Comparative Study of Rhodanese Specific Activity Assay Techniques for Plants and Bacterial Species

Badhane Gudeta

Chemical Engineering Department, Mattu University, Mattu, Ethiopia.

DOI: 10.4236/aer.2025.134004                

Abstract

Rhodanese plays a critical role in cyanide detoxification by catalyzing the conversion of toxic cyanide into the less harmful thiocyanate. This comprehensive review examines recent studies on the purification and characterization of rhodanese from two distinct biological species such as plants and bacteria. In this study, different rhodanese assay techniques and purification methods are evaluated. The results showed that bacterial species, particularly Bacillus cereus and Klebsiella oxytoca, indicated the highest specific activity of 25.30 and 52.7 RU/mg, respectively, outperforming plant sources under similar purification conditions. Factors affecting enzyme activity such as temperature, pH, metal ion inhibition, and substrate specificity were also reviewed to minimize their effects on comparison by selecting nearly optimum temperature and pH, the highest affinity to the same substrate, and minimum effect of metal ion. The study concluded that bacterial species have greater rhodanese specific activity, and Lee modified by Agboola and Okonji and Bradford methods were the most used for rhodanese assay and protein quantification, respectively. Ion exchange chromatography is also the most effective rhodanese purification technique.

Keywords

Rhodanese, Specific Activity, Assay, Substrate Specification, Enzyme, Chromatography

1. Introduction

1.1. Background

Biochemical reactions within living organisms are catalyzed by enzymes. Enzymes play an essential role in various life processes, and their activity must be precisely regulated to ensure proper cellular function [1] [2]. Enzymes are categorized into industrial enzymes, analytical enzymes, and medical enzymes based on their purposes, and they are highly specific catalysts with an active site in their structure where the substrate is converted into a product [3]. Furthermore, research has verified that a broad spectrum of enzymes has been isolated and identified from a wide array of biological origins [4]. Enzymes such as cyanide hydratase [5], 3-mercapto pyruvate sulfurtransferase [6] [7], cyanase [8], cyanide dihydratase [9], cyanide monooxygenase [10], and rhodanese [3]-[8] [11] have been identified as capable of detoxifying cyanide [12].

Rhodanese (thiosulfate: cyanide sulfur transferase, EC 2.8.1.1) is a universal enzyme found in all living organisms and catalyzes the conversion of cyanide into less toxic thiocyanate using a sulfur donor compound [13]-[15]. In a laboratory setting, it undergoes the irreversible transfer of a sulfur atom from thiosulfate to cyanide, resulting in the production of sulfite and thiocyanate [5]-[7] [16].

1.2. Conformational Structure of Rhodanese

Crystalline rhodanese, as isolated from a mammalian source, is a small monomeric sulfurtransferase with a molecular weight of ∼32 kDa containing one reaction site per molecule. The bovine mitochondrial enzyme (Rhobov) is composed of a single polypeptide chain containing 293 amino acid residues [17]. Crystallographic analyses show the polypeptide chain is folded into two domains of equal size and similar three-dimensional tertiary structure [18]. The two structures are arranged around an approximately two-fold axis crossing the domain-domain interface Rhobov Fold as shown in Figure 1. Each domain is composed of a five-stranded parallel sheet, flanked on both sides by α-helices [19]. Hydrophobic interactions stabilize the inter-domain interface to form the active catalytic site. The catalytic residues are in the C-terminal domain, whereas the N-terminal domain stabilizes the active structure [20].

The inactive N-terminal and active C-terminal structurally show identical α/β topology in spite of low sequence identity [α-helix: cyan; β-sheets] [11] [18] [21]. Figure 1 indicates the catalytic and non-catalytic structure of Rhodanese.

1.3. Mechanism of Action

The main process of eliminating cyanide from the body is facilitated by the rhodanese enzyme, which is found in liver mitochondria [15] [22] [23]. A close relationship has been found between rhodanese activity and cyanogenesis, suggesting that the enzymes play a role in enabling plants to convert cyanide through biotransformation processes [15] [24]. The active site of rhodanese is composed of six amino acid residues, with cysteine [Cys] at the first position, facilitating the catalytic process, and the Cys residue is localized in the N-terminal domain, where it is positioned adjacent to an aspartic acid [Asp] residue lacking catalytic function. The compound contains sulfhydryl and aromatic groups nearby, and its catalytic activity is mediated through a double displacement reaction, also known as

Source: [21].

Figure 1. Conformational structure of mitochondrial rhodanese (Rhobov).

a ping-pong mechanism, where a stable persulphide-containing intermediate enzyme is formed [25] [26]. The process consists of two separate half-reactions. In the initial stage of the reaction, sulphur from the substrate is passed to the cysteine residue at the active site, where it forms a persulphide enzyme intermediate [27]. In the latter part of the reaction sequence, a thiophilic acceptor binds to the enzyme-bound persulphide intermediate, giving rise to the product and restoring the free enzyme [15] [27]. The sulfur being transferred is attached to a catalytic cysteine residue that remains constant. The thiocyanate is formed through a nucleophilic reaction with cyanide, formally denoted as S⁰, which results in the regeneration of the active Cys residue for an additional cycle of the reaction. The catalytic mechanism of rhodanese involves transferring thiosulfate to a nucleophilic acceptor [28] [29]. Figure 2 illustrates the stepwise double displacement (ping-pong) catalytic mechanism of rhodanese, showing how the enzyme transfers sulfur between substrates through a covalent enzyme-sulfur intermediate.

The general reaction is

$\underset{\text{Thiosulphate}\text{ion}}{{\text{S}}_2{\text{O}}_3^{2-}}+\underset{\text{Cyanide}\text{ion}}{{\text{CN}}^{-}}\stackrel{\boxed{\text{rhodanese}}}{\to }\underset{\text{Thiocyanide}\text{ion}}{{\text{SCN}}^{-}}+\underset{\text{Sulphite}\text{ion}}{{\text{SO}}_3^{2-}}$(1)

In animal systems, roughly 65% - 80% of the cyanide dosage is converted into thiocyanate via a sulfur donor and free cyanide [30] [31]. Significantly increasing the rhodanese dosage results in a substantial enhancement of the HCN degradation efficiency rate [14]. Rhodanese has been extracted, purified, and characterized from several classes of species due to its cyanide detoxification function [19].

TST = Thiosulphate Sulphur Transferase (Rhodanese).

Figure 2. Stepwise double-displacement catalytic mechanism of rhodanese.

1.4. Biological Source of Rhodanese

Rhodanese was first identified by Lang in 1933 and is found in a broad range of organisms, including both prokaryotes and eukaryotes [15] [32] [33]. It exists in all living organisms, ranging from bacteria to humans. The organism’s function has been assessed in bacteria, yeast, plants, and animals, but how it is distributed across various tissues seems unique to each species [19]. The organism has been found in various animal tissues such as the liver, proventriculus, esophagus, gizzard, cecum, brain, kidney, and lungs, where it is involved in the mitochondrial chain. The liver is the primary source of rhodanese, with the kidney being a secondary source, and rhodanese is also highly prevalent and extracted from a variety of cyanogenic and non-cyanogenic plants [15]. Rhodanese activity in higher plants is significantly lower than in animals, and significantly higher in cyanogenic plants compared to non-cyanogenic plants [34].

This review paper assesses the specific activity of recently published research on purification of rhodanese from bacteria and plants. The main goal of assessing rhodanese activity is to identify the most common assaying and quantifying techniques used in the rhodanese enzyme extraction, and its presence in samples under specific conditions, thereby enabling the comparison of activity levels across different samples and measurement methodologies. The detailed procedure is outlined in the Rhodanese specific activity determination section.

2. Rhodanese Specific Activity Determination Techniques

Rhodanese enzyme activity has been documented in multiple plant species, including cassava leaves, Pentadiplandra brazzeana root, almond, snake tomatoes, meal tomatoes, locust bean seed, maize, and corn. In bacteria like Bacillus cereus, Bacillus licheniformis, wild Klebsiella oxytoca, and Klebsiella edwardsi, the activity of rhodanese is examined. The total rhodanese assay and total protein concentration need to be established before calculating the specific activity.

2.1. Methods for the Rhodanese Assay

Several methods for rhodanese assay have been described. All the rhodanese assay methods, including the Agboola and Okonji method, the modified Lee method, the classical Sorbo method, the Sorbo-Aminlari method, and the Modified Sorbo method, are fundamentally based on the same colorimetric principle: enzymatic conversion of cyanide to thiocyanate, followed by formation of a ferric-thiocyanate complex measured at 460 nm [35]. However, these methods differ in practical parameters such as reaction volume, enzyme volume, substrate concentrations, incubation time, and temperature. Despite these procedural variations, all techniques operate as minor modifications of the same underlying thiocyanate-based colorimetric assay for quantifying rhodanese activity.

The majority of the rhodanese assay methods utilized the approach developed by Agboola and Okonji. The rhodanese assay method measures enzyme activity, which is typically quantified in Rhodanese Units (RU). According to reference [28], the amount of enzyme that yields an optical density measurement of 1.08 at a wavelength of 460 nm is the unit of one RU. The amount of rhodanese that catalyzes the conversion of one micromole (1 μmol) of cyanide ion to thiocyanate in a minute is defined as a rhodanese unit [14] [36]. The details of these methods are explained as follows.

The Lee method, as modified by Agboola and Okonji, is articulated through the utilization of a reaction mixture comprising 0.5 ml of 50 mM borate buffer (pH = 9.4), 0.2 ml of 250 mM potassium cyanide (KCN), and 0.2 ml of 250 mM sodium thiosulfate (Na₂S₂O₃), along with 20 µl of enzyme solution, culminating in a total volume of 1.0 ml; this mixture is subsequently incubated for 1 minute at a temperature of 37 °C, after which the reaction is terminated by the introduction of 0.5 ml of 15% formaldehyde, followed by the incorporation of 1.5 ml of Sorbo reagent. Ultimately, the absorbance is measured spectrophotometrically at a wavelength of 460 nm [14] [37] [38].

The methodology proposed by Agboola and Okonji is characterized by a reaction mixture comprising 250 mM Na₂S₂O₃, 250 mM KCN, and 50 mM borate buffer at pH 9.4, alongside 100 µl of enzyme solution within a final volume of 1.0 ml. 1.5 ml of Sorbo reagent and 0.5 ml of 15% formaldehyde are added to stop the reaction after it has been running for 1 minute at 37˚C. A ferric nitrate solution containing 0.025 g Fe(NO₃)₃∙9H₂O, diluted in 0.74 ml of water and 0.26 ml of concentrated nitric acid, is combined to create the Sorbo reagent. Finally, a wavelength of 460 nm is used to measure the absorbance [25] [35] [36] [38]-[40].

The Sorbo method is explained with an assay mixture of 50 mM borate buffer, pH 9.4, 0.25M KCN, 0.25M Na₂S₂O₃, and 0.1 ml of the enzyme solution in a total volume of 1 ml. After 1 minute of incubation at 37˚C, the reaction is terminated by adding 0.5 ml of 15% formaldehyde. The concentration of thiocyanate produced is determined by the addition of 1.5 ml of Sorbo reagent, where the Sorbo reagent is made up of 10 g Fe(NO₃)₂∙9H₂O, 20 ml HNO_3, and 80 ml distilled water. Then, the absorbance of the reaction medium is read at 460 nm [37].

0.25 ml of 50 mM borate buffer, pH 9.4, 0.1 ml of 250 mM KCN, 0.1 ml of 250 mM Na₂S₂O, and 0.1 ml of the enzyme solution make up the reaction mixture used in the Sorbo and Aminlari technique, which has a total volume of 0.55 ml. After one minute of incubation at 37˚C, 0.25 mL of 15% formaldehyde is added to the reaction mixture to halt it. By adding 2.5 mL of 1% ferric nitrate to 13% nitric acid reagent, the spectrophotometric generation of thiocyanate is measured, and the optical density is measured at 460 nm [37] [41]. A 25 mM citrate-phosphate-borate buffer at a pH of 9.4, 50 mM potassium cyanide (KCN), 50 mM sodium thiosulphate (Na₂S₂O₃), and 50 µl of enzyme solution make up the reaction mixture used in the modified Sorbo technique, which has a total volume of 1.75 ml. After incubating the mixture for two minutes at 25˚C, 0.25 mL of 37% formaldehyde and 1 mL of Sorbo reagent are added to stop the reaction. At 460 nm, the absorbance is measured [41].

The modified Lee method is elaborated as follows: the assay mixture contains 50 mM borate buffer, pH 9.4, 200 mM Na₂S₂O₃, 200 mM KCN, and 200 µl of enzyme in a total volume of 1 ml. The mixture is incubated with 0.5 ml of 15% formaldehyde for 1 min at 37˚C, followed by the addition of an appropriate volume of Sorbo reagent. The absorbance is then read at 460 nm using a spectrophotometer [25].

2.2. Rhodanese Purification Methods

Many enzyme purification methods have been developed over the years. The purification is accomplished in two different stages, i.e., ammonium sulfate precipitation and column chromatography separation [42]. Column chromatography separation includes affinity chromatography on reactive Blue-2 Agarose [43]-[45], ion exchange chromatography on CM-Sephadex C-50 [46] [47], Sephadex G-150 gel filtration chromatography [48] [49], and Sephadex G-75 gel filtration. Traditional enzyme purification procedures often start with ammonium sulfate precipitation [50]. Chromatographic separation of protein mixtures has become one of the most effective and widely used means of purifying individual proteins [51].

Several rhodanese purification techniques have also been observed for various biological extracts. 65% - 80% ammonium sulphate precipitation [52], affinity chromatography on reactive Blue-2 Agarose, ion exchange chromatography on CM-Sephadex C-50 & 25, Sephadex G-150 gel filtration chromatography, and Sephadex G-75 gel filtration are the main rhodanese purification techniques observed during the review.

2.3. Rhodanese Quantification Methods

Determining the concentration of protein is critical in numerous contexts, encompassing enzyme kinetics, enzyme purification, and enzymatic testing. Three of the most widely used methods for protein quantification on a large scale are the Lowry, Bicinchoninic, and Bradford methods [53]-[57]. One of the most commonly used methods for protein purification is the Bradford assay. This method utilizes the binding of Coomassie Brilliant Blue dye to basic amino acid residues in the protein, resulting in a shift in the dye’s absorbance spectrum. The amount of protein present in the sample can then be quantified by measuring the absorbance of the dye-protein complex at 595 nm [57]. This method is utilized to measure the concentration of enzymes after purification, employing bovine serum albumin (BSA) as a standard [28] [58]. Bovine serum albumin [BSA] is the most commonly used standard in protein concentration quantification [59]. For the Bradford enzyme quantification assay to work, basic amino acid residues such as arginine, histidine, phenylalanine, tryptophan, and tyrosine must bind to Coomassie Brilliant Blue G-250 dye at an acidic pH. Hydrophobic interactions in proteins also cause this dye to change colour to blue [39] [54].

The second method applied in the quantification of rhodanese is the Lowry method [60]. This method involves the reduction of copper ions in the presence of protein, resulting in the formation of a complex between the proteins and reduced copper ions. The formation of this complex can be detected by measuring the absorbance of the resulting solution at 750 nm [57]. The Bicinchoninic Acid (BCA) assay is another widely used method for protein quantification. This assay is based on the reduction of Cu²⁺ ions by protein in an alkaline environment to form a complex with BCA, resulting in a color change that can be detected spectrophotometrically at 562 nm [53] [57] [61]. The BCA assay proves to be more specific than the Bradford and Lowry assays, while also displaying reduced sensitivity to interfering substances in the sample [54] [61] [62].

2.4. Calculation of Specific Activity

Enzyme activity assays determine the rate at which an enzyme speeds up a chemical reaction, with that rate directly linked to the amount of enzyme present in the sample. Direct measurement methods can sometimes be less accurate than enzyme activity assays, which consider the actual activity of the enzyme [63]. The specific activity of an enzyme is directly related to its purity within a protein composition. It measures the activity of a specific enzyme on a per-milligram basis of the total protein present. This process is notably important in isolating enzymes from particular proteins to assess their purity. The specific activity is calculated by dividing the number of enzyme units per milliliter by the concentration of protein in milligrams. In the case of Rhodanese, this value is expressed in units of RU per milligram [3] [64].

$\text{Rhodanese}\text{specific}\text{activity}\left(\frac{\text{RU}}{\text{mg}}\right)=\frac{\text{Rhodanese}\text{enzyme}\text{activity,}\text{RU}/\text{ml}}{\text{Total}\text{protein}\text{concentration,}\text{mg}/\text{ml}}$ (1)

2.5. Factors Affecting Enzyme Activity

A range of factors can impact enzyme activity, including temperature [65]-[68], pH levels [67]-[70], the presence of metal ions [71] [72], and substrate concentration levels [73]-[75]. The optimum temperature of rhodanese has been reported from 45˚C - 60˚C, while optimum pH values range between 6 and 8.0 for different sources. Another key consideration is the enzyme’s selectivity for its substrate. The enzyme’s substrate specificity was examined by assessing its activity towards structurally analogous compounds. The effect of various substrates [sodium thiosulphate, sodium metabisulphite, and ammonium persulphate] has been investigated by calculating the Michaelis-Menten constants [Vmax and Km] [37]. Research has documented the substrate specificity of rhodanese isolated from plants, bacteria, and animals. The impact of these heavy metal compounds [BaCl₂, NiCl₂, KCl, NaCl, ZnCl₂, CaCl₂, HgCl₂, SnCl₂, and MnCl₂] at concentrations of 1 and 10 millimolar in the reaction mixture on rhodanese activity is documented [76]. Although the cited studies investigated rhodanese activity across a range of temperature, pH values, substrate concentrations, and metal ion levels, this study focused only on an optimum operating condition, i.e., 50˚C and a pH of 8.0, as well as its strong affinity for sodium thiosulfate and minimal interference from metal ions. Studying these factors is crucial for comparing the enzyme activities obtained from recently isolated rhodanese. Table 1 represents rhodaneses’ assaying methods, purification techniques, and specific activity for plants and bacterial species.

Table 1. Rhodanese-specific activity and its reliance on biological sources.

Biological Sources	Enzyme assay procedures/protein quantification methods	Protein purification methods	Results			Reference
			Total activity, RU	Total protein, mg	Specific activity RU/mg
Cassava leaves	Lee modified by Agboola and Okonji/Bradford	80% Ammonium Sulphate Precipitation	145.5	965.3	2.03	[14]
Bitter cassava	Lee modified by Agboola and Okonji/Bradford	80% Ammonium Sulphate Precipitation	791.32	263.51	3.00	[38]
		Ion-exchange Chromatography on CM-Sephadex C-25	338.51	63.22	5.35
Almond (Prunus amygdalus) nuts	Agboola and Okonji/Bradford	80% Ammonium Sulphate Precipitation	85.77	29.7	2.89	[25]
		Ion-exchange chromatography on CM-Sephadex C-50	54.24	11.32	4.79
		Reactive Blue-2 Agarose Affinity Chromatography	26.79	5.26	5.09
Seed and mesocarp of snake tomatoes (Trichosanthes cucumerina Linn)	Agboola and Okonji/Bradford	80% Ammonium Sulfate Precipitation	S: 41.3 M:81.7	S: 2.1 M: 9.3	S: 19.7 M: 8.8	[77]
Tomato (Solanum lycopersicum Mill)	Agboola and Okonji/Bradford	80% Ammonium Sulphate Precipitation	150.63	53.9	2.79	[78]
		Ion-exchange Chromatography on CM-Sephadex C-50	94.54	31.45	3.01
		Biogel P-100 Size Exclusion Chromatography	56.1	12.62	4.45
Seed, Mesocarp, and Capsule of Snake Tomato Fruit	Lee, modified by Agboola and Okonji/Bradford	80% Ammonium Sulphate Precipitation	S:74.39 M:5.5 C: 6.56	S:1480.86 M:332.65 C:737.87	S: 0.0502 M: 0.0165 C:0.0089		[79]
		Ion-exchange chromatography on CM-Sephadex C-25	S:0.32	S: 10.16	S: 0.0315
		Reactive Blue-2 Agarose Affinity Chromatography	S:7.604	S:0.742	S:0.0621
Locust bean seed (Parkia biglobosa)	Sorbo and Aminlari /Bradford	80% Ammonium Sulfate Precipitation	105.77	33.7	3.14		[80]
		Ion-exchange Chromatography on CM-Sephadex C-50	74.24	20.11	3.69
		Reactive Blue - Affinity Chromatography	36.79	7.06	5.21
Pentadiplandra brazzeana (Baill) root	Agboola and Okonji/Bradford	80% ammonium sulphate precipitation	71.80	31.12	2.31		[81]
		Reactive Blue - Affinity Chromatography	15.6	3.24	4.82
Malted maize (Zea mays)	Lee, modified by Agboola and Okonji/Bradford	85% Ammonium Sulphate Precipitation	102.87	21.06	4.88		[82]
		Ion-exchange Chromatography on CM-Sephadex C-25	9.88	1.95	5.07
Seed of Guinea Corn (Sorghum arundinaceum Stapf)	Lee, modified by Agboola and Okonji/Bradford	Acetone Precipitation	43.09	5924.29	0.007		[83]
		Biogel P-100 Gel Filtration	1.59	57.88	0.027
		DEAE–Sephadex ion-exchange chromatography	0.18	7.00	0.0026
Bacillus cereus	Sorbo/Bradford Sorbo	80% Ammonium Sulphate Precipitation	52.095	6.053	8.74		[37]
		Ion-exchange Chromatography on CM-Sephadex C-50	52.40	2.27	23.08
		Biogel P-100 Size Exclusion Chromatography	33.88	1.339	25.30
Bacillus licheniformis	Lee/Bradford	85% Ammonium Sulfate Precipitation	59.08	20.25	2.92		[40]
		Ion-exchange chromatography on CM-Sephadex C-50	4.77	0.39	12.23
		Sephadex G-100 gel filtration chromatography	20.42	1.50	13.63
Klebsiella edwardsii	Modified Sorbo/Bradford	85% Ammonium Sulphate Precipitation	4.80	205.625	0.0234		[41]
		Ion exchange on DEAE Cellulose	8.07	170.625	0.0473
Wild Klebsiella Oxytoca	Agboola and Okonji/Bradford	70% Ammonium Sulfate Precipitation	239.7	6.2	38.6		[39] [84]
		Sephadex G-100 gel filtration chromatography	336.13	97.02	3.47
		Ion-exchange chromatography on CM-Sephadex C-25	213.5	4.1	52.7

3. Discussion

The evaluation and comparison of rhodanese distribution in plants and bacteria, and the effectiveness of different purification techniques, were assessed in this review. Studies from diverse plant and bacterial species consistently demonstrate substantial variation in rhodanese specific activity, reflecting the strong influence of biological source and purification methodology.

In nearly all studies, ammonium sulfate precipitation from 70% - 85% concentration served as the initial crude purification step. This produced moderate specific activities in plant sources such as cassava (2.02 RU/mg) and almond (2.89 RU/mg), but extremely low activity in guinea corn seeds (0.007 RU/mg). In contrast, Bacillus cereus and Bacillus licheniformis exhibited considerably higher values of 8.74 RU/mg and 2.92 RU/mg, respectively, after the initial precipitation procedure. Further purification through ion-exchange chromatography markedly enhanced activity to 23.08 RU/mg in Bacillus cereus and 12.23 RU/mg in Bacillus licheniformis. The highest activity reported across all studies was found in wild Klebsiella oxytoca, reaching 52.7 RU/mg after ion-exchange purification.

Plant materials such as tomato and snake tomato generally showed only modest improvements after purification in chromatography. However, Reactive Blue-2 agarose affinity chromatography demonstrated strong potential for plant extracts, yielding specific activities of 5.05 RU/mg in almond, 5.21 RU/mg in locust bean, and 4.82 RU/mg in Pentadiplandra brazzeana root. This suggests that affinity chromatography may be particularly suitable for enhancing plant rhodanese purity.

Conversely, size-exclusion chromatography techniques, Biogel P-100 and Sephadex G-100, were more effective for bacterial sources, producing activities of 25.3 RU/mg and 13.63 RU/mg in Bacillus licheniformis. Despite undergoing multiple purification steps, Klebsiella edwardii consistently displayed low specific activities of 0.0234 - 0.0473 RU/mg, indicating limited intrinsic enzyme abundance or stability in that species.

In comparing the rhodanese activity assay techniques, most studies relied on the Abgoola and Okonji and modified Lee methods, while others applied variations of the classical Sorbo assay. Although all are based on the same thiocyanate-ferric complex colorimetric principle, differences in reaction volumes, incubation times, enzyme quantities, and substrate concentrations can yield slight variation in measured activity. Likewise, for protein quantification, the Bradford assay dominated due to its dye-protein interaction and minimal interference, while fewer studies employed the Lowry and BCA methods. Such differences in assay and quantification techniques may introduce small variations in calculated specific activity.

Although the cited studies evaluated rhodanese activity under a broad range of temperatures, pH values, substrate concentrations, and metal-ion levels, the comparison presented in this review relies extensively on the optimum conditions and minimal-inhibitor states reported in each study. By focusing only on the peak activity parameters, major variability among methods was minimized. Nonetheless, minor differences in extraction procedures, buffer compositions, and assay setups may still contribute to small variations in reported activity. These methodological differences represent a minor limitation when making direct cross-study comparisons.

The consistently observed rhodanese activities in bacteria may arise from several biological and biochemical factors. Bacterial systems often exhibit higher enzyme expression levels as part of their adaptive response to harsh or cyanide-rich environments. In addition, bacterial rhodanese may possess inherently greater catalytic efficiency and superior folding stability compared to plant enzymes (36). The simpler cellular matrix of bacteria, lacking complex polysaccharides, secondary metabolites, and phenolic compounds common in plant tissues, also facilitates more efficient extraction and purification, reducing potential interference that can depress measured activity.

4. Conclusion

This review highlights the substantial differences in rhodanese specific activity among various biological sources and purification techniques. Across all examined studies, bacterial species consistently demonstrated significantly higher specific activity compared to plant sources, particularly when purified by ion exchange and gel-filtration chromatography. The practical implication of this trend is that bacterial species, especially Bacillus cereus, Bacillus licheniformis, and wild Klebsiella oxytoca, represent promising candidates for industrial and environmental applications. Overall, the evaluated purification and assay approaches confirm that bacteria provide a more efficient and scalable source of rhodanese for future industrial applications. The Agboola and Okonji and modified Lee assay techniques emerged as the most widely applied methods for determining rhodanese activity, while the Bradford assay remained the dominant approach for protein quantification.

Acknowledgements

I highly acknowledge my family for their continuous and invaluable support.

Funding

This research did not receive any specific grant from funding agencies in the public, commercial, or not-for-profit sectors.

Availability of data and materials

The datasets used are available from the corresponding author upon reasonable request.

Ethical statement

This article is a review and does not involve any new experiments on humans or animals. All data discussed are taken from previously published studies and are properly cited.

Author’s Contribution

Badhane G. Collect recently published data that are relevant to the title, and write the manuscript, recheck, and evaluate by comparing with various published papers.

Abbreviation

BCA	Bicinchoninic Acid
BSA	Bovine Serum Albumin
CM	Carboxymethyl
CM-Sephadex	Carboxymethyl Sephadex
DEAE	Diethylaminoethyl
Fe(NO3)3∙9H2O	Ferric Nitrate Nonahydrate
G-100/G-150	Gel Filtration Chromatography Media (Sephadex)
HNO3	Nitric acid
KCN	Potassium cyanide
Kₘ	Michaelis constant
RU	Rhodanese unit
${\text{SO}}_3^{2-}$	Sulphite ion
Vₘₐₓ	Maximum velocity (in enzyme kinetics)

Conflicts of Interest

The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this review paper.

References

[1]	Emma, R. (2023) Exploring Methods for Accurate Measurement of Enzyme Concentration in Biological Samples. Journal of Clinical and Bioanalytical Chemistry, 7, Article No. 141.
[2]	Bevilacqua, A., Maciel, M.R., Pascarelli, S., Dindo, M., Shen, A.Q. and Laurino, P. (2024) Enzyme Activity Regulates Substrate Diffusion by Modulating Viscosity in Crowded Milieu.
[3]	Vitolo, M. (2020) Brief Review on Enzyme Activity. World Journal of Pharmaceutical Research, 9, 60-76.
[4]	Morgab, M.A.H., Hussein, A.D. and Hadi, S.T. (2023) Extraction and Purification of Polyphenol Oxidase from Edible Mushroom (Agaricus bisporus) and Its Use in the Manufacture of Pastries. Food Research, 7, 64-70.[CrossRef]
[5]	Malmir, N., et al. (2021) Cyanide Hydratase Modification Using Computational Design and Docking Analysis for Improved Binding Affinity in Cyanide Detoxification. Molecules, 26, Article No. 1799.
[6]	Kaczor-Kamińska, M., Kaminski, K. and Wróbel, M. (2021) The Expression and Activity of Rhodanese, 3-Mercaptopyruvate Sulfurtransferase, Cystathionine γ-Lyase in the Most Frequently Chosen Cellular Research Models. Biomolecules, 11, Article No. 1859.[CrossRef] [PubMed]
[7]	Rao, S.P., Dobariya, P., Bellamkonda, H. and More, S.S. (2023) Role of 3-Mercaptopyruvate Sulfurtransferase (3-MST) in Physiology and Disease. Antioxidants, 12, Article No. 603.[CrossRef] [PubMed]
[8]	Kennedy, A., Brennan, A., Mannion, C. and Sheehan, M. (2021) Suspected Cyanide Toxicity in Cattle Associated with Ingestion of Laurel—A Case Report. Irish Veterinary Journal, 74, Article No. 6.[CrossRef] [PubMed]
[9]	Šťastná, K., Martínková, L., Rucká, L., Křístková, B., Příhodová, R., Bojarová, P., et al. (2024) Design and Development of Spectrophotometric Enzymatic Cyanide Assays. Analytical and Bioanalytical Chemistry, 417, 697-704.[CrossRef] [PubMed]
[10]	Malmir, N., Fard, N.A., Aminzadeh, S., Moghaddassi-Jahromi, Z. and Mekuto, L. (2022) An Overview of Emerging Cyanide Bioremediation Methods. Processes, 10, Article No. 1724.[CrossRef]
[11]	Motl, N., Skiba, M.A., Kabil, O., Smith, J.L. and Banerjee, R. (2017) Structural and Biochemical Analyses Indicate That a Bacterial Persulfide Dioxygenase-Rhodanese Fusion Protein Functions in Sulfur Assimilation. Journal of Biological Chemistry, 292, 14026-14038.[CrossRef] [PubMed]
[12]	Jiang, Y. (2023) Analysis of the Toxicity and Treatment Methods of Cyanide. Theoretical and Natural Science, 27, 235-240.[CrossRef]
[13]	Bebarta, V.S. and Nath, A.K. (2025) Redirecting Intermediary Metabolism to Counteract Cyanide Poisoning. The FASEB Journal, 39, e70709.[CrossRef] [PubMed]
[14]	Funke Adedugbe, O., Desola Owolala, O. and Samuel Ilesanmi, O. (2021) Properties of Partially Purified Rhodanese from Leaves of Cassava in Owo Southwestern Nigeria. International Journal of Bioorganic Chemistry, 6, Article No. 21.[CrossRef]
[15]	Buonvino, S., Arciero, I. and Melino, S. (2022) Thiosulfate-Cyanide Sulfurtransferase a Mitochondrial Essential Enzyme: From Cell Metabolism to the Biotechnological Applications. International Journal of Molecular Sciences, 23, Article No. 8452.[CrossRef] [PubMed]
[16]	Wodu, E., Oputu, A.F., Asheshemi, O.F. and Ozomah, C.I. (2022) Characterization of Rhodanese Extracted from Synodontis Schall Liver. EAS Journal of Nutrition and Food Sciences, 4, 46-53.[CrossRef]
[17]	Libiad, M., Sriraman, A. and Banerjee, R. (2015) Polymorphic Variants of Human Rhodanese Exhibit Differences in Thermal Stability and Sulfur Transfer Kinetics. Journal of Biological Chemistry, 290, 23579-23588.[CrossRef] [PubMed]
[18]	Liu, D., Yang, F., Liu, Z., Wang, J., Huang, W., Meng, W., et al. (2020) Structure of TBC1D23 N-Terminus Reveals a Novel Role for Rhodanese Domain. PLOS Biology, 18, e3000746.[CrossRef] [PubMed]
[19]	Benisch, R. and Giessen, T.W. (2024) Structural and Biochemical Characterization of an Encapsulin-Associated Rhodanese from Acinetobacter baumannii. Protein Science, 33, e5129.[CrossRef] [PubMed]
[20]	Benisch, R. (2024) Structural and Biochemical Characterization of an Encapsulin-Associated Rhodanese from Acinetobacter baumannii.
[21]	Zakwani, A.D., et al. (2023) University of Groningen Discovery of a Small Molecule Allosteric Activator of Human Thiosulfate Sulfurtransferase Al Dahmani-Al Zakwani, Zayana.
[22]	Kruithof, P.D. (2023) University of Groningen Discovery of a Small Molecule Allosteric Activator of Human Thiosulfate Sulfurtransferase Al Dahmani-Al Zakwani, Zayana. 4-7.
[23]	Al-Dahmani, Z.M., Li, X., Wiggenhauser, L.M., Ott, H., Kruithof, P.D., Lunev, S., et al. (2022) Thiosulfate Sulfurtransferase Prevents Hyperglycemic Damage to the Zebrafish Pronephros in an Experimental Model for Diabetes. Scientific Reports, 12, Article No. 12077.[CrossRef] [PubMed]
[24]	Tutić, A., Vuković Domanovac, M., Kučić Grgić, D. and Miloloža, M. (2025) Biological Degradation of Cyanide, Thiocyanate, and Phenolic Compounds in Wastewater. Kemija u industriji, 74, 219-234.[CrossRef]
[25]	Ehigie, A.F., Abdulrasak, M.A. and Ehigie, O.L. (2018) Biochemical Properties of Rhodanese from Almond (Prunus amygdalus) Nuts. Pan African Journal of Life Sciences, 1, 17-24.[CrossRef]
[26]	Wang, Q., Li, H., Xia, Y., Xun, L. and Liu, H. (2021) Saccharomyces Cerevisiae Rhodanese RDL2 Uses the Arg Residue of the Active-Site Loop for Thiosulfate Decomposition. Antioxidants, 10, Article No. 1525.[CrossRef] [PubMed]
[27]	Selles, B., Moseler, A., Rouhier, N. and Couturier, J. (2019) Rhodanese Domain-Containing Sulfurtransferases: Multifaceted Proteins Involved in Sulfur Trafficking in Plants. Journal of Experimental Botany, 70, 4139-4154.[CrossRef] [PubMed]
[28]	Supapong, C. and Cherdthong, A. (2021) Rhodaneses Enzyme Addition Could Reduce Cyanide Concentration and Enhance Fiber Digestibility via in Vitro Fermentation Study. Fermentation, 7, Article No. 207.[CrossRef]
[29]	Supapong, C., Sommai, S., Khonkhaeng, B., Suntara, C., Prachumchai, R., Phesatcha, K., et al. (2022) Effect of Rhodanese Enzyme Addition on Rumen Fermentation, Cyanide Concentration, and Feed Utilization in Beef Cattle Receiving Various Levels of Fresh Cassava Root. Fermentation, 8, Article No. 146.[CrossRef]
[30]	Ražná, K., Harenčár, Ľ. and Kučka, M. (2022) The Involvement of microRNAs in Plant Lignan Biosynthesis—Current View. Cells, 11, 2151.[CrossRef] [PubMed]
[31]	Logue, B.A., Hinkens, D.M., Baskin, S.I. and Rockwood, G.A. (2010) The Analysis of Cyanide and Its Breakdown Products in Biological Samples. Critical Reviews in Analytical Chemistry, 40, 122-147.[CrossRef]
[32]	Luo, Y., Chatre, L., Melhem, S., Al-Dahmani, Z.M., Homer, N.Z.M., Miedema, A., et al. (2023) Thiosulfate Sulfurtransferase Deficiency Promotes Oxidative Distress and Aberrant NRF2 Function in the Brain. Redox Biology, 68, Article ID: 102965.[CrossRef] [PubMed]
[33]	Al-Dahmani, Z.M., Hadian, M., Ruiz-Moreno, A.J., Maria, S.A., Batista, F.A., Zhang, R., et al. (2023) Identification and Characterization of a Small Molecule That Activates Thiosulfate Sulfurtransferase and Stimulates Mitochondrial Respiration. Protein Science, 32, e4794.[CrossRef] [PubMed]
[34]	Kaleta, K., Misterka, A., Rydz, L., Wróbel, M. and Jurkowska, H. (2023) Correlation between the Level of Sulfane Sulfur and the Expression/Activity of Sulfurtransferases in Chicken Tissues—A Possible Ways of Cyanide Detoxification. Biologia, 79, 101-108.[CrossRef]
[35]	Wodu, E. and Frank-Oputu, A. (2022) Physicochemical Characterization of Synodontis Schall Gills Rhodanese. Global Academic Journal of Agriculture and Biosciences, 4, 31-35.[CrossRef]
[36]	Wodu, E., Frank-Oputu, A., Lucky-Ben, K., et al. (2021) Studies on Some Physiochemical Properties of Crude Extracts of Rhodanese from Liver and Kidney of an Adult Ram. Global Academic Journal of Agriculture and Bio Sciences, 3, 61-65.
[37]	Babamotemi, O.I., Raphael, E.O., Odunayo, A., Nkem, T., Chinwe, O. and Ademakinwa, O. (2019) Studies on Some Physicochemical Properties of Rhodanese Synthesized by Bacillus Cereus Isolated from the Effluents of Iron and Steel Smelting Industry. African Journal of Biochemistry Research, 13, 1-8.[CrossRef]
[38]	Akintimehin, E.S., Adetuyi, F.O., Karigidi, K.O., Okonji, R.E. and Akinnubi, C.O. (2020) Effects of Postharvest Storage on Cyanide Content and Activity of Partially Purified Rhodanese from Bitter Cassava (Manihot utilissima) Tubers. Food Science and Applied Biotechnology, 3, 157-166.[CrossRef]
[39]	Itakorode, B.O. and Apalowo, O.E. (2022) Isolation and Statistical Optimization of Rhodanese (A Thiosulphate Sulphur Transferase) Production Potential of Klebsiella oxytoca JCM 1665 Using Response Surface Methodology. Nova Biotechnologica Et Chimica, 21, e1083.[CrossRef]
[40]	Ehigie, A.F., Okonji, R.E., Oladiran, W.A. and Ehigie, L. (2019) Isolation and Characterization of Rhodanese from Bacillus Licheniformis Obtained from Industrial Effluents of Steel Rolling Industry in Ilorin. IRA-International Journal of Applied Sciences, 13, 14-34.[CrossRef]
[41]	Adedeji, O.A., Aladesanmi, O.T., Famakinwa, O.A. and Okonji, R.E. (2017) Bioefficiency of Indigenous Microbial Rhodanese in Clean-Up of Cyanide Contaminated Stream in Modakeke, Ile-Ife, Osun State, Nigeria. Journal of Bioremediation & Biodegradation, 8, 1-7.[CrossRef]
[42]	Behera, S.S., Nivedita, S., Behera, H.T. and Ray, L. (2023) Partial Purification and Characterization of an Extracellular Chitin Deacetylase from Streptomyces Griseoinacarnatus RB7AG Isolated from Chilika Lake. The Microbe, 1, Article ID: 100020.[CrossRef]
[43]	Murkar, N.S., Nagrale, K.D., Charde, M.S., Chakole, R.D., Jadhav, V.S. and Karad, V. (2023) Comprehensive Approaches towards Affinity Chromatography. International Journal of Novel Research and Development, 8, 152-161.
[44]	Saladi, V.S., Sc, M. and Phil, M. (2025) Protein Purification—Affinity Chromatography Methods.
[45]	Kumaraswamy, G., Kumar, J.M.R., Sheshagirirao, J.V.L.N., et al. (2021) Affinity Chromatography: A Review. World Journal of Pharmaceutical Research, 4, 1567-1574.
[46]	Asare, M.C., Kubde, J.A., Bakal, R.L., Hatwar, P.R., Kalamb, V.S. and Tambakhe, P.K. (2025) Ion Exchange Chromatography: A Comprehensive Review. GSC Biological and Pharmaceutical Sciences, 31, 26-37.[CrossRef]
[47]	Simanjuntak, B., Julian, H. and Kresnowati, M.T.A.P. (2022) Downstream Process of Xylanase Production from Oil Palm Empty Fruit Bunches: A Review. IOP Conference Series: Earth and Environmental Science, 1034, Article ID: 012046.[CrossRef]
[48]	Bose, K. (2022) Textbook on Cloning, Expression and Purification of Recombinant Proteins. In: Bose, K., Ed., Textbook on Cloning, Expression and Purification of Recombinant Proteins, Springer, 1-311.
[49]	Rehan, S.F.M. and Sambhaji, U.A. (2025) A Review of Scalable and Efficient Techniques for the Purification of Lectins. Analytical and Bioanalytical Chemistry Research, 12, 259-268.
[50]	Alhazmi, L.S. and Alshehri, W.A. (2025) Purification and Biochemical Characterization of α-Amylase from Newly Isolated Bacillus cereus Strain and Its Application as an Additive in Breadmaking. Polish Journal of Microbiology, 74, 48-59.[CrossRef] [PubMed]
[51]	Coskun, O. (2016) Separation Tecniques: Chromatography. Northern Clinics of Istanbul, 3, 156-160.[CrossRef] [PubMed]
[52]	Tamilanban, R., Santhi Velayudhan, S., Rajadas, S.E. and Harshavardhan, S. (2017) Purification and Characterization of an Extracellular Cellulase Produced Using Alkali Pretreated Rice Straw by Stenotrophomonas maltophilia Management of Red Palm Weevil View Project Production of Cellulase View Project Purification and Characterization. International Journal of Biological Research, 2, 45-54. https://www.biologyjournal.in
[53]	Yong, J., Hakobyan, K., Xu, J., Mellick, A.S., Whitelock, J. and Liang, K. (2023) Comparison of Protein Quantification Methods for Protein Encapsulation with ZIF-8 Metal-Organic Frameworks. Biotechnology Journal, 18, 1-16.[CrossRef] [PubMed]
[54]	Dobler, S. (2024) Evaluating the Efficacy of Protein Quantification Methods on Membrane Proteins.
[55]	Zaguri, M., Kandel, S., Rinehart, S.A., Torsekar, V.R. and Hawlena, D. (2021) Protein Quantification in Ecological Studies: A Literature Review and Empirical Comparisons of Standard Methodologies. Methods in Ecology and Evolution, 12, 1240-1251.[CrossRef]
[56]	Löptien, J., Vesting, S., Dobler, S. and Mohammadi, S. (2024) Evaluating the Efficacy of Protein Quantification Methods on Membrane Proteins. Open Biology, 14, 1-26.[CrossRef] [PubMed]
[57]	Benjamin, A. (2023) Protein Quantification Methods: Advantages and Limitations. Journal of Biological Chemistry, 7, 142. https://www.alliedacademies.org/clinical-and-bioanalytical-chemistry/
[58]	Agbo, A.O. and Odibo, F.C.J. (2025) Partial Purification and Characterization of Alkaline Proteases from Maize (Oba Super 2). Bangladesh Journal of Scientific and Industrial Research, 60, 9-24. [Google Scholar] [CrossRef]
[59]	Giles, H., et al. (2025) An Investigative Study into the Suitability of the Bradford Assay for Rapid Protein Determination in Whey. Food Chemistry, 499, 147375.[CrossRef]
[60]	Ahmed, E.M. and Ibrahim, M.M. (2015) Evaluation and Improvement of Rhodanese from Isolated Mortieralla isabellina through Plakett-Burman and Box-Behnken Models. Der Pharmacia Lettre, 7, 173-185.
[61]	Shrief, E. (2021) Factors Affecting Enzyme Activity.
[62]	Bublitz, T.A., Oliva, R.L., Hupe, A. and Joergensen, R.G. (2023) Optimization of the Bicinchoninic Acid Assay for Quantifying Carbohydrates of Soil Extracellular Polymeric Substances. Plant and Soil, 498, 699-709.[CrossRef]
[63]	Sears, P.S. (2020) Units of Enzymatic Activity. Catalysis from A to Z.
[64]	Fares, A. (2025) (ENZYMES 2) Activity—Regulation Medical Biochemistry.
[65]	Daniel, R.M., Danson, M.J., Eisenthal, R., Lee, C.K. and Peterson, M.E. (2007) The Effect of Temperature on Enzyme Activity: New Insights and Their Implications. Extremophiles, 12, 51-59.[CrossRef] [PubMed]
[66]	Kabir, M.F. and Ju, L. (2023) On Optimization of Enzymatic Processes: Temperature Effects on Activity and Long-Term Deactivation Kinetics. Process Biochemistry, 130, 734-746.[CrossRef]
[67]	Arcus, V.L., Prentice, E.J., Hobbs, J.K., Mulholland, A.J., Van der Kamp, M.W., Pudney, C.R., et al. (2016) On the Temperature Dependence of Enzyme-Catalyzed Rates. Biochemistry, 55, 1681-1688.[CrossRef] [PubMed]
[68]	Mohd Zin, Z., Azman, S.N.S., Yahya, F., M., H. and Zainol, M.K. (2022) Effect of pH and Temperature on Antioxidant Enzymes Activities in Morinda citrifolia L. (Mengkudu) Leaves Extract. Food Research, 6, 60-67.[CrossRef]
[69]	Gu, Y. (2024) The Effect of Buffer pH on Enzyme Activity. Theoretical and Natural Science, 33, 137-147.[CrossRef]
[70]	Sharif, N. (2024) Effects of Ph on Enzyme Activity in Pakistan. Journal of Chemistry, 3, 49-59.[CrossRef]
[71]	Prejanò, M., Alberto, M.E., Russo, N., Toscano, M. and Marino, T. (2020) The Effects of the Metal Ion Substitution into the Active Site of Metalloenzymes: A Theoretical Insight on Some Selected Cases. Catalysts, 10, Article No. 1038.[CrossRef]
[72]	Forero, N., Liu, C., Sabbah, S.G., Loewen, M.C. and Yang, T.C. (2023) Assay Development for Metal-Dependent Enzymes-Influence of Reaction Buffers on Activities and Kinetic Characteristics. ACS Omega, 8, 40119-40127.[CrossRef] [PubMed]
[73]	Saladi, V.S., Sc, M. and Phil, M. (2025) Enzyme Activity—Substrate Specificity: A Complete Guide. 1-14.
[74]	Mbira, C. (2024) Influence of Substrate Concentration on Enzyme Activity in Bio Catalysis. Journal of Chemistry, 3, 48-58.[CrossRef]
[75]	Kumari, J.A., Rao, P.C., Padmaja, G. and Madhavi, M. (2020) Effect of Substrate Concentration on Soil Enzyme Urease. International Journal of Current Microbiology and Applied Sciences, 9, 1150-1158.[CrossRef]
[76]	Kaczor-Kamińska, M., Sura, P. and Wróbel, M. (2020) Multidirectional Changes in Parameters Related to Sulfur Metabolism in Frog Tissues Exposed to Heavy Metal-Related Stress. Biomolecules, 10, Article No. 574.[CrossRef] [PubMed]
[77]	Obasi, U.E., Kenneth, A. and Benard, I.O. (2017) A Comparative Analysis of Rhodanese Enzyme Isolated from the Seed and Mesocarp of Snake Tomato (T. cucumerina). Advances in Agricultural Biotechnology, 1, 1-7.
[78]	Ehigie, A.F., Abdulrasak, M.A., Adeleke, G.E. and Ehigie, O.L. (2019) Comparison of Rhodanese Activity and Distribution in Tomato (Solanum lycopersicum Mill.) Plant Parts and Its Physicochemical Characterization. Solanum Journal of Plant Biochemistry & Physiology, 7, 1-8.
[79]	Okonji, R.E., et al. (2017) Comparative Studies on the Partial Purification and Characterization of Rhodanese from Seed and Mesocarp of Snake Tomatoes (Trichosanthes cucumerina Linn.). Journal of Agricultural Biotechnology and Sustainable Development, 9, 9-15. [Google Scholar] [CrossRef]
[80]	Ehigie, A.F., Abdulrasak, M.A., Ojeniyi, F.D. and Ehigie, O.L. (2019) Kinetic Properties of Rhodanese from African Locust Bean Seeds (Parkia biglobosa). Asian Journal of Biomedical and Pharmaceutical Sciences, 9, 18-23. [Google Scholar] [CrossRef]
[81]	Raphael, E.O., Bamidele, S.F., Leonard, O.E., Zainab, A.A. and Olajumoke, O.O. (2017) Physicochemical Properties of Rhodanese: A Cyanide Detoxifying Enzyme from Pentadiplandra brazzeana (Baill) Root. African Journal of Biotechnology, 16, 704-711.[CrossRef]
[82]	Adetuyi, F., Akintimehin, E., Karigidi, K.O. and Raphael, O. (2017) Isolation, Partial Purification and Characterization of Rhodanese from Malted Maize (Zea mays).
[83]	Chukwuejim, S., et al. (2019) Isolation, Purification and Characterization of Rhodanese from the Seeds of Guinea Corn (Sorghum arundinaceum Stapf.). FUOYE Journal of Pure and Applied Sciences (FJPAS), 4, 147-159.
[84]	Oluwasola, B., Emmanuel, O., Duah, I. and Emuebie, R. (2024) Characterization of Rhodanese Synthesized by the Wild and EMS-Mutated Klebsiella oxytoca JCM1665. Journal of Agriculture and Food Research, 18, 101358.[CrossRef]