Determination of Lipolytic Enzyme Activities

Abstract

Pseudomonas aeruginosa is a versatile human opportunistic pathogen that produces and secretes an arsenal of enzymes, proteins and small molecules many of which serve as virulence factors. Notably, about 40 % of P. aeruginosa genes code for proteins of unknown function, among them more than 80 encoding putative, but still unknown lipolytic enzymes. This group of hydrolases (EC 3.1.1) is known already for decades, but only recently, several of these enzymes have attracted attention as potential virulence factors. Reliable and reproducible enzymatic activity assays are crucial to determine their physiological function and particularly assess their contribution to pathogenicity. As a consequence of the unique biochemical properties of lipids resulting in the formation of micellar structures in water, the reproducible preparation of substrate emulsions is strongly dependent on the method used. Furthermore, the physicochemical properties of the respective substrate emulsion may drastically affect the activities of the tested lipolytic enzymes. Here, we describe common methods for the activity determination of lipase, esterase, phospholipase, and lysophospholipase. These methods cover lipolytic activity assays carried out in vitro, with cell extracts or separated subcellular compartments and with purified enzymes. We have attempted to describe standardized protocols, allowing the determination and comparison of enzymatic activities of lipolytic enzymes from different sources. These methods should also encourage the Pseudomonas community to address the wealth of still unexplored lipolytic enzymes encoded and produced by P. aeruginosa.

1 Introduction

Lipases and phospholipases A (PLA) are carboxylester hydrolases (EC 3.1.1) that catalyze the cleavage of ester bonds for example of triacylglycerol and phospholipid substrates, respectively. Both esterases and lipases hydrolyze triacylglycerols; however, esterases show specificity for water-soluble, short-chain fatty acid (less than ten carbon atoms) substrates, while lipases usually hydrolyze water-insoluble, long-chain fatty acid (more than nine carbon atoms) substrates [1–3]. Most lipases show strongly enhanced activity at the interface between the hydrophobic triacylglycerol substrate and water, this phenomenon called “interfacial activation” was proposed to distinguish between esterases and lipases [4]. Meanwhile, structural and kinetic studies of lipases and esterases have challenged this hypothesis [5–7]. Furthermore, broad substrate specificity and substrate promiscuity were observed among carboxylesterases and a number of enzymes were identified that possess esterase, lipase, and PLA activities at the same time [8, 9]. The structural similarity of typical substrates for all three groups of lipolytic enzymes (Fig. 1) suggests a plausible explanation for the observed substrate promiscuity. In living cells, lipases are thought to catalyze exclusively the hydrolysis of ester substrates; however, they are also capable to catalyze the backwards reaction, i.e., the synthesis of esters at low water conditions. Such esterification reactions play an important role for a variety of biotechnological applications, e.g., in the synthesis of enantiopure compounds [10] and acylated carbohydrates [11, 12], thus making lipases the most important group of biocatalysts for the chemical industries [13–15].

Fig. 1

Tripalmitoylglycerol, dipalmitoylphosphatidylcholine, and monopalmitoyl-phosphatidylcholine as typical lipase, phospholipase, and lysophospholipase substrates, respectively, show structural similarity at sn1 and sn2 positions which both represent cleavage sites for these three types of lipolytic enzymes. Lipases can hydrolyze ester bonds at all three positions (sn1, sn2, and sn3); cleavage sites of phospholipases A1 (PLA1), A2 (PLA2), C (PLC), and D (PLD), lysophospholipases A1 (LysoPLA1), A2 (LysoPLA2), C (LysoPLC), and D (LysoPLD) are indicated with arrows. Oxygen, nitrogen, and phosphorus atoms are indicated in red, blue, and brown, respectively (Color figure online)

The genus Pseudomonas comprises a diverse group of medical, environmental, and biotechnological important bacteria [16–19] known to synthesize a variety of different lipolytic enzymes. Pseudomonas aeruginosa represents the best studied member [20] of this group with its genome sequenced and carefully annotated [21]. Here, we focus on lipolytic enzymes from P. aeruginosa although the assays described below are certainly applicable for other lipolytic enzymes as well. The systematic sequence analysis of the P. aeruginosa PA01 genome [21] using amino acid sequences of known lipases and phospholipases and the keywords “phospholipase,” “esterase,” and “lipase” revealed a total number of 104 genes encoding putative and hypothetical lipolytic enzymes. Out of these, 88 genes encode enzymes without homology to any previously characterized lipolytic enzymes, thus underlining the biosynthetic capacity of P. aeruginosa for the production of still unexplored enzymes. A number of lipolytic enzymes including phospholipases C (PlcA, PlcB, PlcH, PlcN), phospholipase D (PldA), phospholipase A2 (ExoU), phospholipase B (PlbF), lysophospholipase (TesA), lipases (LipA, LipC), and esterases (EstA, AchE, PA3859) were identified in P. aeruginosa (Table 1). Recently, we have newly demonstrated PLA2 and LysoPLA activities in P. aeruginosa PA01 associated with membrane and periplasmic localized enzymes [22].

Table 1 Lipolytic enzymes of P. aeruginosa PAO1

The industrial [18, 34, 35] and biomedical [36, 37] potential of lipolytic enzymes still provoke the discovery of novel (phospho)lipases with properties distinct from known enzymes. To this end, metagenomic libraries are constructed and directed evolution methods are applied to identify enzymes with novel properties. These methods require manipulation and screening of a large number of enzyme variants using practical and general activity assays applicable for a wide spectrum of lipolytic enzymes. In contrast, newly discovered enzymes need to be distinguished from already existing ones by using specific and sensitive activity assays with physiologically relevant substrates [38, 39].

A large number of methods have been developed to measure the catalytic activity of lipases [40] comprising mainly three types of assays: (a) quantification of released fatty acids by titrimetric or radiometric methods, (b) determination of released colored or fluorescent products by photometric methods, and (c) detection of changes in biophysical properties of substrates during lipolysis by conductometric, turbidometric, tensidometric, or microscopic methods. A very sensitive and sophisticated method is based on the measurement of changes in surface pressure at the interface of water and lipid substrates, which can be present as a lipid monolayer [40], bilayers [40] or drops [41, 42]. In short, the decrease in surface tension between water and for example a lipid monolayer upon hydrolysis by lipases can directly be measured by a tensiometer (e.g., EasyDyne Krüss, Germany). In general, methods directly detecting changes in biophysical properties of substrates are difficult to establish, require expensive equipment, and do not allow for a high throughput, and hence we do not discuss them in detail here.

Methods that rely on titrimetric measurements are convenient and widely used as long as inexpensive and physiological lipids are used as substrates. However, since large amounts of substrates (usually several hundreds of mg) are needed, the use of synthetic phospholipid substrates renders these assays expensive. Methods utilizing fluorogenic and chromogenic substrates are much more sensitive than titrimetric methods, but they cannot be applied for studying cellular functions of certain enzymes because they rely on the hydrolysis of artificial substrates.

A survey of the literature quickly reveals that even simple assays developed decades ago are used by different researches with slight modifications, thus hampering the comparison of biochemical properties of lipolytic enzymes. Here, we describe detailed protocols of commonly used lipolytic enzyme assays and we will discuss potential drawbacks and strategies how to overcome them.

2 Materials

Prepare all substrate solutions using ultrapure water with electrical resistivity of 18 M Ω cm and analytical grade reagents. All substrates and chemicals can be purchased from Sigma (St. Louis, MO, USA) or Avanti Polar Lipids (Alabaster, Alabama, USA) unless otherwise stated. For agar-plate assays, use distilled water and sterilize all solutions by autoclaving for 20 min at 120 ºC or by filtration through filters with a pore size of 0.22 μm (unless indicated otherwise). We recommend to freshly prepare buffers needed for activity assays rather than to add toxic sodium azide for conservation. The accuracy of any activity assay strongly depends on the careful preparation of all solutions; therefore, pay attention when weighing small amounts of substrates and pipetting small volumes of solutions. Buffers can usually be stored for up to 2 weeks at room temperature or up to 1 month at 8 ºC. Warm up the assay buffers to the temperature chosen for the assays and ensure that pH is adjusted at this same temperature (see Note 1).

2.1 Agar-Plate Assays

2.1.1 Tributyrin Emulsion Assay [43]

• 1. Luria–Bertani (LB) agar medium [44]: 10 g/L bacto-tryptone, 10 g/L sodium chloride, 5 g/L bacto-yeast extract, 15 g/L agar. Add distilled water, dissolve the components, adjust to pH 7 with NaOH, and autoclave.

• 2. Tributyrin emulsion: 50 % (v/v) tributyrin, sterilized by filtration through 0.22 μm membranes, 50 g/L sterile gum arabic dissolved in sterile distilled water. Mix the emulsion for 1 min using a homogenizer (e.g., Ultra Turrax, IKA Labortechnik, Germany) rinsed with 70 % (v/v) ethanol.

2.1.2 Rhodamine B Assay [45]

• 1. Luria–Bertani (LB) agar medium [44]: 10 g/L bacto-tryptone, 10 g/L sodium chloride, 5 g/L bacto-yeast extract, 15 g/L agar. Add distilled water, dissolve the components, adjust to pH 7 with NaOH, and autoclave.

• 2. Rhodamine B solution: 1 mg/mL dissolved in sterile distilled water; sterilize by filtration through 0.22 μm membranes.

• 3. Olive oil emulsion: mix 25 g/L olive oil with sterile distilled water, sterilize by filtration, and emulsify by mixing for 1 min with a homogenizer rinsed with 70 % (v/v) ethanol.

2.1.3 Egg Yolk Assay [46]

• 1. Luria–Bertani (LB) agar medium [44]: 10 g/L bacto-tryptone, 10 g/L sodium chloride, 5 g/L bacto-yeast extract, 15 g/L agar. Add distilled water, dissolve the components, adjust to pH 7 with NaOH, and autoclave.

• 2. Egg-yolk emulsion: 75 g/L egg yolk, 25 g/L taurocholic acid, 20 mM CaCl₂ dissolved in distilled water. Autoclave for 7 min at 120 °C. Emulsify by mixing for 1 min with a homogenizer rinsed with 70 % (v/v) ethanol.

2.2 Titrimetric Assays [47, 48]

• 1. Lipase and PLA substrates: tributyrine, triolein, and olive oil (consists mainly of triolein) are available as liquids; mono- and diacyl-phospholipids and egg yolk are available as solids (Sigma Aldrich, Fluka or Avanti Polar Lipids) (see Note 2).

• 2. Assay buffer: 1 mM Tris–HCl, pH 8; 15 mM NaCl; 2 mM CaCl₂; 4 mM taurodeoxycholate (see Note 3).

• 3. Titrant A: 1 M NaOH in water.

• 4. Titrant B: 50 mM NaOH in water.

• 5. Automatic titrator device (available for example from Mettler Toledo, Switzerland, or Radiometer Analytical, Denmark).

• 6. Homogenizer: bath sonicator, probe sonicator, or blender.

2.3 Colorimetric and Fluorometric Assays

2.3.1 Assays Using p-Nitrophenyl Acyl Ester Substrates

• 1. VIS spectrophotometer: for measurements in cuvettes (0.5–2 mL reaction mixture) or in microplates (20–200 μL reaction mixture) (e.g., microplate reader SpectraMax 250, Molecular Devices Corp.).

• 2. Cuvettes or microplates.

• 3. Plastic reaction tubes of 2 mL capacity.

• 4. Thermoconstant incubator, possibly with agitation.

2.3.1.1 p-Nitrophenyl-Palmitate Assay [49]

• 1. Substrate stock: 20 mM p-NP palmitate in isopropanol.

• 2. Assay buffer: 50 mM Na₂HPO₄; 50 mM KH₂PO₄; 5 mM sodium deoxycholate and 1 g/L arabic gum, adjust to pH 8 with NaOH.

• 3. p-NP standard stock solution: 20 mM p-nitrophenol in assay buffer.

2.3.1.2 p-Nitrophenyl-Butyrate Assay [50, 52]

• 1. Substrate stock: 20 mM p-NP butyrate in acetonitrile.

• 2. Assay buffer: 100 mM K₂HPO₄, pH 7.2.

• 3. p-NP standard stock solution: 20 mM p-nitrophenol in assay buffer.

2.3.2 Assays Using 4-Methylumbelliferyl Acyl Ester Substrates [53, 54]

• 1. Fluorescence spectrometer: for measurements in cuvettes (0.5–2 mL reaction mixture) or in microplates (20–200 μL reaction mixture).

• 2. Glass cuvettes (1 × 1 cm) or microplates suitable for fluorescence measurements (available from PerkinElmer) (see Note 4).

• 3. Plastic reaction tubes of 2 mL capacity.

• 4. Substrate stock: 1 mM 4-MU heptanoate in tetrahydrofuran (see Note 5).

• 5. Assay buffer: 20 mM Tris–HCl, pH 8; 1 mM EDTA; 300 μM taurodeoxycholate.

• 6. 4-MU standard stock solution: 1 mM sodium salt of 4-MU in assay buffer.

2.3.3 Assays Using Natural (Phospho)Lipid Substrates [55, 56]

• 1. VIS spectrophotometer: for measurements in cuvettes (0.5–2 mL reaction mixture) or in microplates (20–200 μL reaction mixture).

• 2. Cuvettes or microplates.

• 3. Plastic reaction tubes of 2 mL capacity.

• 4. Thermoconstant incubator allowing agitation.

• 5. Non-esterified fatty acid HR series (NEFA-HR) kit (Wako Chemicals, Richmond, USA) consists of two sets, reagent 1 set (R1) and reagent 2 set (R2). R1 set contains color reagent A and solvent A, and R2 set contains color reagent B and solvent B. The exact composition of all reagents is listed in the instruction manual provided by the manufacturer, http://www.wako-chemicals.de/DWD/_111327/upload/media_132965.pdf.

• 6. NEFA free fatty acid standards (Wako Chemicals).

• 7. Assay buffer: 40 mM Tris–HCl, pH 7.5; 1 % (v/v) Triton X-100.

• 8. Lipid substrates: 13.4 mM lipid (available from Sigma-Aldrich, Fluka, Avanti Polar Lipids) in assay buffer (see Note 6).

2.4 Adrenalin Fingerprinting Assay [57]

• 1. Substrates: 10 mM of compounds listed in Table 2 dissolved in acetonitrile.

  Table 2 Commercially available polyol acetate substrates used for lipase fingerprinting with the adrenalin assay [58]

• 2. Titrant 1: 10 mM NaIO₄ in water (see Note 7).

• 3. Titrant 2: 15 mM l-adrenaline hydrochloride in water.

• 4. Enzyme in 50 mM aqueous borate buffer, pH 8.0 (see Note 8).

• 5. Microplates.

• 6. Plate reader.

2.5 Quick E Enantioselectivity Assay [75]

• 1. Assay buffer: 50 mM Tris–HCl, pH 8; 4.5 g/L Triton X-100

• 2. Enantiomeric substrate solution S: 7.8 mM (S)-p-nitrophenyl-2-phenylpropanoate dissolved in acetonitrile.

• 3. Enantiomeric substrate solution R: 7.8 mM (R)-p-nitrophenyl-2-phenylpropanoate dissolved in acetonitrile.

• 4. Reference substrate solution: 1.6 mM resorufin tetradecanoate dissolved in acetonitrile.

• 5. Microplates (96-well plates).

• 6. Plate reader (see Note 9).

3 Methods

3.1 Agar-Plate Assays

LB-agar is the most widely used solid medium for the growth of bacteria which can be supplemented with antibiotics to maintain expression plasmids and with inducers of gene expression, e.g., isopropyl-β-d-galactopyranoside (IPTG). One key factor with regard to these assays is how the expression of the genes is regulated by the medium components, e.g., the expression of the plcH and plcN are strongly repressed by phosphate levels in the growth media. Substrates should be freshly emulsified before adding to the agar medium to obtain the emulsion with optimal properties. It is recommended to use fresh agar plates to increase assays sensitivity and reproducibility (see Note 10). The assays need to be adapted to the biophysical properties of the studied enzymes by varying the incubation time (from 1 up to 4 days) and temperature (from 4 °C up to 50 °C) required for detection of an enzymatic activity. Not more than 500 clones per standard petri dish should be plated to achieve good resolution of clones and allow identification of single enzyme-producing clones.

3.1.1 Tributyrin Agar Plate Assay [43]

The most widespread plate assay to detect extracellular but also intracellular esterases and lipases uses tributyrin as the substrate. Clear zones appearing around the bacterial colonies indicate the production of a catalytic active enzyme (Fig. 2a).

Fig. 2

Lipase producing strain P. aeruginosa PAO1 and E. coli DH5α, a negative control, grown overnight on (a) tributyrin agar plate (Subheading 2.1.1); (b) rhodamine B agar plate (Subheading 2.1.2); and (c) egg yolk agar plates (Subheading 2.1.3). Agar plates appear opaque due to the presence of lipid substrate drops in the solidified medium. Lipolytic enzymes produced by P. aeruginosa hydrolyze the substrates resulting in clearance of the medium around P. aeruginosa colonies grown on tributyrin and egg yolk agar plates. The rhodamine B agar plate was photographed under UV light and lipolytic activity around P. aeruginosa colony is visible as light blue fluorescence

• 1. Add 30 mL of tributyrin emulsion, the respective antibiotic and inducer (e.g., IPTG) to 1 L of melted LB agar medium cooled to a temperature of about 60 °C and mix thoroughly.

• 2. Pour 20 mL medium into appropriate petri plates and let it solidify for at least 20 min.

• 3. Plate bacterial clones and incubate at optimal growth temperature for at least 16 h.

• 4. Positive clones are identified after overnight growth or after prolonged (2–4 days) incubation in refrigerator for clones expressing low amounts of active enzymes (see Note 11).

3.1.2 Rhodamine B Agar Plate Assay [45]

This assay, contrary to the tributyrin assay, is specific for true lipases because it uses triolein as a substrate, which is a triglyceride ester containing long-chain (C-18:1) fatty acids. Fluorescent complexes are formed between the cationic rhodamine B and free fatty acids released from the substrate by a lipase. Around lipase-positive clones, a fluorescent halo can be observed after irradiation of the plate with UV light at 350 nm (Fig. 2b). Clones which do not produce lipases appear pink colored, but nonfluorescent (see Note 12).

• 1. Add 31.25 mL of olive oil emulsion, 10 mL RB solution, respective antibiotic and inducer (e.g., IPTG) to 1 L of melted LB agar medium at a temperature of about 60 °C and mix thoroughly.

• 2. Pour 20 mL medium into appropriate plates and let it solidify. Normally, plates are pink colored and have opaque appearance.

• 3. Plate the clones and incubate at optimal growth temperature for at least 16 h.

• 4. Positive clones are identified under UV light (e.g., hand lamp at 350 nm) by fluorescent halos around colonies (see Note 13).

3.1.3 Egg Yolk Agar Plate Assay [46]

This assay is generally used to detect phospholipase A and C activity. The egg yolk substrate consists of about 65 % triacylglycerols, about 30 % phospholipids, and 5 % cholesterol. Hence, the egg yolk assay is applicable for the analysis of phospholipases and lipases, too [48]. The assay is based on the formation of calcium complexes with free fatty acids released by phospholipases A and lipases, indicated as white precipitation halos around the colonies (Fig. 2c). Although assays for PLC are not covered here, it is worth to mention that the egg yolk LB agar assay is applicable for assaying PLC activities, too. Phospholipase C producing clones show precipitation halos as a result of complex formation between diacylglycerol, released by action of PLC, and vitellin, the major egg yolk protein [46].

• 1. Add 200 mL of egg-yolk emulsion to 800 mL melted LB medium at temperature of about 60 °C and mix thoroughly.

• 2. Pour 20 mL medium into appropriate plates and let it solidify for at least 20 min.

• 3. Plate the clones and incubate at optimal growth temperature for at least 16 h.

• 4. Positive clones are identified by appearance of clear halos around colonies.

3.2 Titrimetric Assays [47, 48]

These assays are based on the quantitative titration of carboxylic acids released from various types of ester substrates by the hydrolytic activity of lipases or phospholipases A. Titrimetric methods can be classified in three groups: titration of free fatty acids after extraction, end point titration, and continuous titration. We describe here the continuous automatic titration, also called pH-stat method.

This is a common and reliable method used for the quantification of lipase/PLA activities based on continuous monitoring the amount of NaOH required to neutralize the liberated free carboxylic acids. The advantages of this method include its sensitivity (1 μmol/min of free fatty acids released) and the possibility to use a wide spectrum of naturally occurring lipids (mono-, di-, and tri-acylglycerols for measuring lipase/esterase activities; and phospholipids for PLA activities). Expensive synthetic (phospho)lipids (provided by for example Avanti polar lipids, Alabama, USA; Sigma Aldrich, Missouri, USA), or inexpensive natural (phospho)lipids such as olive oil (for lipases) or egg yolk (for phospholipases) can be used in titrimetric assays.

• 1. Turn on pH-stat device, flush the burette and fill it with titrator. Set up constant temperature to 37 ºC. Follow manufacturer’s instructions regarding maintaining of temperature and preparing the device prior starting measurements (see Note 14).

• 2. Add 10 mM pure triacylglycerol or phospholipid or 50 g/L natural oil or phospholipid in assay buffer. 5 mL of emulsion is usually enough for one reaction, although the volume depends on the reaction vessels available.

• 3. Homogenize the emulsion by using (a) sample sonicator for 4 min (two times 2 min, with cooling in between) at maximum capacity, (b) blender for 10 min (two times 5 min with cooling in between) at a speed of 11,000 min⁻¹, or (c) bath sonicator for 10 min (two times 5 min with cooling in between) at maximal capacity. Emulsions are usually stable for 4–6 h (see Note 15).

• 4. Add substrate emulsion in the reaction vessel and place the electrode, magnetic stirrer and delivery tip into the emulsion. Stir the emulsion at constant speed and adjust the pH of the substrate emulsion to 8 with titrant A. Set up the end point of the titrator to pH 8.0.

• 5. Start the reaction by adding a defined amount of enzyme solution (up to 250 μL per 5 mL of emulsion).

• 6. Titrate the liberated fatty acids continuously with titrant B, usually for 5–10 min.

• 7. Calculate the lipase activity per mg or mL of enzyme using Eq. 1 [47]. For calculation, use only the data with linear response of the titrant spent for acid neutralization with time. The unit of lipase activity is defined as the amount of enzyme releasing 1.0 μmol of fatty acid per 1 min at defined temperature and pH.

$$ {U}_{\mathrm{lipaseactivity}}/\mathrm{mgorml}=\frac{c\left(\mathrm{titrantB}\right)\left[\frac{\mathrm{mol}}{\mathrm{L}}\right]\times V\left(\mathrm{titrantB}\right)\left[\mathrm{mL}\right]\times 1,000}{t\left[ \min \right]\times \mathrm{amountofenzyme}\left[\mathrm{mgorml}\right]} $$

(1)

c = concentration of titrant B, 50 mmol/L NaOH

V = volume of titrant B

t = time of enzymatic reaction

amount of enzyme = amount of mg or mL of enzyme added to the reaction

3.3 Colorimetric and Fluorometric Assays

These methods are based on the hydrolysis of ester substrates that, upon hydrolysis by lipolytic enzymes, release chromogenic or fluorogenic products which can be detected by UV/VIS or fluorescence spectrophotometry, respectively. The artificial profluorogenic or chromogenic substrate analogues are suitable for routine and accurate quantification of lipase activity using standard equipment usually available in biochemical laboratories. Mostly, chromogenic p-nitrophenyl (p-NP) or α/β-naphtyl and fluorogenic 4-methylumbelliferyl (4-MU) or resorufin esters of fatty acids are used as substrates. However, because these substrates naturally do not occur in living cells, the assays are not suitable to define cellular functions of lipolytic enzymes. On the other hand, chromogenic or fluorogenic substrates can be used for high throughput screening of a large number of samples with fluorogenic methods usually being much more sensitive than colorimetric methods.

3.3.1 Assay with p-Nitrophenyl Acyl Ester Substrates [49–52]

This method is based on the quantification of p-nitrophenolate (p-NP) released during the hydrolysis of p-nitrophenyl acyl esters by lipolytic enzymes. The p-NP is chromogenic with an absorption maximum at 410 nm and it can be quantified continuously using a spectrophotometer. As substrates can be used p-nitrophenyl esters with saturated or unsaturated fatty acids differing in chain length. Usually, p-NP butyrate (C4) or caproate (C6) are used for esterase assays and p-NP palmitate (C16) for lipases assays. Because substrates, p-NP acyl esters, are not water soluble they are dissolved in organic solvents and then diluted with assay buffer. The selection of suitable organic solvent is driven by the solubility and stability of short-chain and long-chain fatty acid p-NP esters. Acetonitrile is the most commonly used solvent for short-chain esters and isopropanol for long-chain esters. However, acetonitrile can also be used for assays with p-NP esters in the entire range of C2–C18 fatty acids. We present here two protocols, one for determination of esterase activity using p-NPC4 (also applicable for substrates with fatty acids up to 18 carbon atoms) and another for determination of lipase activity using p-NPC16. The major disadvantage of p-NP acyl ester substrates is their spontaneous hydrolysis under assay conditions. The activity of a purified enzyme should be assayed at the respective optimal temperature, pH, and buffer. Also 5 mM divalent cations (e.g., Ca²⁺ or Zn²⁺) should be added in the buffers if it is required for activity or stability of an enzyme.

Similar to the assays that rely on hydrolysis of p-NP esters, organophosphonate esters of p-NP were used for determination of concentration of catalytically active lipases in solution by active-site titration methods. Standard methods for protein quantification with Bradford reagent or UV spectroscopic measurements do not provide data about concentration of enzymatically active enzyme required for kinetic and structural studies. In the active-site titration experiments, lipase is incubated with an irreversible inhibitor (e.g., methyl p-nitrophenyl n-hexylphosphonate [59]) that specifically binds to the active site serine yielding the chromophore p-nitrophenolate that is quantified spectrometrically. Optionally, 3,4-dichloroisocoumarin may be used as an inhibitor followed by measurement of residual lipase activity using p-NP assay [60].

• 1. Dilute substrate stock solution 20-fold with the assay buffer and vortex for 2–3 min.

• 2. Pipette enzyme sample into reaction tubes (5–50 μL) or in microplates (1–20 μL). Prepare the blank (non-enzymatic control) by adding buffer in which the enzyme is dissolved to the reaction tube or microplate well.

• 3. Set up the temperature of VIS spectrometer at 30 ºC (see Note 16).

• 4. (A) Add substrate solution into reaction tubes (0.5–2 mL) to initiate the reactions. Start the timer, mix and transfer the solution from reaction tube into the cuvette and record absorbance at 410 nm (A _410nm) over the time course of 2–10 min.

  (B) Fill the microplate with the substrate (100–200 μL) (see Note 17) and place it into VIS microplate spectrophotometer (plate reader). Set up the 5 s mixing step prior measurements start and record A _410nm each 10–30 s over the time course of 10–15 min.

• 5. Prepare at least six standard solution of p-NP by diluting standard stock solution of p-NP in the assay buffer and by adding isopropanol (for p-NPC16 assay) or acetonitrile (for p-NPC4 assay) to yield final concentration of 5 % (v/v). Measure the A _410nm of p-NP dilutions in a range of 0.01–0.2 mM under the assay conditions. Prepare the standard curve by plotting the A _410nm versus the p-NP concentration.

• 6. Subtract the fluorescence of the blank from the measured values for the enzyme samples and convert the A _410nm into enzyme activity using the standard curve. Optionally, if the extinction coefficient is known, the absorbance measured may be used to calculate units of lipase activity (U) according to Eq. 2 (see Note 18).

$$ {U}_{\mathrm{lipaseactivity}}/\mathrm{ml}=\frac{\left({A}_{410\mathrm{nm}}\left({t}_1\right)-{A}_{410\mathrm{nm}}\left({t}_0\right)\right)\times {V}_{\mathrm{total}}}{\varepsilon \times {V}_{\mathrm{enzyme}}\times \left({t}_1-{t}_0\right)\times l} $$

(2)

A _410nm(t ₁) = absorbance at 410 nm measured after enzymatic reaction is finished

A _410nm(t ₀) = absorbance at 410 nm measured before enzymatic reaction is started

t ₁ = time point when reaction is finished

t ₀ = time point when reaction is started, usually 0 min

V _total = volume of reaction sample plus enzyme sample (V _enzyme)

ε = molar extinction coefficient [mmol⁻¹ dm³ cm⁻¹]

l = light path length [cm]

3.3.2 Assay with 4-Methylumbelliferyl Acyl Ester Substrates [53, 54]

Hydrolysis of 4-MU acyl esters by lipases can be followed continuously by monitoring the increase of fluorescence intensity due to the production of highly fluorescent 4-methylumbelliferone with an emission spectra maximum at 460 nm. In contrast to emission maxima, the fluorescence excitation maxima of 4-MU is pH dependent and varies from 330 nm at pH 4.6 to 385 nm at pH 10.4 [54]. Esters of 4-MU with different fatty acids (butyric, heptanoic, oleic acid) are commercially available. Short-chain 4-MU butyrate is used for assaying esterases, long-chain 4-MU-oleate is used for true lipases, and 4-MU heptanoate is suitable for both lipases and esterases. For activity calculations, it is necessary to correct for spontaneous hydrolysis of the substrates and for background fluorescence of the protein sample. Enzyme activity is defined as the amount of released 4-MU per minute, and can be calculated with a standard curve obtained by measuring the fluorescence of 4-MU under assay conditions.

• 1. Dilute substrate stock solution 100-fold into the assay buffer and vortex for 2–3 min.

• 2. Pipette enzyme sample into reaction tubes (5–50 μL) or in microplates (1–20 μL). Prepare the blank (non-enzymatic control) by adding the buffer in which enzyme is dissolved to the reaction tube or microplate. Prepare one reaction tube or microplate well with the enzyme solution for measurement of intrinsic enzyme fluorescence.

• 3. Set up the temperature of spectrofluorometer at 30 ºC.

• 4. (A) Add substrate solution into reaction tubes (0.5–2 mL) to initiate the reactions. Start the timer, mix and transfer the solution from the reaction tube into the cuvette and record fluorescence at 460 nm over a time course of 10–15 min.

  (B) Fill the microtiter plate with the substrate (100–200 μL) (see Note 17) and place it into fluorescence plate reader. Set up the 5 s mixing step prior measurements start and record the fluorescence at 460 nm each 10–30 s over a time course of 10–15 min.

• 5. Add the assay buffer only (without the substrate) to the reaction tube or microplate well with the enzyme sample and measure the intrinsic fluorescence of the enzyme.

• 6. Prepare at least six standard solution of 4-MU by diluting standard solution of 4-MU in the assay buffer to yield dilutions in a range 50–500 nM. Measure the fluorescence under the assay conditions. Prepare the standard curve by plotting the fluorescence versus the 4-MU concentration.

• 7. Subtract the fluorescence of the blank and intrinsic enzyme fluorescence from the sample fluorescence and convert the fluorescence into the enzyme activity using the standard curve.

3.3.3 Assay with Natural (Phospho)Lipid Substrates [55, 56]

Lipase, esterase, phospholipases A and lysophospholipase A activities can be determined with (phospho)lipids naturally occurring in living cells. Here, the ASC-ACOD-MEHA enzymatic-coupled method for the determination of released fatty acids can be used. Free fatty acids liberated upon enzymatic hydrolysis of natural (phospho)lipid substrates are determined colorimetrically by means of the NEFA-HR(2) kit according to the manufacturer’s instructions. This method relies upon the synthesis of acyl-CoA from free fatty acid and coenzyme A in the presence of acyl-CoA synthetase (ACS). The acyl-CoA is subsequently oxidized by acyl-CoA oxidase (ACOD) to yield enoyl-CoA resulting in release of H₂O₂ as the side product. A peroxidase then uses this H₂O₂ to oxidize 4-aminoantipyrin (4-AA); which in its oxidized form reacts with 3-methyl-N-ethyl-N-(β-hydroxyethyl)-aniline (MEHA) to yield a quinoneimine which is purple colored and can be quantified spectrometrically at 550 nm. The concentration of non-esterified fatty acids (NEFA) can be calculated from the A _550nm by using the calibration curve. This enzymatic ASC-ACOD-MEHA method gains rapidly increasing interest for measurement of lipolytic activities because it is applicable for all ester substrate available, it is reliable even in the presence of numerous interfering compounds, it is simple, sensitive and requires only low amounts of substrates (in the μg range per reaction).

• 1. Dilute lipid substrate 20-fold with assay buffer by vortexing for 15 min at 37 ºC and then subject to ultrasonication three times for 20 s.

• 2. Combine 25 μL of lipid substrate with 25 μL of enzyme sample in reaction tubes or microplates, tightly close reaction vessels and incubate at 37 ºC in agitation mode (see Note 19). The incubation time, usually 30 min for very active samples and overnight for less active samples, is dependent on the activity of the analyzed samples and therefore it should be empirically estimated in preliminary experiments.

• 3. Prepare respective blank controls by treating them the same like enzyme samples.

• 4. Prepare at least six dilutions of oleic acid in the range of 0.1–1 mM in assay buffer (see Note 20). A standard curve may also be determined by using a commercially available calibrator kit (Wako Chemicals).

• 5. Prepare reagent solution R1 according to the supplier instructions by dissolving color reagent A in solvent A (see Note 21).

• 6. Prepare reagent solution R2 according to the supplier instructions by dissolving color reagent B in solvent B (see Note 21).

• 7. Pipette 2–10 μL of the reaction samples into new reaction tube or microplate and add reagent solution R1 (300 or 100 μL for assays in reaction tubes or microplates, respectively) followed by agitation for 5 min at 37 °C.

• 8. Add reagent B to the samples (150 or 50 μL for assays in reaction tubes or microplates, respectively), agitate at 30 °C for 5 min and measure absorbance at 550 nm.

• 9. Subtract the absorbance of the blank samples from the enzyme samples and convert the A _550nm into the enzyme activity using the standard curve.

3.4 Adrenalin Fingerprinting Assay [57]

The simultaneous measurement of a single enzyme activity toward a range of different substrates provides a data set sufficient for a precise functional definition of a given enzyme, a so-called enzyme activity fingerprint. Because only small variations in substrate structure may significantly influence the enzyme activity, fingerprinting methods are confident for functional characterization of a single enzyme. The fingerprinting data are usually represented as images created by linking each measured data with one pixel which color intensity represents the activity (for example from white corresponding to 0 % to black corresponding to 100 % activity) (see Fig. 3). The assay requirements for such fingerprinting methods include applicability with usually tens of different substrates, high reproducibility and use of the same enzyme solution. These methods are valuable for definition of substrate promiscuity, identification of biological function, quality control, and medical diagnostics.

Fig. 3

The principle of the fingerprint method. The same enzyme sample is assayed under identical conditions with, in this case, 24 different substrates (S1–S24) resulting in an enzyme fingerprint

Although several fingerprinting methods for assaying lipase activities were developed with fluorogenic [61] and chromogenic [62] substrates, the limited range of commercially available tagged substrates hampers the creation of large substrate libraries for fingerprinting. To overcome the rather laborious synthesis of fluorogenic or chromogenic substrate libraries it is possible to use indirect assays with a range of commercially available esters substrates. In these assays, the products released by the hydrolytic activity of lipases a) may initiate a sequence of chemical reactions that result in generation of chromogenic or fluorogenic compounds (e.g., adrenalin assay) or b) may change the color of a pH indicator by changing the pH of the reaction. The amount of hydrolytic product may be indirectly quantified spectrophotometrically or fluorometrically.

Here, we describe the adrenalin assay [57], a fingerprinting method used for profiling substrate specificity of different lipases and esterases. This indirect fingerprinting assay uses 35 different commercially available polyol acetates substrates (Table 1) from which 1,2-diols are released upon enzymatic hydrolysis. The formed diols are oxidized with periodate and the remaining periodate is back-titrated with adrenaline to give a red colored adrenochrome product with an absorption maximum at 490 nm. These end point spectrometric measurements can be used for generating fingerprinting images.

• 1. Mix 10 μL substrate with 10 μL NaIO₄ and 80 μL of enzyme solution and incubate for 60 min at 37 °C.

• 2. Add 10 μL adrenalin solution and incubate for 5 min at 26 °C.

• 3. Measure the absorbance at 490 nm.

3.5 Quick E Enantioselectivity Assay [75]

The ability of many lipolytic enzymes to distinguish between two enantiomers of a given racemic substrate, designated as enantioselectivity, is a key feature for their industrial applications [13–15, 17–19]. The increasing demand for the synthesis of enantiomerically pure compounds by the pharmaceutical industries is driven by the fact that very often only one enantiomer has a desired biological activity, whereas the other enantiomer may cause undesirable side effects [63, 64]. Although several lipases often exhibit enantioselectivity, mostly, it is necessary to improve it for a specific substrate. For this purpose, molecular genetic methods have been developed and applied including rational protein design [65], random mutagenesis, saturation mutagenesis, iterative saturation mutagenesis [66], and DNA shuffling to increase the enantioselectivity of lipases [67]. Notably, the lipase LipA from P. aeruginosa represents the best studied enzyme with respect to directed evolution of enantioselectivity [68–74]. Initially, proof-of principle was demonstrated by increasing the enantioselectivity of LipA in the kinetic resolution of the substrate 2-methyldecanoic acid nitrophenyl ester to E = 25,8 (the wild-type lipase is not selective at all showing an E = 1,1) [72]. Later, diverse directed evolution methods were developed and optimized using this lipase. Finally, an enantioselectivity of E = 106 was achieved for the hydrolysis of 2-phenylbutyric acid p-nitrophenyl esters (the wild-type lipase has E = 2) [66].

The enantioselectivity is usually expressed as enantiomeric ratio (E) defined as the ratio of the initial rate of reaction with the preferred enantiomer (fast enantiomer) and the initial rate of reaction with the second enantiomer (slow enantiomer) (Eq. 3) [76]. The true enantiomeric ratio can be obtained by measuring the initial rates of reactions of each enantiomer in the presence of both enantiomers, under so-called competitive conditions. This limits the choice of detection methods to time consuming methods like for example HPLC or GC. Therefore, colorimetric methods were developed allowing screening of large libraries for enantioselective enzymes. Here, enzyme activities are determined with enantiomerically pure (S)- and (R)- substrates separately, e.g., (S)- and (R)-p-NP 2-methyl decanoate [77] or (S)- and (R)-p-NP-2-phenylpropanoate [78]. It should be noted that these assays reveal only estimated E-values that can significantly deviate from true E-values as shown for Candida antarctica lipase A (true E = 1,4 and estimated E = 4 for hydrolysis of (R)- and (S)-p-NP-2-phenylpropanoate [75]).

$$ \mathrm{Enantiomericratio}=E=\frac{\left(\frac{k_{\mathrm{cat}}}{K_{\mathrm{m}}}\right)\mathrm{fastenantiomer}}{\left(\frac{k_{\mathrm{cat}}}{K_{\mathrm{m}}}\right)\mathrm{slowenantiomer}}=\frac{\mathrm{reactionrate}\left(\mathrm{fastenantiomer}\right)}{\mathrm{reactionrate}\left(\mathrm{slowenantiomer}\right)} $$

(3)

Here, we describe a method for the fast determination of enantioselectivities, termed Quick E method. It was developed to overcome the lack of competition between two enantiomers. It is based on the hydrolysis of each enantiomerically pure substrate, in the presence of a reference substrate that must not be enantiomerically pure [78] but both these substrates must be detectable simultaneously in one solution. The colorimetric assays with pure (S)- and (R)-p-NP-2-phenylpropanoate, and resorufin tetradecanoate as a reference substrate, were performed with several lipases [78]. The p-nitrophenolate and resorufin can be detected spectrophotometrically at 404 and 572 nm, respectively, thus providing initial rates of hydrolysis of each enantiomer. Principally, by performing two measurements, one with a mixture of (S)-p-NP-MD and RT and the second one with (R)-p-NP-MD and RT, the enantioselectivity for both reactions can be calculated using Eqs. 4 and 5 [75]. By dividing these two enantio selectivities, a Quick E-value can be calculated (Eq. 6 [75]). Such simulated competition between enantiomeric and reference substrates results in better agreement of Quick E-values with true E-values than estimated E-values with true E-values, as demonstrated e-.g. for lipases of Pseudomonas cepacia (true and Quick E-values = 29) and Candida rugosa (true and Quick E-values = 3,5) [75].

$$ E=\frac{(R)\ \mathrm{substrate}}{\mathrm{reference}}\mathrm{selectivity}=\frac{\frac{ reaction\ rate\left(R\ enantiomer\;\right)}{\mathrm{concentration}\left(R\ enantiomer\;\right)}}{\frac{ reaction\ rate\left(\mathrm{reference}\ substrate\;\right)}{\mathrm{concentration}\left(\mathrm{reference}\ substrate\right)}} $$

(4)

$$ E=\frac{(S)\ \mathrm{substrate}}{\mathrm{reference}}\mathrm{selectivity}=\frac{\frac{ reaction\ rate\left(S\ enantiomer\;\right)}{\mathrm{concentration}\left(S\ enantiomer\;\right)}}{\frac{ reaction\ rate\left(\mathrm{reference}\ substrate\;\right)}{\mathrm{concentration}\left(\mathrm{reference}\ substrate\right)}} $$

(5)

$$ {E}_{\mathrm{Quick}}=\frac{E1}{E2}=\frac{\frac{(R)-\mathrm{substrate}}{\mathrm{reference}}\mathrm{selectivity}}{\frac{(S)-\mathrm{substrate}}{\mathrm{reference}}}=\frac{(R)-\mathrm{substrate}}{(S)-\mathrm{substrate}}\mathrm{selectivity} $$

(6)

• 1. Add 0.5 mL of (S)-substrate solution and 0.5 mL of reference substrate solution into 9 mL of assay buffer drop wise and vortex 2–3 min until emulsion is clear (see Note 22).

• 2. Add 0.5 mL of (R)-substrate solution and 0.5 mL of reference substrate solution into 9 mL of assay buffer drop wise and vortex 2–3 min until emulsion is clear.

• 3. Add 20 μL enzyme solution to the 180 μL substrate emulsion (containing (S)- and reference substrates) in microplate and measure the increase in absorbance at 404 and 572 nm at 25 °C for 5–30 min with steps of 15 s.

• 4. Add 20 μL enzyme solution to the 180 μL substrate emulsion (containing (R)- and reference substrates) in microplate and measure the increase in absorbance at 404 and 572 nm at 25 °C for 5–30 min with steps of 15 s.

• 5. Measure blank and substrate only samples the same as the enzyme samples (see Note 23).

• 6. Values are in absorbance per second × 1,000. Enantiomeric ratio for (R)- and (S)- enantiomer separately is calculated using Eqs. 4 and 5, respectively. Quick E enantiomeric ratio is calculated using Eq. 6.

4 Notes

• 1. pH of buffers may change with temperature; hence, for measuring temperature optima, use buffers having their pH adjusted accordingly.

• 2. The substrates with different acyl chain length, number of double bonds in the acyl chain and absence, or presence, of phosphoester moiety bound on the glycerol may be chosen for this assay dependent on the available information about the substrate specificity of tested enzyme.

• 3. pH 8 is required because most free fatty acids have a pKa lower than 8. Also, optimal pH and stability of lipases is usually in the range of pH 7–9.

• 4. Black microplates have low background fluorescence and minimal light scatter, therefore they are recommended for fluorometric assays.

• 5. Optionally, 4-MU heptanoate may be dissolved in ethylene glycol monomethyl ether [54].

• 6. Mono-, di-, and tri-acylglycerol, phosphatidylglycerol, phosphatidylcholine, phosphatidyl-ethanolamine, lysophosphatidylcholine lysophosphatidylglycerol. and lysophosphatidyl-ethanolamine are usually used as (phospho)lipase substrates.

• 7. NaIO₄ solution in water should be prepared freshly.

• 8. Sodium phosphate buffer may be used depending on optimal stability and activity of studied enzyme. Do not use buffer that is oxidized with NaIO₄, e.g., Tris or triethanolamine buffers.

• 9. It is preferable to use a plate reader allowing the simultaneous measurement of absorbance at two different wavelengths.

• 10. The spontaneous hydrolysis of substrates can result in its decreased concentration upon long term storage of plates.

• 11. To increase the probability to find lipase-producing clones in gene libraries one can prepare several tributyrin agar plate replicas from one master plate after overnight incubation. Each replica plate may then be incubated at a different temperature, for example 4, 37 and 50 °C in order to detect lipases with different temperature optima.

• 12. Bacteria accumulate intracellularly pink colored rhodamine B which is not fluorescent in absence of free fatty acids.

• 13. To prevent damaging effects of UV light apply UV radiation only for a short time (a few sec) if you want to propagate the clones from the same plate afterwards.

• 14. The temperature should be chosen based on the stability and optimal temperature of the enzyme.

• 15. The reproducibility of assays depends strongly on the emulsion properties, therefore, the same homogenization conditions should always be used.

• 16. The assay can be performed at temperatures up to 80–90 ºC, although the spontaneous hydrolysis of the substrate at higher temperatures is significant.

• 17. Preferably use multichannel pipette to minimize time needed to fill microplate.

• 18. The molar extinction coefficient changes with temperature and pH.

• 19. Already 2 μL of reaction samples is sufficient to measure activities.

• 20. If you are using (phospho)lipid substrates with different fatty acid chain lengths than oleic acid, it is recommended to determine a calibration curve with the same fatty acid attached to the ester substrate. Fatty acids with 6 up to 18 carbon atoms can be detected with this method.

• 21. Preferably prepare and use the solution fresh or not older than 1 month when stored at 8 ºC. Due to instability of the enzymes and ATP present in the solutions R1 and R2, older reagents may give false results.

• 22. So prepared emulsion remained stable for 2–3 h.

• 23. Substrate only sample should not show any increase of absorbance during 30 min.