ have been reported.

11. Characterization of microbial collagenase

A limited number of purified microbial collagenases have been characterized by their activity and stability profiles corresponding to temperature, pH, and the effects of metal ions and chelating agents. The biochemical characteristics of microbial collagenases are given in Table 3. The properties of collagenases depend on the microbial species/strain. Most of these collagenases have a high apparent molecular mass, within the range of 50 to 120 kDa. Very few reports are available on the effect of pH on collagenase activity. The optimum pH of microbial collagenases has been found to be in the range of 5 to 9.5. The optimum temperature for the activity of microbial collagenases may be associated with the temperature required for the growth of the microorganism as well as the microbial species/strain,⁹⁶ and has been found to be in the range of 40 to 65 °C. Ions play a significant role to improve the activity of microbial collagenases. The structural stability and activity of Clostridium collagenases are enhanced in the presence of calcium ions.^28,101 However, collagenase activity is majorly inhibited by Fe²⁺. The effects of different metal ions on the activity of microbial collagenases are summarized in Table 4.

Table 3 The biochemical characteristics of microbial collagenases^a

Organism name	pH optima	Temp. optima (°C)	Molecular mass (kDa)	References
a ND = not determined.
Bacillus licheniformis F11.4	7	50	124	Ace Baehaki^27
Clostridium histolyticum	ND	ND	116	Jung,^133 Matsushita^134
C. perfringens	7.2	42	120	Matsushita^55
C. histolyticum	ND	ND	116	Yoshihara^56
Grimontia (Vibrio) hollisae 1706B	ND	ND	84	Teramura^57
B. subtilis FS-2	9	50	125	Nagano^43
Penicillium aurantiogriseum URM4622	8	45	39.16	Lima^45
Candida albicans URM3622	8.2	45	ND	Lima^46
B. cereus CNA1	7	45	ND	Suphatharaprateep^52
Klebsiella pneumoniae CNL3	6	40	ND
Thermoactinomyces sp. E-21	9.0–9.5	60–65	50	Petrova^40
Rhizoctonia solani	5	40	66	Hamdy^51
B. pumilus Col-J	7.5	45	58.64	Wu^44
Nocardiopsis dassonvillei NRC2aza	8	55	ND	Abdel-Fattah^29

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Table 4 Effects of different metal ions on the activity of microbial collagenases^a

Source of enzyme	Inhibition	Activation	Reference
a ND = not determined.
Clostridium histolyticum	ND	Ca^2+	Ohbayashi^28,101
C. histolyticum	ND	Zn^2+	Jung^133
Bacillus licheniformis F11.4	Fe^2+, Mg^2+, Mn^2+, Co^2+	Ca^2+, Cu^2+	Ace Baehaki^27
B. pumilus Col-J	Mn^2+, Pb^2+	Ca^2+, Mg^2+	Wu^44
Rhizoctonia solani	Fe^2+, Hg	Co^2+, Ca^2+, Cu^2+, Mg^2+, Zn^2+	Hamdy^51
Thermoactinomyces sp. E-21	Fe^2+, Cu^2+, Mn^2+, Sr^2+, Cd^2+, Ba^2+, Zn^2+, Ni^2+	Mg^2+, Ca^2+, Co^2+	Petrova^40
C. perfringens	ND	Zn^2+, Ca^2+, Mg^2+	Matsushita^55

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12. Applications of microbial collagenases

The applications of microbial collagenases are wide, including areas such as the food, tannery, meat, and cosmetic industries, the production of pharmaceutical compounds, and the bio-restoration of frescoes.^21,35,69 (Fig. 4). Microbial collagenases are also applied for the isolation and cultivation of mammalian cells. Therefore, they have a significant role in medical sectors. Microbial collagenases can be used to treat burns, wounds, scar tissue, transplantation of specific organs, Peyronie's disease, destructive fibrosis, liver cirrhosis and blood cleaning to improve screening in medical diagnostics.^21,32

Fig. 4 Applications of microbial collagenases in various sectors.

12.1. In food processing and allied industries

Microbial collagenase can be used in several food applications (Fig. 5). However, the commercial use of microbial collagenase in the food industry is limited to a few applications due to its higher cost.

Fig. 5 Possible applications of microbial collagenases in food and nutrition industries.

12.1.1. In meat tenderization. Tenderness is one of the most important sensory qualities of meat; it is influenced by the length of sarcomeres, the integrity of connective tissue (background toughness) and myofibrils (actomyosin toughness).^{106–108,113} Toughness is one of the most significant causes of unacceptability in meat quality.¹⁰⁶ In developing countries such as India, most animals are reared for dual purposes and slaughtered only after the end of their productive economic life (when they are useless for any other purpose). The meat obtained from these old/spent animals is very tough. Therefore, a tenderization process is required to increase the consumer acceptability of raw meat and meat products.

To enhance meat quality, the tenderization problem can be effectively overcome using enzymatic tools and process modifications.¹⁰⁹ Treatment by proteases (microbial and plant origin) is one of the most progressive methods used to increase meat tenderness. These proteases have broad specificities toward meat proteins. Therefore, microbial/plant protease might result in several undesirable sensory attributes in tenderized meat. Collagen (which accounts for 80% of the connective tissue) is also responsible for the toughness of red meats, and its digestion induces the meat tenderization process. Thus, a microbial collagenase that showed specific hydrolyzing activity towards collagen compared to other proteins would be more advantageous.¹¹¹ Therefore, microbial collagenases have been proposed to be an attractive alternative to non-specific plant/microbial proteases in meat tenderization processes.¹¹⁰ Collagenases from C. histolyticum and Vibrio B-30 have shown promising results in meat tenderizing studies.^110,112,127 However, concerns regarding safety issues such as pathogenicity and other unfavorable effects have limited the industrial use of microbial collagenases from these organisms in the process of meat tenderization. An ideal meat tenderizer should be a specific enzyme with high activity at room temperature and should be easily inactivated during the cooking process of meat/meat products.¹¹³ Microbial collagenase from non-pathogenic and safe microorganisms with improved specificity toward collagens could be used as an ideal meat tenderizer. Zhao et al. demonstrated that a cold-adapted collagenolytic enzyme from Pseudoalteromonas sp. SM9913 significantly reduced beef meat shear force and preserved the fresh color and moisture compared to commercially used papain and bromelain. The collagenolytic enzyme had a strong selectivity for the degradation of collagen/myofibril proteins and had a distinct tenderization mechanism.¹¹³

12.1.2. In collagen extraction. Collagen and its hydrolyzed form, gelatin, are widely used in food industries. Traditionally, collagen is extracted using an acidic solution with or without the aid of an enzyme.⁴ The yield of collagen extraction can be increased by using low concentrations of microbial collagenases.⁵² Microbial collagenases facilitate collagen extraction via the cleavage of the telopeptide region.⁵² A combination of collagenases from B. cereus and K. pneumoniae strains with acid treatment yielded a higher collagen recovery from salmon skin preparations than using an acid treatment alone.⁵² Therefore, microbial collagenase might be used for the extraction of collagen for industrial applications.

12.1.3. In the preparation of bioactive collagen hydrolysate and collagen peptides. In publications, “collagen hydrolysate” is defined by several terms, such as “collagen peptides”, “collagen degradation products”, “hydrolyzed collagen”, and “hydrolyzed gelatin”, to designate the same product.¹¹⁶ Collagen hydrolysate is usually prepared from gelatin using proteolytic treatment. Proteases such as trypsin, chymotrypsin, pepsin, alcalase, collagenase, bromelain and papain are the most regularly used enzymes for collagen hydrolysate preparation. The average molecular weight of commercially available collagen hydrolysate ranges between 0.5 to 20 kDa.^7,114–116 The structure, composition, molecular weight distribution and functional properties of collagen hydrolysate depend on the processing conditions and raw material as well as the specificity of the enzyme used to hydrolyze the gelatin. Collagen hydrolysate has been approved as Generally Recognized as Safe (GRAS) by the Center for Food Safety and Nutrition, US Food and Drug Administration (USFDA).⁷ From a nutritional point of view, collagen hydrolysates are well acknowledged for their safety.⁷ Therefore, collagen hydrolysates/peptides derived from animal/seafood based collagen are of great interest as potential ingredients in functional food and beverage products.⁷

Collagen hydrolysate (collagen-derived ingredients) is widely used in the food, cosmetic and pharmaceutical industries due to its gelling capacity and its texturizing, thickening, and water binding capacities, as well as its swelling and solubility properties, emulsion, foam formation and stabilization, adhesion and cohesion, protective colloid function and film forming capacity.^7,116 It is commonly used as a dietary supplement or included in various foodstuffs. The ingestion of collagen peptides has been shown to induce numerous biological processes.^114–119 Some clinical studies reported that daily oral intake of collagen hydrolysate decreased joint pain, reduced skin wrinkles and improved skin properties. Other studies have also suggested that a hydrolysate enriched diet can improve bone collagen metabolism.⁷ Collagen hydrolysate can be used as a relevant alternative in the designing of future nutritional approaches to managing osteoarthritis and osteoporosis prevention.¹¹⁶ Collagen hydrolysate/collagen peptides have exhibited strong antioxidant, anti-fatigue, antimicrobial and angiotensin-converting-enzyme (ACE) inhibitory activities.^8,128

Microbial collagenases are also used as hydrolyzing agents for the preparation of collagen hydrolysates/collagen peptides. Preparations of strong antimicrobial and radical scavenging properties containing bovine collagen hydrolysates were reported using P. aurantiogriseum URM 4622 collagenase.⁸ Ding et al.¹²⁸ demonstrated in vivo anti-fatigue activity and antioxidant activity of jellyfish collagen hydrolysate. Specific antifreeze peptides derived from shark skin collagen hydrolysate have been reported.¹³¹ Guo et al.¹²⁹ isolated Alaska Pollock skin collagen-derived mineral chelating peptides that may have potential applications as a functional food ingredient in the management of mineral deficiencies. Squid skin collagen hydrolysates have anti-hyaluronidase, antityrosinase, and antioxidant activites.¹³⁰

12.1.4. In meat and seafood processing byproducts valorization. Microbial collagenase may also be used to recover biomolecules/biopolymers such as collagen, gelatin, collagen peptides, collagen hydrolysate, and protein hydrolysates from different animal/seafood processing by-products.¹³⁹ The cost-effective production of collagenase from B. tequilensis was achieved by utilizing meat industry wastes as the sole nitrogen and carbon source. However, a decrease in extracellular collagenase production from B. tequilensis can be negotiated by the utilization of waste animal skin for useful product synthesis and waste management.³⁴

12.2. In leather processing

Due to water pollution, careless solid waste disposal and gaseous emissions, leather industries are categorized as a red industry and are under deep pressure to develop environmentally efficient leather-making processes to meet the terms of modern pollution and discharge legislation. Conventional leather processing involves the use of a large number of different chemicals, such as lime.¹³⁵ Due to global concerns about the environmental impact of the leather industry have led tanners to reduce the elements of toxicity and the organic and inorganic chemicals in effluents.¹³⁶ Therefore, it is a great challenge today to find environmentally friendly alternatives to the chemical processing of hides and leather. In this respect, the development of enzymatic processes as alternatives to some chemical-based processes is being encouraged because they not only yield products with improved quality but also reduce the use of hazardous and polluting chemicals.^135–137

Collagenases also have potential applications in the leather industry.^21,32 They are used as biocatalysts to improve the dye exhaustion process. Dyeing is an important process of the leather industry. Synthetic colorants that are used in leather dyeing are also a major source of environmental pollution. The unexhausted dyes present in leather industry effluents are frequently resistant to bioremediation, which is a major concern.¹²⁰ After leather tanning, the use of bacterial collagenases results in the opening-up of the fibrous leather network. In the leather matrix, the diffusion of dyes can be enhanced by opening up this fibrous collagen network. This process results in an uptake of dye of up to 99% using this eco-friendly approach and also improves bulk properties, such as the softness, fullness, grain smoothness, feel and general appearance of the leather.¹²¹

Dehairing is the first step in the leather making process. In the leather industry, the dehairing process creates a high level of environmental pollution. Therefore, the enzymatic dehairing process may help to reduce pollution load and collagen damage. A high degree of control over the process is the major limitation of the enzymatic dehairing process.¹²² The use of collagenase enzymatic formulations has been proposed for the dehairing process.¹²³ However, the efficacy of collagenases and their cost are restricting their commercial applications in dehairing processes.¹²³

12.3. In the laboratory as an experimental reagent

In the laboratory, microbial collagenases are widely used as experimental reagents in laboratory-scale studies such isolation of rat liver cells and scission of collagen-like peptide infusion proteins.^35,58 Cell culture has been widely used as an essential tool in biotechnology, molecular biology, and toxicology to address important economic, technical and scientific issues in biology. There are various types of cultured cells, such as primary cultures, cell lines, and cell strains, in which different types of enzymes are frequently used for tissue disaggregation. Microbial collagenase formulation has been successfully used for the isolation of cells from bone, endothelial cells, and neuronal cells as well as isolation of the islets of Langerhans, among other applications.²¹ Fibrous tissues with high collagen content are particularly resistant to trypsin digestion. Microbial collagenases are especially valuable in the case of fibrous or sensitive tissues in which trypsin use would be ineffective or damaging.²⁵ Furthermore, the Clostridial collagenases, in association with other enzymes, have been increasingly applied for numerous medical purposes. For example, the creation of autologous dermal tissue from fibroblasts was isolated using Clostridium collagenase.^21,25

12.4. In human health and nutrition

One of the most significant applications of microbial collagenase is found in medical industries. Collagen constitutes one-third of the body protein in humans, reflecting its extraordinary role in health and disease.^4,38 Microbial collagenases have directly been employed in clinical therapy (debridement of burns, wound healing, sciatica, retained placenta, lumbar disc herniation, chronic total occlusion treatment, and enhancement of adenovirus-mediated cancer genetic therapy or electro-genetic therapy).^9,21 The use of Clostridium collagenase is considered as a capital addition in wound debridement, helping to avoid surgery complications and also limiting the progress and enlargement of necrotic tissue.^21,124,125 Clostridium collagenase is also applied for treatment of Dupuytren's disease (DD).²¹ Enzymatic fasciotomy with Clostridial collagenase targets excessive collagen deposition and ruptures the fibrous cords that cause contractures in DD. Historically, treatment options for Dupuytren's contracture were limited to surgical procedures. The FDA has approved Clostridium collagenase for DD treatment.²¹ Microbial collagenases are used for various types of destructive fibrosis. A recent application of microbial collagenases is related to genetic therapy. The administration of naked nucleic acids into cells and tissues is considered to be the simplest and safest method for gene delivery.²¹ The application of microbial collagenase (with hyaluronidase) increases transfection efficiency in tumors without promoting the spreading of metastases.¹²⁶

12.5. In environmental protection

In addition to the pathogenic microorganisms, various collagenase-producing microorganisms have been isolated from marine and terrestrial sources. Although a huge quantity of collagen is produced annually in the aquatic/marine biosphere alone, there is no substantial accumulation of collagen in ocean sediments because they are biodegraded by the naturally occurring bioconversion process mediated by collagenolytic marine microorganisms.⁶⁹ Zhang et al. revealed that the mechanisms of environmental microorganisms on collagen degradation are helpful for the study of the global nitrogen cycle.⁶⁹ Furthermore, the role of Clostridium collagenases in the putrefaction process of the carcass is well documented.

13. Challenges associated with microbial collagenases in the food and nutrition industries

The market share of microbial collagenases is considered to be higher compared to the proportion of animal and plant collagenases. The properties of microbial collagenases (Clostridium collagenases) are well documented, and these collagenases are widely used in medical sectors. However, studies of collagenase-producing microorganisms and their importance in food and nutrition are rare. Major challenges and limiting factors for microbial collagenase applications in the food and nutrition sectors are discussed.

13.1. Sources of collagenase-producing microorganisms

The ecosystem contains a vast abundance of microorganisms. The microorganisms involved in the degradation of animal waste/by-products may be capable of producing extracellular collagenase. Marine microorganisms have recently emerged as excellent sources of collagenase. Most of the commercially available and well-studied collagenase-producing microorganisms are anaerobic and pathogenic. The collagenase produced from pathogenic microorganisms creates specific concerns about the pathogenicity of the enzyme. Presently, very few manufacturers are involved in the production of microbial collagenases (Clostridium collagenases) globally. Due to the higher cost and safety concerns of collagenases, their potential applications in the food and nutrition sectors are scanty. Therefore, in view of food and nutrition applications, there is a vast requirement to search non-pathogenic, extracellular collagenase-producing microorganisms.

13.2. Screening, optimization and production of collagenase

Numerous controversies can be found for the screening of collagenase-producing microorganisms in the scientific literature, due to the low denaturation temperature (∼30 to 37 °C) and acidic solubility of collagen. Furthermore, the search for a simple, cost-effective and high throughput method for the determination of collagenase activity is still underway. However, radiolabeled and synthetic peptide methods used to determine collagenase activity are very laborious, time-consuming and expensive. Cost-effective collagenase production is a critical challenge to the scientific community. Sustainable collagenase production for the food and nutrition sectors is becoming a priority due to the current dependency upon Clostridium collagenase. The use of cheaper agro-industrial by-products as fermentation media can also help to reduce the enzyme cost. The optimization of processes parameters (medium composition nutrients and culture conditions) for collagenase production is very rare. It is a critical challenge to define the level of each variable (physico-chemical, culture conditions, and medium composition), and their possible interactions between the variables. Collagenases producing microbial strains can be typically designed using combinations of gene manipulations (addition and removal of genes). This process may afford the potential for the successful production of microbial collagenases. The challenges to improving the production of collagenase may also be overcome using synthetic and systems biology.

13.3. Classification, crystal structure and collagen degradation mechanism of microbial collagenase

A new overview to classify and purify the microbial collagenases is needed for the identification of true microbial collagenase from other, similar collagenolytic proteases. Still, there are no specific recommendations for the purification and classification of microbial collagenases. The mechanism of microbial collagenase for the degradation of native collagen is still unclear. To best of our knowledge, no crystal structure of a microbial collagenase has been reported, except Clostridium collagenase. Hence, microbial collagenase classification, purification, crystal structure and collagen degradation mechanisms are very challenging tasks for the scientific community. Microbial collagenases can also be used for the production of novel true collagen peptides (native collagen peptides). These collagen peptides have been reported to have health promoting activities. Currently, the development of true/native collagen peptides and their potential applications as functional food ingredients may create opportunities for their diverse applications in the food and nutrition industries.

14. Directions and recommendations for future research

Potential application studies of microbial collagenases are not common and evolving in the direction of food, nutrition, and other related industrial applications. Currently, very few studies on the fermentative production and application of microbial collagenases have been reported.^8–10,45,46 There is a huge controversy regarding the proper and unambiguous identification and classification of collagenase-producing microorganisms. To reduce the cost of collagenases, we have to look for non-pathogenic microbial sources for collagenase production using cheap agroindustrial by-products. The production of collagenase from non-pathogenic strains can increase food industry applications by preventing pathogenicity and addressing safety concerns. Most of the reported microbial collagenases have not been structurally or enzymatically characterized. Therefore, the exploration of the specificity, structure, and functions of microbial collagenases could provide further understanding of collagen degradation mechanisms and also uncover various potential biotechnological applications.

It would be interesting to study the role of microbial collagenases to improve the quality of food, meat, and other related value-added products. Future research and studies should be focused on multidisciplinary areas such as the production of bioactive collagen peptides, collagen hydrolysate, and the development of novel functional food and beverage ingredients. Furthermore, it will also be important in the extraction of collagen and its application in the food and nutrition industries. Hence, studies on microbial collagenase production and applications would be interesting, rewarding and beneficial to improve the growth of food, nutrition, and other allied sectors.

Conflict of interest

The authors declare that we have no conflict of interest.

Acknowledgements

Gaurav Kumar Pal thanks the Department of Science and Technology (DST), Govt. of India, New Delhi, India for the award of a Research Fellowship. The authors would like to thank anonymous reviewers for the valuable comments provided to improve the manuscript.

References

1. K. E. Kadler, C. Baldock, J. Bella and R. P. Boot-Handford, J. Cell Sci., 2007, 120, 1955–1958.
2. U. Eckhard, P. F. Huesgen, H. Brandstetter and C. M. Overall, J. Proteomics, 2014, 100, 102–114.
3. G. B. Fields, J. Biol. Chem., 2013, 288, 8785–8793.
4. G. K. Pal, T. Nidheesh and P. V. Suresh, Food Res. Int., 2015, 76, 804–812.
5. F. H. Silver, J. W. Freeman and G. P. Seehra, J. Biomech., 2003, 36, 1529–1553.
6. S. W. Manka, F. Carafoli, R. Visse, D. Bihan, N. Raynal, R. W. Farndale, G. Murphy, J. J. Enghild, E. Hohenester and H. Nagase, Proc. Natl. Acad. Sci. U. S. A., 2012, 109, 12461–12466.
7. S. E. Bilek and S. K. Bayram, J. Funct. Foods, 2015, 14, 562–569.
8. C. A. Lima, J. F. Campos, J. L. Lima Filho, A. Converti, M. G. C. da Cunha and A. L. Porto, J. Food Sci. Technol., 2015, 52, 4459–4466.
9. C. A. Lima, J. L. Lima Filho, B. B. Neto, A. Converti, M. G. C. da Cunha and A. L. Porto, Biotechnol. Bioprocess Eng., 2011, 16, 549–560.
10. C. A. Lima, D. A. Viana Marques, B. B. Neto, J. L. Lima Filho, M. G. Carneiro-da-Cunha and A. L. Porto, Biotechnol. Prog., 2011, 27, 1470–1477.
11. Z. Ruszczak and W. Friess, Adv. Drug Delivery Rev., 2003, 55, 1679–1698.
12. C. A. Miles, N. C. Avery, V. V. Rodin and A. J. Bailey, J. Mol. Biol., 2005, 346, 551–556.
13. S. Jus, I. Stachel, W. Schloegl, M. Pretzler, W. Friess, M. Meyer, R. Birner-Gruenberger and G. M. Guebitz, Mater. Sci. Eng., C, 2011, 31, 1068–1077.
14. F. J. O'Brien, B. A. Harley, I. V. Yannas and L. Gibson, Biomaterials, 2004, 25, 1077–1086.
15. D. F. Holmes, H. K. Graham, J. A. Trotter and K. E. Kadler, Micron, 2001, 32, 273–285.
16. D. I. Zeugolis, R. G. Paul and G. Attenburrow, Mater. Sci. Eng., C, 2010, 30, 190–195.
17. D. I. Zeugolis, G. R. Paul and G. Attenburrow, J. Biomed. Mater. Res., Part A, 2009, 89, 895–908.
18. E. C. Soller, D. S. Tzeranis, K. Miu, P. T. So and I. V. Yannas, Biomaterials, 2012, 33, 4783–4791.
19. M. Schlapp and W. Friess, J. Pharm. Sci., 2003, 92, 2145–2151.
20. F. Shahidi and Y. J. Kamil, Trends Food Sci. Technol., 2001, 12, 435–464.
21. A. S. Duarte, A. Correia and A. C. Esteves, Crit. Rev. Microbiol., 2014, 1–21.
22. M. TR Gomes, M. L. Oliva, M. T. P. Lopes and C. E Salas, Curr. Protein Pept. Sci., 2011, 12, 417–436.
23. M. Kim, S. E. Hamilton, L. W. Guddat and C. M. Overall, Biochim. Biophys. Acta, Gen. Subj., 2007, 1770, 1627–1635.
24. N. Tsuruoka, T. Nakayama, M. Ashida, H. Hemmi, M. Nakao, H. Minakata, H. Oyama, K. Oda and T. Nishino, Appl. Environ. Microbiol., 2003, 69, 162–169.
25. Z. Yu, R. Visse, M. Inouye, H. Nagase and B. Brodsky, J. Biol. Chem., 2012, 287, 22988–22997.
26. T. S. Thring, P. Hili and D. P. Naughton, BMC Complementary Altern. Med., 2009, 9, 27.
27. A. Ace Baehaki, M. T. Suhartono, S. Sukarno, D. Syah, A. B. Sitanggang, S. Setyahadi and M. Friedhelm Meinhardt, et al., Afr. J. Microbiol. Res., 2012, 6, 2373–2379.
28. N. Ohbayashi, N. Yamagata, M. Goto, K. Watanabe, Y. Yamagata and K. Murayama, Appl. Environ. Microbiol., 2012, 78, 5839–5844.
29. A. M. Abdel-Fattah, Egypt. Pharm. J., 2013, 12, 109.
30. L. Liu, M. Ma, Z. Cai, X. Yang and W. Wang, Food Technol. Biotechnol., 2010, 48, 151–160.
31. S. M. Daboor, S. M. Budge, A. E. Ghaly, M. S. Brooks and D. Dave, Adv. Biosci. Biotechnol., 2012, 3, 191–203.
32. S. M. Daboor, S. M. Budge, A. E. Ghaly, S. L. Brooks and D. Dave, Adv. Biosci. Biotechnol., 2010, 6, 239–263.
33. R. Jain and P. C. Jain, Indian J. Exp. Biol., 2010, 48, 174–178.
34. S. Preet Kaur and W. Azmi, Curr. Biotechnol., 2013, 2, 17–22.
35. K. Watanabe, Appl. Microbiol. Biotechnol., 2004, 63, 520–526.
36. B. Raskovic, O. Bozovic, R. Prodanovic, V. Niketic and N. Polovic, J. Biosci. Bioeng., 2014, 118, 622–627.
37. A. S. Adhikari, E. Glassey and A. R. Dunn, J. Am. Chem. Soc., 2012, 134, 13259–13265.
38. U. Eckhard, E. Schönauer, D. Nüss and H. Brandstetter, Nat. Struct. Mol. Biol., 2011, 18, 1109–1114.
39. A. Gousterova, I. Goshev, P. Christov, R. Tsvetkova and P. Nedkov, Biotechnol. Biotechnol. Equip., 2003, 17, 81–86.
40. D. Petrova, S. Vlahov and P. Dalev, Biotechnol. Biotechnol. Equip., 2001, 15, 31–38.
41. D. H. Petrova, S. A. Shishkov and S. S. Vlahov, J. Basic Microbiol., 2006, 46, 275–285.
42. D. Petrova, A. Derekova and S. Vlahov, Folia Microbiol., 2006, 51, 93–98.
43. H. Nagano and K. A. To, Biosci., Biotechnol., Biochem., 2000, 64, 181–183.
44. Q. Wu, C. Li, C. Li, H. Chen and L. Shuliang, Appl. Biochem. Biotechnol., 2010, 160, 129–139.
45. C. A. Lima, A. C. F. Júnior, J. L. Lima Filho, A. Converti, D. A. V. Marques, M. G. Carneiro da Cunha and A. L. F. Porto, Biochem. Eng. J., 2013, 75, 64–71.
46. C. A. Lima, P. M. Rodrigues, T. S. Porto, D. A. Viana, J. L. Lima Filho, A. L. Porto and M. G. C. da Cunha, Biochem. Eng. J., 2009, 43, 315–320.
47. S. K. Kim, P. J. Park, J. B. Kim and F. Shahidi, J. Biochem. Mol. Biol., 2002, 35, 165–171.
48. P. J. Park, S. H. Lee, H. G. Byun, S. H. Kim and S. K. Kim, J. Biochem. Mol. Biol., 2002, 35, 576–582.
49. G. F. Fasciglione, S. Marini, S. D'Alessio, V. Politi and M. Coletta, Biophys. J., 2000, 79, 2138–2149.
50. U. Eckhard, E. Schönauer, P. Ducka, P. Briza, D. Nüss and H. Brandstetter, Biol. Chem., 2009, 390, 11–18.
51. H. S. Hamdy, Indian J. Biotechnol., 2008, 7, 333–340.
52. W. Suphatharaprateep, B. Cheirsilp and A. Jongjareonrak, New Biotechnol., 2011, 28, 649–655.
53. R. Bauer, J. J. Wilson, S. T. L. Philominathan, D. Davis, O. Matsushita and J. Sakon, J. Bacteriol., 2013, 195, 318–327.
54. P. Ducka, U. Eckhard, E. Schönauer, S. Kofler, G. Gottschalk, H. Brandstetter and D. Nüss, Appl. Microbiol. Biotechnol., 2009, 83, 1055–1065.
55. O. Matsushita, K. Yoshihara, S. Katayama, J. Minami and A. Okabe, J. Bacteriol., 1994, 176, 149–156.
56. K. Yoshihara, O. Matsushita, J. Minami and A. Okabe, J. Bacteriol., 1994, 176, 6489–6496.
57. N. Teramura, K. Tanaka, K. Iijima, O. Hayashida, K. Suzuki, S. Hattori and S. Irie, J. Bacteriol., 2011, 193, 3049–3056.
58. B. U. Rosso, C. de Albuquerque Lima, T. S. Porto, C. de Oliveira Nascimento, A. Pessoa, A. Converti, M. das Graças Carneiro-da-Cunha and A. L. F. Porto, Fluid Phase Equilib., 2012, 335, 20–25.
59. P. Medina and L. Baresi, J. Microbiol. Methods, 2007, 69, 391–393.
60. L. H. Tran and H. Nagano, J. Food Sci., 2002, 67, 1184–1187.
61. C. J. Doillon, R. Drouin, M.-F. Cote, N. Dallaire, J.-F. Pageau and G. Laroche, J. Biomed. Mater. Res., 1997, 37, 212–221.
62. S. L. Voytik-Harbin, US Pat., 13/192, 276, 2011.
63. S. L. Voytik-Harbin, S. Kreger, B. Bell and J. Bailey, US Pat., 8,512,756, 2013.
64. Y. Zhang, Y. Fu, S. Zhou, L. Kang and C. Li, Anal. Biochem., 2013, 437, 46–48.
65. Y. Sakurai, H. Inoue, W. Nishii, T. Takahashi, Y. Iino, M. Yamamoto and K. Takahashi, Biosci., Biotechnol., Biochem., 2009, 73, 21–28.
66. J. Mukherjee, N. Webster and L. E. Llewellyn, et al., PLoS One, 2009 DOI:10.1371/journal.pone.0007177.
67. K. Thanzami and I. Roy, Electrophoresis, 2008, 29, 1585–1588.
68. M. Okamoto, Y. Yonejima, Y. Tsujimoto, Y. Suzuki and K. Watanabe, Appl. Microbiol. Biotechnol., 2001, 57, 103–108.
69. Y. Z. Zhang, L. Y. Ran, C. Y. Li and X. L. Chen, Appl. Environ. Microbiol., 2015, 81, 6098–6107.
70. D. J. Harrington, Infect. Immun., 1996, 64, 1885–1891.
71. N. D. Rawlings, A. J. Barrett and A. Bateman, Curr. Protoc. Bioinformatics, 2014, 1–25.
72. N. D. Rawlings, M. Waller, A. J. Barrett and A. Bateman, Nucleic Acids Res., 2014, 42, 503–509.
73. N. D. Rawlings, A. J. Barrett and A. Bateman, Nucleic Acids Res., 2012, 40, D343–D350.
74. A. S. Duarte, E. Cavaleiro, C. Pereira, S. Merino, A. C. Esteves, E. P. Duarte, J. M. Tomás and A. C. Correia, Lett. Appl. Microbiol., 2015, 60, 288–297.
75. B. R. Park, R. A. Zielke, I. H. Wierzbicki, K. C. Mitchell, J. H. Withey and A. E. Sikora, J. Bacteriol., 2015, 197, 1051–1064.
76. S. K. Kim, J. Y. Yang and J. Cha, Gene, 2002, 283, 277–286.
77. R. Bauer, K. Janowska, K. Taylor, B. Jordan, S. Gann, T. Janowski, E. C. Latimer, O. Matsushita and J. Sakon, Acta Crystallogr., Sect. D: Biol. Crystallogr., 2015, 71, 565–577.
78. O. Matsushita, C. M. Jung, S. Katayama, J. Minami, Y. Takahashi and A. Okabe, J. Bacteriol., 1999, 181, 923–933.
79. O. Matsushita, T. Koide, R. Kobayashi, K. Nagata and A. Okabe, J. Biol. Chem., 2001, 276, 8761–8770.
80. T. Toyoshima, O. Matsushita, J. Minami, N. Nishi, A. Okabe and T. Itano, Connect. Tissue Res., 2001, 42, 281–290.
81. U. Eckhard and H. Brandstetter, Biol. Chem., 2011, 392, 1039–1045.
82. U. Eckhard, E. Schönauer and H. Brandstetter, J. Biol. Chem., 2013, 288, 20184–20194.
83. U. Eckhard, D. Nüss, P. Ducka, E. Schönauer and H. Brandstetter, Acta Crystallogr., Sect. F: Struct. Biol. Cryst. Commun., 2008, 64, 419–421.
84. R. Miyake, Y. Shigeri, Y. Tatsu, N. Yumoto, M. Umekawa, Y. Tsujimoto, H. Matsui and K. Watanabe, J. Bacteriol., 2005, 187, 4140–4148.
85. A. Kurata, K. Uchimura, S. Shimamura, T. Kobayashi and K. Horikoshi, Appl. Microbiol. Biotechnol., 2007, 77, 311–319.
86. H. K. Kim, Y. R. Ha, H. S. Yu, H. H. Kong and D. I. Chung, Korean J. Parasitol., 2003, 41, 189–196.
87. W. T. Kim, H. H. Kong, Y. R. Ha, Y. C. Hong, H. J. Jeong, H. S. Yu and D. I. Chung, Korean J. Parasitol., 2006, 44, 321–330.
88. R. T. Biaggio, R. R. da Silva, N. G. da Rosa, R. S. R. Leite, E. C. Arantes, T. P. d. F. Cabral, M. A. Juliano, L. Juliano and H. Cabral, Prep. Biochem. Biotechnol., 2015 DOI:10.1080/10826068.2015.1031387.
89. X. L. Chen, B. B. Xie, J. T. Lu, H. L. He and Y. Zhang, Microbiology, 2007, 153, 2116–2125.
90. L. Y. Ran, H. N. Su, G. Y. Zhao, X. Gao, M. Y. Zhou, P. Wang, H. L. Zhao, B. B. Xie, X. Y. Zhang and X. L. Chen, et al., Mol. Microbiol., 2013, 90, 997–1010.
91. L. Y. Ran, H. N. Su, M. Y. Zhou, L. Wang, X. L. Chen, B. B. Xie, X. Y. Song, M. Shi, Q. L. Qin and X. Pang, et al., J. Biol. Chem., 2014, 289, 6041–6053.
92. Y. K. Wang, G. Y. Zhao, Y. Li, X. L. Chen, B. B. Xie, H. N. Su, Y. H. Lv, H. L. He, H. Liu and J. Hu, et al., J. Biol. Chem., 2010, 285, 14285–14291.
93. G. Y. Zhao, X. L. Chen, H. L. Zhao, B. B. Xie, B. C. Zhou and Y. Z. Zhang, J. Biol. Chem., 2008, 283, 36100–36107.
94. T. Nidheesh, P. G. Kumar and P. V. Suresh, Int. Biodeterior. Biodegrad., 2015, 97, 97–106.
95. T. Nidheesh, G. K. Pal and P. V. Suresh, Carbohydr. Polym., 2015, 121, 1–9.
96. N. Thadathil and S. P. Velappan, Food Chem., 2014, 150, 392–399.
97. P. A. Kumar and P. V. Suresh, Mar. Biotechnol., 2014, 16, 202–218.
98. P. V. Suresh, World J. Microbiol. Biotechnol., 2012, 28, 2945–2962.
99. S.-I. Kang, Y.-B. Jang, Y.-J. Choi and J.-Y. Kong, Biotechnol. Bioprocess Eng., 2005, 10, 593–598.
100. A. Kawasaki, H. Nakano, Y. Tsujimoto, H. Matsui, T. Shimizu, T. Nakatsu, H. Kato and K. Watanabe, Acta Crystallogr., Sect. F: Struct. Biol. Cryst. Commun., 2007, 63, 142–144.
101. N. Ohbayashi, T. Matsumoto, H. Shima, M. Goto, K. Watanabe, A. Yamano, Y. Katoh, K. Igarashi, Y. Yamagata and K. Murayama, Biophys. J., 2013, 104, 1538–1545.
102. J. P. Acevedo, V. Rodriguez, M. Saavedra, M. Munoz, O. Salazar, J. A. Asenjo and B. A. Andrews, J. Appl. Microbiol., 2013, 114, 352–363.
103. W. Janwitthayanan, S. Keelawat, S. Payungporn, A. Lowanitchapat, D. Suwancharoen, Y. Poovorawan and C. Chirathaworn, Microbiol. Res., 2013, 168, 268–272.
104. J. Xu, X. Chen, X. Guo and X. Jiang, J. Shanghai Jiaotong Univ., Med. Sci., 2010, 9, 017.
105. L. Rainbow, M. C. Wilkinson, P. J. Sargent, C. A. Hart and C. Winstanley, Curr. Microbiol., 2004, 48, 300–304.
106. C. M. Kemp, P. L. Sensky, R. G. Bardsley, P. J. Buttery and T. Parr, Meat Sci., 2010, 84, 248–256.
107. D. L. Hopkins, P. G. Allingham, M. Colgrave and R. J. van de Ven, Meat Sci., 2013, 95, 219–223.
108. D. L. Hopkins, T. A. Lamb, M. J. Kerr and R. J. van de Ven, Meat Sci., 2013, 93, 838–842.
109. J. Lepetit, Meat Sci., 2008, 80, 960–967.
110. E. A. Foegeding and D. K. Larick, Meat Sci., 1986, 18, 201–214.
111. M. Ha, A. E. D. Bekhit, A. Carne and D. L. Hopkins, Food Chem., 2013, 136, 989–998.
112. A. J. Miller, E. D. Strange and R. C. Whiting, J. Food Sci., 1989, 54, 855–857.
113. G. Y. Zhao, M. Y. Zhou, H. L. Zhao, X. L. Chen, B. B. Xie, X. Y. Zhang, H. L. He, B. C. Zhou and Y. Z. Zhang, Food Chem., 2012, 134, 1738–1744.
114. T. Lafarga and M. Hayes, Meat Sci., 2014, 98, 227–239.
115. T. Lafarga, P. O'Connor and M. Hayes, Peptides, 2014, 59, 53–62.
116. A. Daneault, V. Coxam and Y. Wittrant, Crit. Rev. Food Sci. Nutr., 2015 DOI:10.1080/10408398.2015.1038377.
117. S. Sai-Ut, S. Benjakul, P. Sumpavapol and H. Kishimura, J. Food Process. Preserv., 2015, 39, 394–403.
118. I. Lassoued, L. Mora, R. Nasri, M. Jridi, F. Toldrá, M. C. Aristoy, A. Barkia and M. Nasri, J. Funct. Foods, 2015, 13, 225–238.
119. I. Lassoued, L. Mora, A. Barkia, M. C. Aristoy, M. Nasri and F. Toldrá, J. Proteomics, 2015, 128, 8–17.
120. C. R. Corso, E. J. R. Almeida, G. C. Santos, L. G. Morão, G. S. L. Fabris and E. K. Mitter, Water Sci. Technol., 2012, 65, 1490–1495.
121. S. V. Kanth, R. Venba, B. Madhan, N. K. Chandrababu and S. Sadulla, Dyes Pigm., 2008, 76, 338–347.
122. P. Thanikaivelan, J. R. Rao, B. U. Nair and T. Ramasami, Crit. Rev. Environ. Sci. Technol., 2005, 35, 37–79.
123. S. Sivasubramanian, B. M. Manohar, A. Rajaram and R. Puvanakrishnan, Chemosphere, 2008, 70, 1015–1024.
124. J. Ramundo and M. Gray, J. Wound Ostomy Continence Nurs., 2009, 36, S4–S11.
125. J. Ramundo and M. Gray, J. Wound Ostomy Continence Nurs., 2008, 35, 273–280.
126. M. Cemazar, M. Golzio, G. Sersa, J.-M. Escoffre, A. Coer, S. Vidic and J. Teissie, Hum. Gene Ther., 2011, 23, 128–137.
127. A. L. Cronlund and J. H. Woychik, J. Food Sci., 1987, 52, 857–860.
128. J. F. Ding, Y. Y. Li, J. J. Xu, X. R. Su, X. Gao and F. P. Yue, Food Hydrocolloids, 2011, 25, 1350–1353.
129. L. Guo, P. A. Harnedy, M. B. O'Keeffe, L. Zhang, B. Li, H. Hou and R. J. FitzGerald, Food Chem., 2015, 173, 536–542.
130. L. Nakchum and S. M. Kim, Prep. Biochem. Biotechnol., 2016, 46, 123–130.
131. S. Wang, J. Zhao, L. Chen, Y. Zhou and J. Wu, LWT–Food Sci. Technol., 2014, 55, 210–217.
132. N. J. Stanford, P. Millard and N. Swainston, Front. Cell Dev. Biol., 2015, 3 DOI:10.3389/fcell.2015.00017.
133. C. M. Jung, O. Matsushita, S. Katayama, J. Minami, J. Sakurai and A. Okabe, J. Bacteriol., 1999, 181, 2816–2822.
134. O. Matsushita, C. M. Jung, J. Minami, S. Katayama, N. Nishi and A. Okabe, J. Biol. Chem., 1998, 273, 3643–3648.
135. S. Saran, R. V. Mahajan, R. Kaushik, J. Isar and R. K. Saxena, J. Cleaner Prod., 2013, 54, 315–322.
136. J. Kanagaraj, T. Senthilvelan, R. C. Panda and S. Kavitha, J. Cleaner Prod., 2015, 89, 1–17.
137. H. B. Khandelwal, S. V. More, K. M. Kalal and R. S. Laxman, Clean Technol. Environ. Policy, 2015, 17, 393–405.
138. H. Rosen, Arch. Biochem. Biophys., 1957, 67, 10–15.
139. P. V. Suresh, T. Nidheesh and G. K. Pal, in Enzymes in food and beverage processing, ed. M. Chandrasekaran, Taylor & Francis, CRC Press, New York, 2015, ch. 15, pp. 353–377.

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