Optimisation of a general procedure for limonene synthase enzyme isolation, purification, and characterisation

Introduction

As a key designer of specialised plant metabolites, limonene synthase is a crucial plant enzyme that is a member of the structural terpene synthase (TPS) family. Geranyl Pyrophosphate (GPP), a universal ten-carbon precursor, is complexly cyclized by this enzyme to produce limonene, a monocyclic monoterpene. This metabolic process is a crucial branch point in the cytoplasmic Mevalonate (MVA) and plastidial Methylerythritol Phosphate (MEP) pathways, directing carbon flow into volatile essential oils and specialised defence mechanisms. Zerbe and Karunanithi (2019) [15] . Strict stereospecificity characterises limonene synthase's function. The active-site pocket controls the production of different mirror-image enantiomers depending on the particular plant species. For instance, it creates (+)-limonene in the flavedo tissues of citrus fruits while producing (-)-limonene in the Lamiaceae family, which includes peppermint and spearmint. The initial ionisation of the GPP substrate, the creation of a temporary Linalyl Pyrophosphate (LPP) intermediate, and a final cyclisation followed by exact deprotonation are the coordinated stages that make up the reaction process. The enzyme is a great model for structural biology and contemporary protein engineering because of its exquisite control over unstable carbocation intermediates. Wüst et al., (2001) [24] . From an industrial and economic Standpoint, the enzyme is a crucial bio-engine for producing limonene, a high-value platform chemical whose global market value is expected to surpass$2.7 billion by 2034. Total product yield and overall cost efficiency in large-scale Commercial operations are determined by the enzyme's accuracy and catalytic efficiency. Limonene is a recognised safe natural flavouring agent that is frequently utilised in the food and beverage industries. At the same time, it serves as a key top-note ingredient for contemporary perfumes in the fragrance and cosmetics sectors. The substance serves as a potent, biodegradable, non-toxic degreaser in addition to its conventional aromatic uses, taking the place of dangerous industrial solvents like xylene or chlorinated hydrocarbons. Tufvesson et al., (2011) [22] . Spearmint (Mentha spicata) and peppermint (Mentha x piperita), two aromatic mint species, are great model systems for researching and separating monoterpene cyclases. Citrus crops provide significant amounts of the terpene overall, but mint plants have unique biological advantages that enable quick, target-specific enzyme recovery. The many peltate glandular trichomes that cover the surfaces of immature leaves are what propel this efficiency. These specialised structures serve as active, localised biological factories that are nearly entirely focused on the synthesis and storage of terpenes. Prakash et al., (2016) [19] . Mint concentrates the expression of limonene synthase directly within the secretory cells of these exterior glands, in contrast to other plant systems where secondary enzymes are dispersed throughout intricate internal tissue matrices. As a result, it is possible to recover the active target protein with low background contamination and high initial specific activity. Mint plants are also perfect for protocol optimisation because of their quick growth cycles, which guarantee a consistent supply of new, increasing leaves (1 to 3 cm in length) where transcript expression peaks. However, it is still very difficult to transfer high-resolution separation techniques to simple academic labs. Automated cell disrupters, low-temperature ultracentrifugation, and fast protein liquid chromatography (FPLC) apparatus are commonly used in established protocols. In order to close this operational gap, this research project offers an efficient and economical workflow that uses readily available chemical methods, such as visible-spectrum colorimetric assays and Aqueous Two-Phase Systems (ATPS), to isolate, purify, and characterise active limonene synthase using simple lab equipment. Maffei et al., (2007) [18] .

Materials and methods

Chemical reagents and laboratory infrastructure

Pecialized buffer engineering and optimization

The foundational extraction buffer was designed to preserve enzyme tertiary structure and prevent degradation during mechanical cell breakdown. The formulation was prepared by dissolving precise quantities of chemical stabilizers in deionized water: 1.017 g of magnesium chloride hexahydrate, 0.931 g of EDTA, 6.302 g of sodium sulfite, 10.0 g of PVP polyphenol binder, and 2.0 g of purified BSA protection fraction. These components were thoroughly blended into 500 mL of deionized water, followed by the addition of 200 mL of stabilizing glycerol, 100 mL of 0.5 M Tris-HCl stock (pH 7.5), and 10 mL of β-Mercaptoethanol. The mixture was stirred continuously until completely clear and uniform, transferred to a 1000 mL volumetric flask, brought to volume with deionized water, and inverted repeatedly. The completed buffer was stored at 4 °C to prechill prior to plant tissue contact. The specific structural and biochemical role of each Component is detailed in Table 1. 2 mg / mL Acts as a sacrificial target to limit proteolysis and surface Adsorption.

Phytochemical mapping via thin layer chromatography

To evaluate the background secondary metabolites coextracted with the protein, a qualitative Thin Layer Chromatography profile was developed. The stationary matrix was prepared by mixing 22.5 g of silica Gel G (containing a gypsum binder) with 45 mL of distilled water (1:2 w/v ratio). The mixture was ground in a mortar to remove air bubbles, poured uniformly over clean glass slides, and allowed to air-dry. The dried plates were activated in an oven at 110 °C for 45 minutes, then cooled in a moisture-free desiccator.

The plant extract was prepared by crushing 1.0 g of fresh mint leaves in 10 mL of 70% analytical Ethanol (1:10 w/v ratio), followed by fine filtration. Chromatographic development was conducted using a glass chamber saturated with an optimized polar mobile phase: ethyl acetate, glacial acetic acid, formic acid, and distilled water in a strict 50:5.5:5.5:13 (v/v) ratio. Using a micro-capillary pipette, small spots of the extract were applied along a baseline 1.0 cm from the plate edge. The plate was placed in the chamber, ensuring the liquid level remained below the sample dots. The solvent migrated upward via capillary action. When the solvent front approached the top margin, the run was stopped, the plate dried, and the front line marked. Separated spots were visualized under long-wave UV light (366 nm) and sprayed with a 5% ferric chloride (FeCl3) solution to detect phenolic hydroxyl groups. Retardation factors were calculated using the standard formula RF = Distance traveled by solute / Distance traveled by solvent front

Tissue homogenization and enzyme isolation

Soluble enzyme extraction used a low-temperature processing method to prevent thermal denaturation. First, 1.0 g of frozen mint leaves was thoroughly pulverized in a pre-chilled mortar and pestle with 10 mL of ice-cold extraction buffer. After primary tissue breakdown, an additional 30 mL of buffer was introduced, and the slurry was stirred continuously on ice for 15 minutes to maximize protein recovery. The crude mixture was filtered through four layers of cheesecloth to remove large cellular fragments. The collected filtrate was centrifuged at 15,000 rpm for 30 minutes at 4 °C to pellet chloroplast fragments and cellular aggregates. The supernatant was recovered and centrifuged again under identical conditions for 30 minutes to yield a highly clarified crude enzyme extract, which was stored on ice for immediate purification.

Queous two-phase system (ATPS) liquid-liquid extraction

To isolate the target enzyme without risk of structural damage, a liquid-liquid Aqueous Two-Phase System was constructed on a weight-to-weight (w/w) baseline. The system combined PEG 2000 (15-20% w/w) as the polymer phase and dipotassium phosphate (10-15% w/w) as the highdensity salt phase. The clarified crude extract was added at a loading concentration of 15% (w/w), and the total mass was balanced with distilled water to cross the system's binodal curve. The complete 10 g system was gently inverted for 45 minutes at 4 °C to allow the protein partition to reach thermodynamic equilibrium across the liquid interface. Phase separation was accelerated by low-speed centrifugation at 3,500 rpm for 8 minutes, creating a sharp boundary between the upper polymer phase and the lower salt phase. The hydrophobic target enzyme partitioned into the upper PEG layer, which was recovered using a fine micropipette. A back-extraction step using a fresh phosphate salt wash transferred the purified enzyme back into an aqueous salt environment for activity validation.

Uantitative total protein estimation (lowry method)

Total protein concentration was determined via the classic Lowry technique. Solution I was prepared with 1% sodium potassium tartrate mixed with 0.5% copper sulfate pentahydrate in distilled water. Solution II consisted of 4% sodium carbonate dissolved in 0.1 N sodium hydroxide. The active working Reagent A was prepared fresh by blending equal parts of Solution I and Solution II. Reagent B was constructed by a 1:3 dilution of Folin-Ciocalteu phenol reagent with distilled water. A standard calibration curve was established using eleven test tubes (0 to 10) containing Increasing amounts of a 1 mg/mL BSA stock solution (0 to 100 μL), balanced to a total volume of 100 μL with distilled water. Tube 0 served as the blank. Purified enzyme samples were added to separate tubes at identical volumes. Next, 2.0 mL of Reagent A was added to each tube and incubated for 10 minutes at room temperature to form copper-peptide complexes. Then, 500 μL of Reagent B was added, mixed quickly, and left for 30 minutes to complete the color reduction. Absorbance values were measured at 750 nm on a digital spectrophotometer, and sample values were plotted against the standard curve.

Pectrophotometric enzyme activity assay

Enzyme activity was monitored via a colorimetric assay using a 1% potassium permanganate (KMnO4) solution as an oxidative indicator that targets the reactive double bonds of the produced limonene. A control reference line was created using 1 mL of fresh leaf extract mixed with 4 mL of 0.1 M phosphate buffer (pH 7.0) to establish baseline background compounds. The assay was initiated by diluting 250 μL of the purified enzyme with 750 μL of phosphate buffer. Next, 100 μL of this enzyme dilution was combined with 1.0 mL of phosphate buffer and 1.0 mL of the control sample. The tracking run began with the addition of 500 μL of the 1% KMnO4 indicator. The mixture was diluted fivefold with 30 mL of phosphate buffer to maintain optimal absorbance readability. The reaction was incubated at room temperature, and changes in optical density were recorded at 525 nm at fixed intervals (0, 15, 30, 45, 60, 75 minutes, and a terminal overnight 20-hour endpoint) to assess substrate conversion over time.

Molecular weight verification via sds-page electrophoresis

Molecular weight and sample purity were verified using discontinuous Sodium Dodecyl Sulfate Polyacrylamide Gel Electrophoresis. The gel assembly used vertical glass plates sealed at the base with a boiled 1% agarose layer. A 12% polyacrylamide resolving gel was prepared with 12 mL of 30% acrylamide stock, 6 mL of water, 12 mL of 2.5X resolving buffer (pH 8.8), 250 μL of 10% APS, and 36 μL of TEMED catalyst. The mixture was poured between the plates and overlaid with water for one hour to solidify. After removing the water layer, a 5% stacking gel composed of 2.6 mL acrylamide stock,10 mL water, 3.0 mL of 5X stacking buffer (pH 6.8), 150 μL of APS, and 20 μL of TEMED was cast on top, and a sample comb was inserted. Prepared protein samples (20 μL extract mixed with 5 μL of 5X sample buffer) were denatured in a boiling water bath at 100 °C for 5 minutes. The samples and a prestained molecular weight ladder (10-250 kDa) were loaded into the wells. Electrophoresis ran at a constant 100 V and 10 mA in 1X Tris-glycine-SDS running buffer. The gel was stained with Coomassie Brilliant Blue R-250 solution, rinsed with water, and cleared in a destaining solution under gentle agitation until the protein bands were clearly defined.

Results

Buffer properties and extraction performance

The preparation of the optimized extraction buffer yielded a completely clear, transparent solution with no visible precipitation or suspended particles. Post-formulation tracking confirmed an optimal pH of 7.5, which remained stable during extended storage at 4 °C. Pre-chilling the buffer successfully limited sample warming during leaf grinding, effectively preventing the rapid structural breakdown typically caused by heat-sensitive proteases.

Phytochemical migration profiles

The thin-layer separation of the Mentha extract successfully resolved its major secondary metabolites. Under visible light, several yellow spots migrated along the plate, indicating the presence of flavonoid glycosides. Spraying with the ferric chloride reagent produced distinct blue-grey and dark grey bands, confirming the location of free hydroxyl groups on the aromatic rings of the phenolic compounds. Calculating the retardation factors identified three major compound groups: a dominant phenolic band appearing between 0.40 and 0.60 (corresponding to rosmarinic acid), a distinct grey-green zone at 0.45 (indicative of caffeic acid), and a highly polar flavonoid cluster at 0.42 (representing rutin, eriocitrin, and Luteolin). These results are summarized in Table 2. The optimized PEG/phosphate system demonstrated excellent phase separation, crossing the binodal curve to form a sharp interfacial boundary after low-speed centrifugation. Due to its hydrophobic surface features, the active enzyme partitioned preferentially into the upper PEGrich phase. This step successfully isolated the target protein from highly polar plant sugars and dense debris, which remained trapped in the lower salt-rich layer.

Total protein quantification values

Plotting the absorbance values of the standard BSA series at 750 nm produced a reliable, linear calibration curve. The line of best fit is defined by the following regression equation Y = 0.0025x + 0.0125 (R² = 0.9583)

Where y represents the recorded absorbance and x is the calculated protein mass in micrograms.

The absorbance values of the purified enzyme samples ranged from 0.545 to 1.723, exceeding the highest standard point due to the high density of the recovered protein. Midrange sample volumes (40 to 80 μL) provided the most reliable values within the linear range of the assay, yielding an estimated average stock concentration of 5.27 ± 1.05 μg/μL, as detailed in Table 3.

Concentration

During the active enzyme assay, the initial absorbance at 525 nm dropped rapidly from 1.597 to 1.328 within the first 45 minutes, reflecting the rapid oxidation of the newly formed monoterpene double bonds by the KMnO_4 reagent. This was followed by a brief stabilization phase between 45 and 75 minutes. Over extended incubation, secondary degradation caused the absorbance to drop to 0.362 at the 20-hour endpoint, corresponding to a total color loss of 77.3%. These colorimetric changes confirm active limonene production, as detailed in Table 4.

Discussion

The optimal composition of the extraction buffer, which was created to counteract the difficult chemical environment inside mint cells, was crucial to the effective extraction of catalytically active limonene synthase from Mentha tissue.

In order to maintain the protein's natural folding and stop the acid-catalyzed degradation of its volatile substrate, it was essential to maintain an alkaline pH of 7.5 using the 50 mM Tris-HCl system. The quick oxidation of natural phenolic chemicals is a significant problem when working with mint proteins. These reactive compounds were effectively sequestered in this work by adding 1% PVP and 50 mM sodium bisulfite, which stopped the extract from turning brown and prevented protein aggregation. Maffei et al., (2007) [18] . By adding 5 mM MgCl2, which supplies the necessary divalent metal cofactors required to initiate the ionisation of geranyl pyrophosphate, the isolated enzyme's catalytic preparedness was preserved. The combination contained 2.5 mM EDTA, but its concentration was regulated to chelate trace heavy metal impurities without competing for the essential magnesium ions. Additionally, the protein was stabilised during extraction by the addition of 20% glycerol and 2 mg/mL BSA. By successfully reducing non-specific adsorption and thermal breakdown, these additives produced a superior crude extract that is perfect for downstream colorimetric tracking and structural validation. Shine (1973) [20] . This study's optimised whole-tissue extraction strategy offers straightforward substitutes for conventional techniques. Although gland-specific separation methods, like bead-beating surface strips, exclude inner leaf proteins and produce high initial specific activity, they are timeconsuming and require specialised equipment. Our wholetissue approach successfully reduced protein breakdown while optimising overall enzyme recovery when paired with stringent temperature control at 4 °C and a fast centrifuge run at 15,000 rpm. Khan et al., (2026) [16] . The Mentha extract's secondary metabolite profile was effectively mapped by the thin-layer chromatography results. The selectivity required to resolve complicated phenolic compounds was obtained by using a highly polar solvent mixture consisting of ethyl acetate, glacial acetic acid, formic acid, and water. By reducing band tailing, inhibiting ionisation, and stabilising the phenolic hydroxyl groups, the acidity of the mobile phase was crucial. The equilibrium between compound polarity and migration speed is shown in the vertical distribution of bands on the silica gel G plate. Strong hydrogen bonds with the silanol groups on the plate caused the polar flavonoid cluster (rutin and eriocitrin) to exhibit lower Rf values, whereas less polar phenolic acids migrated farther. Hawrył et al., (2015) [12] . The hydrophobic surface characteristics of limonene synthase are highlighted by its preferential migration into the upper PEG phase during ATPS purification. The protein has non-polar outer surfaces that help bind lipophilic substrates, just like many plant terpene cyclases. To achieve thermodynamic stability, the enzyme is drawn into the top layer of our optimised system by the non-polar, friendly environment created by the polyethylene glycol chains. The potassium phosphate buffer's salting-out action, which raised the aqueous phase's surface tension and guided the target protein into the polymer layer, further improved this partitioning. This liquid-liquid separation eliminates the possibility of denaturation, which is frequently brought on by harsh chemical solvents. Soni (2025) [21] . Strong internal consistency is shown in the quantitative data obtained using the colorimetric assays. The accuracy of our protein values is confirmed by the BSA curve's good linear fit (R² = 0.9583). A dependable stock concentration of 5.27 μg/μL was obtained by employing mid-range quantities, despite the fact that our sample absorbance values exceeded the standard range because of the high protein density. The reported values for active plant gland tissue are in line with this high yield. Kummari et al., (2022) [17] . In a similar vein, all three tracking wavelengths demonstrated good linearity (R² = 1.0) in our KMnO4 activity experiment. The indicator's quick reactivity with the double bonds of the generated limonene is reflected in the sharp decline in absorbance at 525 nm over the first 45 minutes, indicating active catalysis. The relevance of our consistent 60-minute incubation period for producing repeatable data is shown by the stabilisation phase that follows. (Hoang et al., (2025) [13] . Lastly, structural validation of successful enzyme isolation is provided by our SDS-PAGE data. Our observed band of 50-60 kDa corresponds to the size of the mature monomer following the removal of the N-terminal plastid transit peptide, whereas native plant limonene synthases normally migrate approximately 70-75 kDa as complete pre-proteins. The target bands' sharpness and clarity attest to the protein's structural stability and lack of major breakdown during the extraction process. Tsakalidou et al., (1994) [23] .

Conclusion

Using simple institutional laboratory equipment, this study effectively developed and validated an optimised, economical framework for isolating, purifying, and characterising active plant limonene synthase. We were able to reduce the problems of phenolic oxidation and quick protein breakdown that are common in plant tissues by using a specialised stabilisation buffer. By utilising the hydrophobic surface characteristics of the enzyme to efficiently partition it into the top polymer layer, the integration of Aqueous Two-Phase Systems (ATPS) provided a mild liquid-liquid substitute for costly chromatography column setups.

Strong internal consistency was shown in the quantitative results from the Lowry and potassium permanganate colorimetric tests, offering a dependable way to monitor enzyme concentrations and activity kinetics without the need for sophisticated gas chromatography systems. Lastly, structural confirmation was obtained by SDS-PAGE analysis, which resolved a clear, crisp band in the 50-60 kDa range that represents the enzyme's mature monomer. By bridging the gap between sophisticated biochemical theory and real-world laboratory application, this proven workflow shows that excellent, professional enzymology research can be successfully carried out in conventional academic settings.

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