Proteins and Enzymes

Chapter 8

Proteins and Enzymes 8.1. INTRODUCTION Because of its importance in many analytical, industrial and medical applications, the behaviour of proteins and enzymes at interfaces has attracted the attention of workers from various fields and with diverse interests. This is attested by the numerous reports and reviews that have appeared at intervals over the past five decades (Boyd and Mortland, 1990; Brash and Lyman, 1971; Burns, 1986; Burns and Dick, 2002; Fenstermaker et al., 1974; Fraser, 1957; Gray, 2004; Haynes and Norde, 1994; Loeb, 1965; Lynch and Dawson, 2008; McLaren and Packer, 1970; Nakanishi et al., 2001; Norde, 1986; Quiquampoix, 2000; Quiquampoix and Burns, 2007; Wahlgren and Arnebrant, 1991; Zimmerman and Ahn, 2010). A classic example is the separation and purification of proteins and enzymes by adsorption to inorganic and organic supports (Zittle, 1953), setting the scene to ‘enzyme immobilization’ by attachment to, and entrapment in, solid (mineral) and gel matrices (Bornscheuer, 2003; De Boeck, 1975; Hudson et al., 2005; Katchalski-Katzir, 1993; Kim et al., 2006; Messing, 1975; Mosbach, 1976, 1987; Rouxhet, 1990; Sedaghat et al., 2009a; Sheldon, 2007; Shen et al., 2002; Thomas and Kernevez, 1976; Tischer and Kasche, 1999; Unsworth et al., 2007; Zaborsky, 1973). By this means, both the activity and stability of enzymes can be controlled and modified to meet specific requirements, providing the basis for ‘enzyme engineering’ (Gray, 2004; Okada et al., 1990; Svendsen, 2004; Weetall and Suzuki, 1975; Wingard, 1972). Equally relevant is the ability of adsorbed proteins to alter the surface properties of the adsorbent. In such seemingly unrelated phenomena as blood clotting, dental plaque formation and marine fouling, for example, the adhesion of arriving cells to the solid substrate is markedly influenced by the presence of a pre-adsorbed layer of protein at the surface (Baier, 1975; Burns and Holmberg, 1996; Fletcher et al., 1980; Gray, 2004; Milleding et al., 1999; Norde and Lyklema, 1989). More recent developments in the clay–protein interaction include the behaviour of insecticidal proteins from Bacillus thuringiensis at clay mineral surfaces (Helassa et al., 2009; Tapp and Stotzky, 1995; Venkateswerlu and Stotzky, 1992; Zhou et al., 2005), the fate of pathogenic proteins (prions) in soil (Cooke and Shaw, 2007; Johnson et al., 2006; Leita et al., 2006; Quiquampoix and Burns, 2007; Rigou et al., 2006) and the Developments in Clay Science, Vol. 4. DOI: 10.1016/B978-0-444-53354-8.00008-6 # 2012 Elsevier B.V. All rights reserved.

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layer-by-layer construction of clay–protein films for electroanalytical (biosensing) and catalytic applications (Li and Hu, 2003; Liu et al., 2005; Lojou and Bianco, 2006; Lvov et al., 1995; Mousty, 2004, 2010; Patil et al., 2005; Szabo´ et al., 2007; Zhou et al., 2002a,b). From a soil science perspective, it is worth recalling that nearly all of the nitrogen in soil is organically bound (Bremner, 1965, 1967; Parsons and Tinsley, 1975). A large proportion (up to 50%) of this nitrogen can be released as a-amino acids on acid hydrolysis of whole soils or their organic matter extracts (Friedel and Scheller, 2002; Rillig et al., 2007). Although the form in which these amino acids occur is still a matter of some debate, there is good evidence to indicate that they are primarily derived from peptide-like structures (Knicker et al., 1993, 2000; Ladd and Butler, 1975). The inherent resistance to microbial decomposition of soil organic nitrogen is consistent with the observation that only a small proportion (<5%) of this nitrogen can be converted into inorganic forms (‘mineralized’) during the growing season (Bartholomew, 1965; Bremner, 1967). The persistence of proteins in soil has been ascribed to complex formation with tannins, lignins, carbohydrates and humic substances. The extensive literature on this topic has recently been reviewed by Rillig et al. (2007). Alternatively, proteinaceous compounds may be stabilized by interaction with the mineral constituents of soil. Because of its extensive surface area and reactivity, the clay fraction plays a dominant role in this process (Boyd and Mortland, 1990; Chevallier et al., 2003; Kleber et al., 2007; Nielsen et al., 2006; Yuan and Theng, 2011). Here we are primarily concerned with the formation and properties of complexes of clay minerals with (globular) proteins and enzymes. Apart from its intrinsic value, the clay–protein interaction is of great interest to agronomists, soil scientists and biogeochemists alike because of its profound influence on the biostability and persistence in soil and sediment of amino acid polymers and humic substances, in general (Bremner, 1965, 1967; Curry et al., 1994; Goh, 1972; Hedges and Keil, 1995; McLaren and Peterson, 1965; Riboulleau et al., 2002; Scharpenseel, 1971; Skujin¸sˇ and McLaren, 1968, 1969; Zang et al., 2000; Zimmerman and Ahn, 2010). Direct evidence for the occurrence of proteins in soil is not easy to obtain because of practical difficulties in their extraction and isolation from the soil matrix (Nielsen et al., 2006). A notable exception is that of glomalins, a class of glycoproteins secreted by arbuscular mycorrhizal fungi. Because of their abundance and persistence in soil, these compounds can be extracted with relative ease (Wright and Upadhyaya, 1996). As already remarked on, the release of amino acids on acid hydrolysis is indicative of the presence in soil of clay–protein complexes. Additional evidence is provided by the appearance of an intense amide signal in the 15N-NMR spectra of whole soils (Friedel and Scheller, 2002; Knicker et al., 1993; Rillig et al., 2007). On the other hand, complexes of clay minerals with proteins can readily be prepared in the laboratory. Accordingly, information on the clay–protein

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interaction has largely derived from a systematic examination of the factors affecting the formation of such ‘synthetic’ complexes. Nevertheless, the data obtained from laboratory measurements are valuable in that they provide insight into the fate of proteins when they come into contact with clay and mineral colloids in the soil environment (Theng and Yuan, 2008). Unlike the situation with proteins, the occurrence of enzymes in soil has been amply documented (Burns, 1978, 1982; Durand, 1965; Gianfreda and Ruggiero, 2006; Hofmann and Hoffmann, 1966; Kiss et al., 1975; Shukla and Varma, 2010; Skujin¸sˇ, 1967; Tabatabai and Dick, 2002). Indeed, a number of enzymes, either as such or in combination with humic substances, have been successfully extracted from soils, leaving little doubt that complex formation with soil colloids does not generally lead to enzyme inactivation (Burns et al., 1972a,b; Ceccanti et al., 1978; Ladd and Butler, 1975; Marx et al., 2005; McLaren et al., 1975; Nannipieri et al., 1982, 2002; Ruggiero et al., 1996). Rather, complex formation tends to protect soil enzymes from microbial attack. What is readily measurable, of course, is the activity of a given enzyme acting on its substrate under some specified conditions, rather than its (absolute) quantity (Burns, 1982; Gianfreda and Ruggiero, 2006; Nielsen et al., 2006). We should also mention that enzyme reactions in soil occur at solid/liquid or solid/gel interfaces rather than in homogeneous, aqueous solutions (Burns, 1986; Fraser, 1957; McLaren and Packer, 1970; Nannipieri and Gianfreda, 1999; Quiquampoix and Burns, 2007). Thus, soil enzymes express their activity within an organized structural framework where surface effects may be as important as, or even take precedence over, ordinary mass–action relationships (Katchalski et al., 1971; McLaren, 1960; McLaren and Babcock, 1959). By forming complexes with extracellular enzymes, both clay minerals and humic substances provide such a framework in the soil environment (Burns, 1986, 1990; Ladd and Butler, 1975; Skujin¸sˇ, 1967, 1976). Since the early work by Mattson (1932) and Ensminger and Gieseking (1939, 1941, 1942), a large amount of information has accumulated on the clay–protein interaction. Although the different data sets are broadly concordant, they do not always agree in detail. On reflection, this is hardly surprising because complex formation between clay minerals and proteins is influenced by many variables. Even the manner in which the reactants are brought together may affect the properties of the resultant complexes. For these reasons, the reactions of clay minerals with proteins can at best be described in semi-quantitative terms.

8.2. GENERAL ASPECTS OF THE CLAY–PROTEIN INTERACTION Protein molecules are organized into several levels of structural complexity: (a) a primary structure denoting the sequence of amino acids linked through peptide bonds in a polypeptide chain; (b) a secondary structure arising from the hydrogen bond-induced coiling of this chain into a more stable conformation, such as the a-helix or (pleated) b-sheet and (c) a tertiary structure

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Formation and Properties of Clay-Polymer Complexes

describing the folding of this helix into a compact, globular molecule stabilized by interfold hydrogen bonding, van der Waals forces and hydrophobic interactions. Disulphide links in both secondary and tertiary structures may further contribute to holding the molecule in a particular conformation (Creighton, 1993; Teegarden, 2004). Such a structural organization imposes severe internal constraints thereby limiting the extent to which the molecule can rearrange itself at the clay mineral surface. A partial breakdown of the secondary structure with a reduction in a-helix or b-sheet content would increase the conformational entropy of the protein and hence favour adsorption. In this connection, we may also distinguish between ‘hard’ proteins having a strong internal coherence (stability) and ‘soft’ proteins that display a weak internal coherence (stability). As a rule of thumb, ‘hard’ proteins would only adsorb to hydrophilic surfaces (e.g. montmorillonite) if they are electrostatically attracted. ‘Soft’ proteins, on the other hand, can attach to hydrophilic surfaces carrying charges of the same sign because the accompanying structural changes give rise to an increase in conformational entropy. Both ‘hard’ and ‘soft’ proteins, however, can interact with hydrophobic surfaces (e.g. talc, organically modified clay minerals), irrespective of the charge characteristics of either protein or sorbent, induced by surface dehydration. These points are summarized in Table 8.1. Proteins are polyampholytes in that the molecules may either be positively or negatively charged depending on whether the ambient solution pH lies on the acid or the alkaline side of the isoelectric point. At this point, the sum of positive charges arising from protonation of basic (

NH2) side groups is equal to that of negative charges due to dissociation of acidic (

COOH) side groups in the primary (polypeptide chain) structure. Protein molecules also contain apolar (amino acid) groups. Because of their tendency to expel water molecules, these groups reside in the interior of the globular structure, making up the densely packed hydrophobic core. On adsorption, some of the internal (‘buried’) hydrophobic groups may come into contact with the surface, although they remain shielded from the aqueous phase (Kim et al., 2002; Norde, 2008). Similarly, the charge characteristics of clay mineral surfaces are pH dependent. As indicated in Chapter 1, this dependency is pronounced with the kaolinite group of minerals where the amphoteric edge surface makes up an appreciable proportion of the total particle area (cf. Figure 1.3), and more so with allophane where amphoteric (OH)Al(OH2) groups are exposed at perforations in the spherule wall (cf. Figure 1.18). On the other hand, the charge characteristics of the smectites are relatively insensitive to variations in solution pH since the dominant (negative) basal surface charge arises from isomorphous substitution within the layer structure (cf. Figure 1.9). Because of their peculiar molecular structure, globular shape, and pHdependent charge characteristics, proteins can associate with clay and mineral

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TABLE 8.1 Predictive Scheme of Protein Adsorption. Internal Cohesion

Protein

Strong (‘hard’)

Weak (‘soft’)

Electrical charge

Sorbent Surface Hydrophobic

Hydrophilic

Electrical Charge

Electrical Charge

þ




þ




þ

Yes

Yes

No

Yes




Yes

Yes

Yes

No

þ

Yes

Yes

Yes

Yes




Yes

Yes

Yes

Yes

Hydrophobic dehydration dominates adsorption

Electrical interactions or structural changes in proteins dominate adsorption

Positive and negative signs refer to the charges on the protein or surface. Conditions at which adsorption is predicted to occur are indicated by ‘Yes’, while conditions predicting the absence of adsorption are marked ‘No’. From Norde (2008) and Norde et al. (2008).

surfaces through both enthalpic (electrostatic and van der Waals forces) and entropic (hydrophobic and conformational) interactions. Electrostatic interactions between solute and surface are generally dominant, although van der Waals (‘dispersion’) forces together with hydrophobic interactions between amino acids, inside the protein secondary structure, and surface (siloxane) sites can make a significant contribution to the overall adsorption energy (Lynch and Dawson, 2008; Norde, 2008; Quiquampoix, 2008a,b; Quiquampoix and Burns, 2007; Roth and Lenhoff, 1995). Since protein molecules can establish numerous points of contact with the solid surface, the isotherms for protein adsorption are often of the H-type (cf. Figure 2.2). Further, little desorption occurs on dilution because the likelihood of all solute–surface bonds breaking simultaneously is very low (Nakanishi et al., 2001; Norde, 2008; Wahlgren and Arnebrant, 1991). However, the train-loop-tail surface conformation adopted by linear, flexible polymers (cf. Figure 2.1; Chapter 3) does not generally obtain with globular proteins. Fenstermaker et al. (1974) and Morrissey and Stromberg (1974), for example, found a bound fraction of 0.11 for bovine serum albumin (BSA) and prothrombin adsorbing to silica, irrespective of the amount adsorbed. This value was only 1/5 to 1/3 of that predicted and commonly observed for flexible polymers (Greenland, 1972; cf. Chapter 2). In other words, adsorption does not usually lead to extensive unfolding of the polypeptide chain, and the

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Formation and Properties of Clay-Polymer Complexes

thickness of adsorbed proteins is often comparable with the dimension of their counterparts in solution (Brash and Lyman, 1971; Norde, 2008) Adsorption to mineral surfaces, however, may induce a partial breakdown (rearrangement) of the ordered (secondary) structure, giving rise to an increase in conformational entropy (Haynes and Norde, 1994; Norde, 1986, 2008; Quiquampoix, 2008a,b). Expanding 2:1 type layer silicates, of which montmorillonite is the outstanding example, are capable of intercalating a large variety of proteins. When this happens, the thickness and probable conformation of the intercalated species may be deduced from adsorption and X-ray diffraction data. In the absence of intercalation, or when interlayer penetration is restricted for steric reasons, these parameters may be estimated by ellipsometry, viscometry and various spectroscopic techniques such as circular dichroism, fluorescence, infrared (IR) and nuclear magnetic resonance (Andrade and Hlady, 1986; Gray, 2004; Lynch and Dawson, 2008; Norde, 2008; Quiquampoix, 2008a, b; Wahlgren and Arnebrant, 1991). Ellipsometry together with total internal reflectance fluorescence and NMR spectroscopy have been useful in assessing the thickness and density of polymers and proteins at solid/liquid interfaces but have not been developed to full potential to studying the properties of clay-adsorbed proteins (Andrade and Hlady, 1986; Beaglehole et al., 1998; Berg et al., 2001; Fenstermaker et al., 1974; Gray, 2004; Malmsten, 1995; Nakanishi et al., 2001; Stromberg et al., 1970; Svendsen et al., 2008; Vroman and Adams, 1969). The same can be said for viscometry and related rheological techniques (Garvey et al., 1976; Heath and Tadros, 1983; Liang et al., 1995; Srivastava and Chauhan, 1975). Circular dichroism has been used to good effect in probing the structure of proteins adsorbed to nanosize solids where light scattering effects do not interfere with the measurement (Billsten et al., 1995; Kondo et al., 1991; Maste et al., 1997; Nakanishi et al., 2001; Norde and Favier, 1992; Vertegel et al., 2004). IR spectroscopy, however, remains the single, most widely used technique for assessing the conformation of adsorbed proteins at clay mineral surfaces (Baron et al., 1999; Fusi et al., 1989; Noinville et al., 2002; Quiquampoix, 2008a; Servagent-Noinville et al., 2000; Tarasevich et al., 1975). It is generally accepted that electrostatic attraction, involving cation exchange, is the primary mechanism underlying the adsorption and intercalation of proteins by montmorillonite, at least at pH values below the isoelectric point (Baron et al., 1999; Boyd and Mortland, 1990; Ding and Henrichs, 2002; Garwood et al., 1983; Norde et al., 2008; Servagent-Noinville et al., 2000; van der Veen et al., 2004). An elegant demonstration of this process has been provided by Szabo´ et al. (2007) during the layer-by-layer construction of clay/protein films. Using Naþ–saponite and papain–phosphate, they noted that phosphate was completely missing from the films because the

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positively charged papain replaced sodium ions which, together with phosphate, were removed when the films were washed with water. The cation exchange reaction, however, may not be stoichiometric in that less than the equivalent amount of inorganic cations, originally present at the clay surface, is replaceable (Quiquampoix and Ratcliffe, 1992; Szabo´ et al., 2008) because the surface requirement of the adsorbing protein far exceeds the area per exchange site (0.8 nm2 for montmorillonite). Following Hendricks (1941), this phenomenon may be referred to as the ‘cover-up’ effect. Another noteworthy feature of the intercalation process is layer segregation (‘demixing’), giving rise to protein-rich and inorganic cation-rich layers that are randomly interstratified within a single montmorillonite particle. Since the basal spacing of the fully exchanged clay–protein complex is normally greater than that of the original (parent) clay, the basal spacing of partially exchanged (up to 55%) samples is intermediate between the two end-members. Thus, layer segregation is characterized by an increase in basal spacing (of the clay–protein complex) as intercalation progresses (cf. Figure 8.4). In addition, the basal or d(001) peaks are often diffuse, and there is a non-integral (irrational) series of basal reflections. To complicate matters further, the layer charge of montmorillonite is often inhomogeneous, varying from layer to layer within a single particle (Lagaly and Weiss, 1976). It is against this background that we shall attempt to rationalize the scattered data on clay–protein systems and hopefully present them as a more or less coherent body of information.

8.3. FORMATION AND PROPERTIES OF COMPLEXES Ensminger and Gieseking (1939, 1941) pioneered the systematic investigation into the clay mineral–protein interaction. To this end, they prepared complexes of montmorillonite with gelatin and albumin by acidifying an alkaline (pH 10) suspension of the sodium clay containing different amounts of protein. X-ray diffraction analysis (of the dry complexes) showed that intercalation had occurred, the extent of which increased as the suspension pH decreased (from 7 to 2.7). This finding indicated that complex formation primarily involved an exchange between the cationic (

NH3þ) groups on the amino acid side chains of the protein and the interlayer Naþ counterions. Further support for the cation exchange mechanism was provided by the observation that prior treatment of the protein with nitrous acid, formaldehyde or lignin, all of which would have deactivated the amino groups on the molecule, resulted in a marked decrease in the extent of intercalation. In addition, complex formation caused a reduction in the cation exchange capacity of the clay sample. They also noted that the basal spacing of the complexes increased (up to5 nm) with the amount of protein in the system, indicative of layer

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Formation and Properties of Clay-Polymer Complexes

segregation. Similar results were obtained with casein, haemoglobin, pancreatin, pepsin and protamine. In examining the interaction of casein and gelatin with montmorillonite, Chiang and Chao (1964) also found that the extent of adsorption correlated with the cation exchange capacity of the clay mineral sample. The ease with which these proteins could replace the inorganic counterions was influenced by cation valency, providing further evidence for a cation exchange process. Morgan and Corke (1976) have proposed similarly for the interaction of glucose oxidase with various clay minerals. A more detailed study of protein intercalation by montmorillonite was made by Talibudeen (1950, 1955) using X-ray diffraction analysis. The interlayer complexes were obtained by immersing air-dry oriented flakes of Naþ–montmorillonite in aqueous solutions (usually 2% w/v), and at a pH below the isoelectric point (pI), of the protein. In the case of gelatin (pI4.5) at pH 2.5, the basal spacings were indicative of intercalation of a discrete number of extended molecular layers. What was interpreted as a two-layer complex (basal spacing1.77 nm) appeared to be the most stable arrangement, while a one-layer complex (basal spacing1.4 nm) generally obtained when the amount of protein in solution was relatively low (<0.5% w/v) and the pH was greater than 3.5. Under special conditions, and then only rarely, was a spacing of 2.63 nm recorded (in addition to the 1.77-nm line) due possibly to the formation of a four-layer complex. More recently, Zheng et al. (2002) reported a basal spacing of 4.42 nm for an interlayer complex of Naþ–montmorillonite with gelatin, while Martucci et al. (2007) and Bae et al. (2009) obtained an exfoliated complex showing no diffraction peaks (cf. Chapter 7) by mixing gelatin with ultrasonically treated Naþ–montmorillonite. Subtracting the assumed thickness of 0.95 nm for an individual montmorillonite layer (cf. Figure 1.9) from the observed basal spacing gave an interlayer separation (‘D-value’) of 0.45 nm for the one-layer complex and 0.82 nm for the two-layer complex. After allowing for the normal amount of recession (‘keying’) into the ditrigonal basal oxygen network (cf. Figure 1.1), these values were comparable to those obtained for single- and double-layer complexes with amino acids and small peptides (Greenland et al., 1962, 1965; Theng, 1974; Weiss, 1969). Thus, the intercalated proteins appeared to adopt an extended, slightly buckled, b-type conformation with the amino acid side chains lying more or less flat on the interlayer surface. Assuming that the gelatin molecules consisted of single chain, monodisperse, random coils in solution (Finer et al., 1975), such an interlayer conformation would imply that the overall adsorption energy was sufficiently large to offset the loss of entropy accompanying the transition from coil to extended chain. The combination of cation exchange, entropy effects associated with the displacement of surface-adsorbed water, and van der Waals attractive forces could conceivably meet this requirement.

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The adopted b-keratin type structure, however, is apparently not restricted to fibrous proteins such as gelatin. Under similar conditions, edestin and pepsin also yielded stable complexes with a basal spacing of 1.8 nm, suggesting the presence of two layers of extended polypeptide chains in the interlayer space (Talibudeen, 1955). If so, intercalation into montmorillonite appeared to induce a conformational change affecting both the tertiary and secondary structure of the proteins as Simpson and Hughes (1978) have suggested for arylsulphatase. In the absence of analytical and kinetic data, however, we can only speculate that the molecules underwent a conformational change just prior to, or immediately after, coming into contact with the external particle surface of montmorillonite. Still possessing some mobility, the partially unfolded protein could then slowly penetrate the interlayer space where it attained its final extended form. Such a mechanism is in keeping with the kinetic data of McLaren et al. (1958) who noted that although the initial uptake of proteins by montmorillonite was very rapid, many hours were required for the system to attain equilibrium. Weiss (1969) subsequently found that the basal spacing of montmorillonite complexes with a number of proteins (gelatin, gliadin, human albumin, egg albumin, pepsin, serum albumin, salmin and zein) never exceeded 1.8 nm, although the amount adsorbed varied over a fairly wide range. This would also indicate intercalation of up to two layers of extended polypeptide chains as Talibudeen (1950) has proposed. For a given protein and at pH values when the molecule carried a net positive charge, adsorption was greater for Naþ– than for Ca2þ–montmorillonite. This might be because Naþ–montmorillonite shows extensive interlayer swelling, allowing a large surface to be accessible to the protein. In addition, the amount adsorbed tended to rise as the solution pH was increased from 2.7 to near the isoelectric point of the proteins. Other examples of this behaviour will be described more fully below. Pinck and co-workers (Pinck, 1962; Pinck and Allison, 1951; Pinck et al., 1954) assessed the biostability of clay-adsorbed proteins by preparing interlayer complexes of montmorillonite with some proteins (e.g. gelatin and ovalbumin), incubating the materials in sand or soil and measuring the amount of carbon dioxide evolved over a period of some weeks. By concurrently examining the complexes using X-ray diffractometry, the extent of decomposition could be related to the changes in basal spacing that took place during incubation. Following Talibudeen (1950), the complex with a basal spacing of 1.5 nm, containing 10 wt% protein, was assumed to have a single layer of intercalated protein molecules (Pinck et al., 1954). Since the basal spacing remained sensibly constant over the incubation period, the interlayer material was apparently protected from enzymatic degradation (proteolysis). On this basis, 18% or so of protein carbon that evolved as CO2 would represent material adsorbed to the external particle surface of montmorillonite and, as such, readily accessible to extracellular enzymes. On the other hand, close to 80%

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Formation and Properties of Clay-Polymer Complexes

of the interlayer material in complexes with two or more layers, containing 35–54 wt% protein, was degradable. At the same time, the basal spacing fell from an initial value of 3 nm to 1.2 nm at the end of the incubation period. It would therefore appear that a basal spacing of 1.8 nm or greater is required for extracellular enzymes—in reality, their active sites—to gain access to interlayer substrates. Earlier, Ensminger and Gieseking (1942) noted that albumin and haemoglobin adsorbed to montmorillonite were largely resistant to hydrolysis by pepsin (at acidic pH) and pancreatin (at alkaline pH). By contrast, extensive protein decomposition occurred for the corresponding complexes with kaolinite. Similar results were reported by Birch and Friend (1956) for gelatin. Likewise, Berg et al. (2001) reported that gelatin adsorbed on (external) silica/glass surfaces was partially degradable by a proteolytic enzyme (krilase). The extensive studies by Stotzky and co-workers, as summarized by Stotzky (2004), on the fate in soil of insecticidal proteins from B. thuringiensis also show that attachment to external clay surfaces provides partial protection against microbial decomposition, enabling the (bound) proteins to retain their biological activity. Although Ensminger and Gieseking (1942) did not measure basal spacings, it seemed likely that the protective effect of montmorillonite was related to its capacity for intercalating various proteins, something that kaolinite, silica and glass did not possess. What should add that interlayer sorption per se does not invariably or necessarily impart ‘immunity’ to the intercalated molecules against enzymatic attack (Estermann and McLaren, 1959; Estermann et al., 1959). The decomposition rate of intercalated molecules, however, would be much lower than that of externally adsorbed species. Further, intercalation may induce changes in protein conformation which would interfere with enzyme–substrate ‘recognition’ (Rabbi et al., 2010). As already mentioned, steric factors related to the extent of interlayer expansion influence the biostability of intercalated proteins. This point will be taken up below when we discuss the montmorillonite–lysozyme interaction. The interactions of clay minerals with proteins and enzymes have been extensively and systematically studied by McLaren and his colleagues and reported in a series of papers (Benesi and McLaren, 1975; McLaren, 1954a,b, 1975; McLaren and Estermann, 1956, 1957; McLaren and Packer, 1970; McLaren and Peterson, 1965; McLaren et al., 1958; Skujin¸sˇ et al., 1959). A good proportion of their work deals with the formation and properties of complexes with kaolinite. Besides being incapable of intercalating proteins and enzymes, kaolinite has a low exchange capacity and a limited propensity for adsorbing proteins. This is not to say, however, that the interaction process is any less complicated. In order to preserve the thread of our discussion, the focus of attention will first be directed to the montmorillonite–protein interaction.

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In an early paper McLaren et al. (1958) characterized the reaction of Naþ– montmorillonite with some proteins (chymotrypsinogen, gelatin, haemoglobin, b-lactoglobulin, lysozyme, pepsin) by determining the adsorption isotherms at different pH and relating the basal spacing of the resultant complexes to the amount adsorbed. In common with lysozyme (Figure 8.1), the isotherms were of the H-type showing a steep initial rise and levelling off to a long plateau at fairly low solute concentrations. Such isotherms are typical of polymer adsorption to clay and mineral surfaces, indicating a high affinity of the solute for the surface (Giles, 1970; Giles et al., 1960). Another feature of the clay–protein and the solid–protein interaction, in general, is that the curve relating plateau adsorption (Gp) to pH commonly shows a maximum close to the isoelectric point (pI) of the solute (Brash and Lyman, 1971; Claus and Filip, 1988; Haynes and Norde, 1994; Kragh, 1964; Norde, 2008; Norde et al., 1986, 2008; Quiquampoix, 2008a,b). Figures 8.2 and 8.3 illustrate this behaviour for a number of proteins adsorbing to montmorillonite. The decrease in adsorption at pH>pI, represented by the ‘right branch’ of the Gp versus pH curve, may be ascribed to electrostatic repulsion between the protein anion and the negatively charged montmorillonite surface. This effect is further enhanced by competition for surface sites between the cations in solution (e.g. Naþ) and the protein. That adsorption also 2.0

10.3 9.2 1.5

6.7 6.1

Amount adsorbed (g/g)

FIGURE 8.1 Isotherms at 298K for the adsorption of lysozyme by montmorillonite at the indicated pH values. The curves were determined at an ionic strength (I) of 0.0375 except for the pH 1.9 curve (I¼0.053). The data for pH 1.9, 4.1, 6.1 and 6.7 were obtained in sodium citrate buffer and those at pH 7.0, 9.2 and 10.3 in sodium borate buffer. From McLaren et al. (1958).

7.0 4.1 1.0

1.9 0.5

0

0

0.5 1.0 1.5 Final concentration (mg/mL)

2.0

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Formation and Properties of Clay-Polymer Complexes

2.0

La

Amount adsorbed (g/g)

1.5

1.0 Ch Pe 0.5

0 1.0

3.0

5.0

7.0 pH

9.0

11.0

13.0 2.0

He Ge

Amount adsorbed (g/g)

1.5

Ly 1.0

0.5

1.0

3.0

5.0

7.0 pH

9.0

11.0

0 13.0

FIGURE 8.2 Relationship between the maximum extent of adsorption and suspension pH for various proteins adsorbing to montmorillonite. The ionic strength was 0.0375. Vertical dotted lines indicate the isoelectric point (pI) of the respective proteins: Ch, chymotrypsinogen; Ge, gelatin; He, haemoglobin; La, b-lactoglobulin; Ly, lysozyme; Pe, pepsin. From McLaren et al. (1958).

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1.5

100

1.2

80

0.9

60

0.6

40 Methods NMR spectroscopy: UV absorption:

0.3

0.0

Mn2+ displaced (%)

Maximum amount of BSA adsorbed (g/g)

Chapter

20 0

2

3

4

5

6

7

pH FIGURE 8.3 Diagram showing the effect of medium pH on the maximum adsorption of bovine serum albumin by montmorillonite and on the percentage of Mn2þ ions displaced from the clay surface during protein adsorption, determined by NMR and UV absorption spectroscopy. From Quiquampoix and Ratcliffe (1992) as modified by Quiquampoix (2008b).

decreases at pH
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Formation and Properties of Clay-Polymer Complexes

protein adsorbed as a function of solution pH. At pH>pI, the amounts of counterion released and protein adsorbed decreased in the same proportion, indicating a reduction in surface coverage due to electrostatic repulsion with no major change in conformation (Quiquampoix et al., 1989). At pH
Amount adsorbed (g/g)

1.5

1.0

0.5

0

1

2

3 4 Basal spacing (nm)

5

FIGURE 8.4 Variation in basal spacings of montmorillonite–lysozyme complexes with the amount of protein adsorbed. The pH varied from 1.9 to 10.3 and the ionic strength from 0.0375 to 0.1375. Most of the points were obtained at pH 6.6 and an ionic strength of 0.0375. From McLaren et al. (1958).

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4.4 nm (for complexes containing 1.0–1.6 g/g). The 2.7-nm spacing probably arose from random interstratification within a particle of fully expanded layers (d(001)¼4.4 nm) and fully collapsed layers (d(001)¼1.0 nm) in a 1:1 proportion, that is, (4.4þ1.0)/2¼2.7. If so, the 4.4-nm basal spacing would indicate intercalation of a single layer of (globular) lysozyme molecules. This interpretation was called into question by Laura (1975) who suggested that a basal spacing of 4.4 nm was due to intercalation of multimolecular layers of protein. This suggestion was made in the context of the hypothesis that the decomposition of organic matter in soil could be explained in terms of the protolytic action of water (Laura, 1974). Being mobile, multilayers of clay-adsorbed organic molecules were susceptible to protolysis (by water), whereas the thermodynamically stable monolayer arrangement was not. Thus, the basal spacing of montmorillonite–protein complexes, in terms of the shape and thickness of the intercalated species, became the subject of a polemic between Laura (1975, 1976), on the one hand, and McLaren and Barshad (1976), on the other. Although it is not our intention to enter into the debate, we wish to make the following comments. First, it is entirely feasible that some clay-associated organic compounds in soil can decompose through an abiotic pathway, since clay minerals are capable of activating and catalyzing a variety of organic reactions (Adams and McCabe, 2006; Ruggiero et al., 1996; Theng, 1974, 1982). Nevertheless, it is incontestible that the decomposition of organic compounds in soil, at least under unsterilized conditions, is a microbially mediated process, catalyzed by extracellular enzymes (Naidja et al., 2000; Ruggiero et al., 1996; Zimmerman and Ahn, 2010). In response to Laura’s (1974) hypothesis, Stotzky and Kunc (1975) have pointed out that the microbial (enzymatic) concept of degradation is supported by a large, solid body of experimental evidence. Our second comment relates to the conformation of intercalated proteins in montmorillonite. As remarked on earlier, ‘hard’ proteins (e.g. lysozyme) tend to retain their native globular conformation on adsorption. Thus, a basal spacing of 4 nm is consistent with the presence of a single layer of molecules in the interlayer space, given that the lysozyme molecule is slightly ellipsoidal with a dimension of 4.533 nm (Imoto et al., 1972; Malmsten, 1995). In accordance with Pinck’s (1962) observation, the interlayer material would be accessible to extracellular enzymes, and if so, its degradability ceases to be problematic. Nevertheless, the X-ray diffraction data of McLaren et al. (1958) are amenable to a different interpretation, especially in ascribing the 4.4-nm spacing to formation of a monolayer complex, containing 1.25 g lysozyme/g clay (Figure 8.4). We suggest that the increase in interlayer expansion (as adsorption progressed) was more gradual than step-wise, although the presence of two expansion stages was discernible. The first stage was apparently completed at a basal spacing of 3.3 nm, while the second stage would appear

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to terminate near or slightly below 6 nm. Further, the diffraction peaks in the second expansion stage were rather broad, indicating an irrational series of basal reflections. Thus, formation of a one-layer complex was presumably completed at a basal spacing of 3.3 nm, giving a monolayer capacity of 0.85 g/g. As uptake continued beyond this level, a second layer of protein was intercalated, attaining completion at 1.7 g/g. Basal spacings recorded in the range of 1– 3.3 nm and 3.3–6 nm probably arose from segregation of zero and one and of one and two molecular layers within a single particle, respectively. If so, the 4.4-nm spacing represented a complex in which single and double layers of molecules co-existed, rather than a monolayer complex. The observed spread of basal spacings for a given amount adsorbed reflected the effect of pH and ionic strength on molecular conformation, since neither factor was kept constant. Subsequent measurements by Armstrong and Chesters (1964) indicate that montmorillonite is indeed capable of intercalating more than one layer of lysozyme molecules depending on solute concentration and pH. Using Mg2þ–montmorillonite as the parent material, they were able to prepare interlayer complexes with basal spacings of 3.5, 5.9 and 7.4 nm. As McLaren et al. (1958) reported, the initial reaction was rapid with over 90% of the added lysozyme being adsorbed within the first hour and uptake continuing up to 12 h after mixing the protein with the clay suspension. The isotherms were again of the H-type, and the extent of adsorption increased to a maximum near the isoelectric point of lysozyme (pI11). The basal spacings quoted above were for complexes formed at pH 10.5 at three (increasing) protein/clay ratios, corresponding to plateau adsorption (Gp) values of 0.93, 1.91 and 2.45 g/g, respectively. In this instance, however, the basal peaks were sharp, and at least one higher (rational) order of basal reflection was recorded. The apparent absence of layer segregation and the persistence of the basal reflection at 3.5 nm in the X-ray diffractograms strongly suggest that the first (lowest) level of expansion is due to the formation of a single-layer complex. Szabo´ et al. (2007) measured a similar basal spacing for an interlayer complex with Naþ–saponite. This suggestion accords with our interpretation of the data shown in Figure 8.4 (McLaren et al., 1958). Taking 350 m2/g as the available interlayer surface of montmorillonite, and assuming no overlap between close-packed molecules, the monolayer capacity of 0.93 g/g is equivalent to 2.7 mg/m2. The observed basal spacing of 3.5 nm for the monolayer complex corresponds to an interlayer separation (D-value) of 2.55 nm. This value is slightly smaller than the short axis (3 nm) of the ellipsoidal lysozyme molecule (Imoto et al., 1972; Malmsten, 1995), or the diameter (3.3 nm) of an equivalent sphere (McLaren and Peterson, 1961; McLaren et al., 1958). These observations would suggest that lysozyme intercalates into montmorillonite as a flattened ellipsoid in a ‘side-on’ orientation with its short axis perpendicular to the interlayer surface. Using ellipsometry, Malmsten (1995)

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similarly obtained a layer thickness of 30.5 nm for a monolayer complex (3.5 mg/m2) of silica with lysozyme, indicative of a ‘side-on’ type adsorption. The suggested ‘flattening’ of intercalated lysozyme molecules is also consistent with the loss of a-helix content on adsorption to silica nanoparticles (Billsten et al., 1995; Vertegel et al., 2004). On this basis, the complex with a basal spacing of 5.9 nm (D-value¼4.95 nm) and containing 1.91 g/g may be identified with the presence of a double layer of intercalated lysozyme molecules adopting essentially the same (‘side-on’) orientation in the interlayer space. Formation of double-layer complexes between lysozyme and silica has also been reported by Wahlgren et al. (1995). Although lysozyme can form multilayers on muscovite and silica (Kim et al., 2002; Vertegel et al., 2004), the complex with a basal spacing of 7.4 nm (Armstrong and Chesters, 1964) cannot be explained in terms of the intercalation into montmorillonite of three molecular layers of lysozyme. This is because the interlayer separation (D-value¼6.45 nm) and amount adsorbed (2.45 g/g) for such an arrangement are much smaller than three times the values for the monolayer complex. Armstrong and Chesters (1964) suggested intercalation of a double layer of lysozyme molecules oriented ‘end-on’ with their long axis perpendicular to the silicate surface. The X-ray diffraction and adsorption data, combined with the molecular dimension of lysozyme, however, are more consistent with the presence in the interlayer space of a monolayer of flattened ellipsoidal molecules adsorbed in an ‘end-on’ orientation than with intercalation of two protein layers. Malmsten (1995) has proposed similarly for the orientation of immunoglobulin adsorbed to silica gel. Thus, globular proteins with an ellipsoidal shape tend to adsorb in a ‘sideon’ orientation at low solute concentrations and adopt an ‘end-on’ configuration when the protein concentration in the bulk solution is high (Nakanishi et al., 2001). A change in orientation from a ‘side-on’ to an ‘end-on’ arrangement, giving rise to a decrease in surface requirement of the adsorbing solute, has also been reported for many interlayer complexes of montmorillonite with small, well-defined organic species (Theng, 1974). Further, adsorption to clay mineral surfaces of ‘hard’ globular proteins does not commonly induce drastic changes in their secondary structure. Accordingly, unless the adsorption data would indicate otherwise, basal spacings in excess of 3 nm might be expected to obtain for many montmorillonite–protein complexes, containing a single layer or, at most, a double layer of intercalated molecules. The question arises why some proteins, such as gelatin and pepsin, can apparently unfold when they intercalate into montmorillonite. The reason for this may be sought in the solution properties of the proteins concerned as much as in the experimental approach used in forming the clay–protein complex. Being fibrous, gelatin can intercalate as multilayers of extended molecules depending on the pH and solute concentration as described earlier. Similarly, pepsin tends to unfold, and behave more or less like a linear polyelectrolyte, when the solution pH falls below the isoelectric point due to

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Formation and Properties of Clay-Polymer Complexes

electrostatic repulsion between cationic groups on the molecule (McLaren and Lewis, 1950). The ability of BSA to unfold at the solid/liquid interface is also influenced by the solution pH and protein concentration (Bull, 1956; Chattoraj and Bull, 1959; MacRitchie, 1972). In the final analysis, the mode of conformation that a protein would adopt at the interface is determined by the balance between internal forces tending to maintain the structural integrity of the molecule and the affinity the molecule has for the adsorbing surface. Tarasevich et al. (1975) made an early attempt at applying IR spectroscopy to assess the conformation of adsorbed proteins in montmorillonite. The complex that they examined in some detail was one between Naþ– montmorillonite and serum albumin (molecular weight65,000 Da), formed at pH 3.5 and containing 250 mg protein/g clay. The IR spectrum of the airdry complex (basal spacing¼2.3 nm) showed NH stretching bands at frequencies close to those given by polypeptides and albumin in the solid state. This observation was interpreted in terms of the formation of N

H  O¼¼C hydrogen bonding between adjacent, more or less extended, polypeptide chains in the adsorbed protein. Heating the complex to 423–573 K disrupted these interpeptide links, inducing the formation of N

H  O

Si bonding between the interlayer protein (or its decomposition products) and the silicate surface. The occurrence of certain absorption bands in the frequency range of 1800–1200 cm
1, particularly the presence of an amide II component with a maximum at 1524 cm
1, indicated increased development (in the interlayer species) of a b-type conformation at the expense of the a-helix structure relative to the solid protein. Similarly, Fusi et al. (1989) observed a decrease in a-helix content of albumin on adsorption to Naþ–montmorillonite. The conformational changes of ‘soft’ BSA, adsorbed from buffered D2O solutions to montmorillonite and talc, have been investigated in some detail by Servagent-Noinville et al. (2000), using Fourier-transform IR spectroscopy. Adsorption to (hydrophilic) montmorillonite over the pH range of 4.3–5.6 led to extensive unfolding of both external and internal a-helix structures, involving about 20% of the backbone. Some secondary structure modification was observed even at the isoelectric point (pH 4.8), while at pH 2.9, only the external a-helix structures appeared to unfold. Less pronounced conformational changes were observed with uncharged (hydrophobic) talc where only external helical regions appeared to unfold at pH 4.8. By contrast, a-chymotrypsin (a ‘hard’ protein) underwent only minimal changes in its secondary structure on adsorption to montmorillonite (Baron et al., 1999). More recent measurements by Rytwo et al. (2010) also indicate that adsorption to montmorillonite induced a change in the secondary structure (of a viral protein), while no conformational modification occurred on adsorption to an organically modified (hydrophobic) clay sample. Interestingly, even a ‘hard’ protein like lysozyme can undergo appreciable structural changes when it adsorbs to hydrophilic silica nanoparticles (Billsten et al., 1995). Further, the loss of a-helix content is strongly influenced by particle size and

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solution pH (Vertegel et al., 2004). The effect of clay particle size on molecular conformation of adsorbed proteins merits close investigation. Leaving aside the effect of adsorption on protein conformation and stability, we may ask what determines layer ordering in montmorillonite–protein complexes. The literature would indicate that layer segregation, giving rise to randomly interstratified structures within a montmorillonite particle, occurs when the protein solution is mixed with a suspension of the clay mineral in the sodium form (Ensminger and Gieseking, 1939, 1941; McLaren et al., 1958; Pinck et al., 1954). On the other hand, little, if any, segregation occurs when air-dry oriented flakes of Naþ–montmorillonite are used (Talibudeen, 1950, 1955) or when the protein is added to a suspension of divalent cationexchanged montmorillonite (Armstrong and Chesters, 1964). The answer to our question must lie in the stacking mode of the silicate layers making up a particle. In an aqueous low-salt or salt-free suspension of Naþ–montmorillonite, the layers tend to be displaced relative to each other and separated by distances that are large (>2 nm) but by no means uniform (Norrish, 1954). This would allow various proteins to be intercalated, but the mutual alignment of silicate layers in the interlayer complexes that form would remain somewhat disordered. At a solute concentration in excess of what is required for monolayer coverage, some interlayers may contain a single layer, and others a double layer, of protein molecules. The variation in basal spacing as a function of the amount adsorbed would, therefore, show the behaviour depicted in Figure 8.4. On the other hand, for air-dry oriented Naþ–montmorillonite flakes and polyvalent ion clay systems, the stacking of the silicate layers in a particle is relatively ordered. Although this arrangement would not preclude layer segregation from occurring during protein intercalation, more ordered single- and double-layer complexes would be expected to form by comparison. Since the parent clays show limited interlayer swelling, little intercalation would occur unless the protein in question can uncoil as it apparently did for Talibudeen (1950, 1955) and Weiss (1969). That lysozyme could penetrate the interlayer space of Mg2þ–montmorillonite (Armstrong and Chesters, 1964) would suggest that some interlayer swelling preceded protein intercalation. This preexpansion was probably induced by the exchange of interlayer Mg2þ ions for Naþ ions supplied by the (borate) buffer solution. The X-ray diffraction data of Harter and Stotzky (1973), using unbuffered systems, were consistent with our suggestion. Vansant and Uytterhoeven (1972) have argued that layer segregation may not occur until the complexes are dried prior to X-ray diffraction analysis. There is good evidence to suggest, however, that segregation would obtain in both wet and dry montmorillonite systems, unless the wet clay is more fully expanded in suspension, in which case cation size is less important than the extent of interlayer expansion (McBride and Mortland, 1973). This proviso would apply to the aqueous Naþ–montmorillonite system at low protein

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Formation and Properties of Clay-Polymer Complexes

concentration and pH. Homogeneity might thus be expected to obtain until drying restricts the uniform distribution of protein cations in every layer throughout the montmorillonite particle. Thus, caution should be exercised in interpreting basal spacing data of clay–organic complexes in terms of the thickness and disposition of the intercalated molecule, especially for systems involving macromolecular compounds. Nevertheless, there is little doubt that some globular proteins (and enzymes) can intercalate into Naþ-exchanged 2:1 type layer silicates that can show extensive interlayer swelling (in aqueous suspension). On the other hand, adsorption is confined to external particle surfaces in situations where interlayer expansion of the parent clay is either absent (as in kaolinite and illite) or limited (as in caesium- and polyvalent cation-exchanged montmorillonites) (De Cristofaro and Violante, 2001; Fiorito et al., 2008; Szabo´ et al., 2008; Vettori et al., 1999). There is no unequivocal evidence, however, for the occurrence in soil of interlayer montmorillonite–protein complexes (Perez Rodriguez et al., 1977). This might be because the concentration of ‘free’ proteins in the soil solution is very low. Even if such complexes were to form in montmorillonite-rich soils, the interlayer protein would generally be accessible to, and degradable by, soil enzymes (Estermann and McLaren, 1959; Estermann et al., 1959). Only if the basal spacing of the complex is limited to 1.4 nm, or has fallen to this value after partial decomposition of the interlayer protein, might intercalation offer protection against enzymatic attack (Pinck, 1962). We have already seen how the exchangeable inorganic cations at the montmorillonite surface may influence protein uptake. Besides being directly involved in exchange reactions with the protein, the nature of these cations influences the extent of interlayer swelling of the clay and, hence, the ease with which the protein may be intercalated. These and related aspects have been more closely examined by Harter and Stotzky (1971, 1973). Their approach was to react montmorillonite samples containing different exchangeable cations (Hþ, Naþ, Ca2þ, Al3þ, La3þ, Th4þ) with a number of proteins, such as a-casein, catalase, a-chymotrypsin, edestin, b-lactoglobulin, lysozyme, ovalbumin, ovomucoid and pepsin, covering a wide spectrum of molecular weights (14,000–310,000 Da) and isoelectric points (2–11). Adsorption and retention (‘binding’) isotherms were determined, the latter being obtained by measuring the amount of protein remaining in the respective complexes after repeated washing with distilled water. The use of buffers was avoided so that the initial and final pH values of the system were somewhat dependent on the clay mineral and protein species used. The pH of an aqueous suspension of the various clay–protein complexes, however, was generally close to neutrality. Positive adsorption was recorded for all clay–protein combinations, even when the net charge on the solute molecule was negative. The binding curves indicated that uptake depended on the valency of the exchangeable cations,

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decreasing in the order Hþ>Naþ>Ca2þ>Al3þ>La3þ>Th4þ. This behaviour reflects the decreasing ability of a given protein to replace the inorganic cations from their exchange sites as well as the relative accessibility of the interlayer surface to the solute. The position of Naþ relative to Ca2þ in the series accords with the results of Weiss (1969), Tarasevich et al. (1975), Morgan and Corke (1976), Hamzehi and Pflug (1981), Lozzi et al. (2001), Kelleher et al. (2003) and Sinegani et al. (2005). X-ray diffraction analysis indicated that, with the exception of catalase, the proteins were able to penetrate the interlayer space of Hþ– and Naþ–montmorillonites (at clay/protein ratios>1/5). Failure of catalase to intercalate was related to its relatively high molecular weight (238,000 Da) and large size (9 nm diameter). Interestingly, the lysozyme complex with the monovalent ion-exchanged clay samples gave a basal spacing of 3.5 nm, close to the value reported by Armstrong and Chesters (1964) for their single-layer complex with Mg2þ–montmorillonite. The lysozyme complex with Ca2þ–montmorillonite, on the other hand, did not expand beyond a basal spacing of 1.9 nm. This observation provided further support to our suggestion that the Mg2þ ions in Armstrong and Chesters’ (1964) montmorillonite sample were replaced by Naþ ions of the buffer solution, enabling the interlayers to expand for intercalation to occur. Thus, in the absence of buffers, all the proteins used (except possibly for casein with La3þ–montmorillonite) failed to intercalate into polyvalent ion-exchanged montmorillonites because these minerals showed only limited interlayer expansion in water. For the same reason, AmpliTaqÒ DNA polymerase (Vettori et al., 1999) and cellulase (Sinegani et al., 2005) were not intercalated by Ca2þ–montmorillonite. The effect of molecular weight and size was also reflected in the extent of adsorption (Harter and Stotzky, 1973). If the amount adsorbed was expressed on a w/w basis, adsorption rose with molecular weight although the magnitude of the increase was not proportional to molecular weight. If expressed on a mol/g basis, however, uptake tended to increase with decreasing molecular weight. This might be because the smaller and more compact the molecule, the easier it is to establish contact with, and close packing at, the surface. Albert and Harter (1973) described the interactions of lysozyme and ovalbumin with different layer silicates, such as biotite–vermiculite, illite, kaolinite as well as montmorillonite. Again, no buffers were used, the suspension pH being varied by titrating the acid-washed clays with NaOH. Adsorption increased as the ambient pH rose to the isoelectric point of the proteins. Following Armstrong and Chesters (1964), they suggested that the net positive charge on the protein decreased with increasing pH, resulting in more solute being required to satisfy the negative surface charge. As pointed out earlier, the decrease in protein solubility, and the adoption of a more compact molecular conformation as the pH approaches the isoelectric point, would also contribute to the observed increase in adsorption. With the possible exception of kaolinite, an ion exchange mechanism was involved since the amount of

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Formation and Properties of Clay-Polymer Complexes

sodium ions released into solution (at pH7) was proportional to the quantity of protein retained by the clay minerals. Less sodium, however, was released from biotite–vermiculite and illite (as a proportion of their respective cation exchange capacity) than from montmorillonite. It would therefore appear that the ‘cover-up’ effect was not important for montmorillonite because of its relatively low surface charge density. Subsequently, Harter (1975) extended the investigation to montmorillonite saturated with different cations, namely, the first five of the alkali and alkaline-earth series as well as Hþ, Al3þ and La3þ. For cations of equal valency, adsorption decreased with an increase in ionic radius. This observation may be ascribed to surface accessibility. Thus, the sodium-exchanged clay adsorbed nearly five times as much lysozyme as its caesium counterpart. Further, adsorption was progressively reduced by adding increasing amounts of a neutral electrolyte to the system. Thus, in the presence of 2 M NaCl, uptake by Naþ–montmorillonite was only half as much as in NaCl solutions of 0.1 M. In the former situation, the diffuse double layers on interlayer surfaces were presumably so compressed as to inhibit protein intercalation. In addition, protein solubility would increase as the NaCl concentration is raised. This ‘salting-in’ effect of 1:1 electrolytes would extend to high ionic strengths (Mahler and Cordes, 1968) and contribute to the observed decrease in adsorption. Desorption studies indicated that only a small proportion (<10%) of adsorbed lysozyme and ovalbumin could be eluted from their complexes with water or moderately concentrated (up to 2 M) solutions of sodium chloride, sodium acetate and ammonium acetate (pH 4.8 and 7). This finding further supports the view that interactions other than cation exchange are involved in the adsorption process. So far, our attention has largely been focused on the interactions of proteins with 2:1 type layer silicates, with special reference to montmorillonite. We now wish to summarize the available information on the behaviour of proteins at the surface of other clay mineral species. As might be expected, interactions of kaolinite with proteins have many features in common with those of 2:1 type layer silicates (smectites). Thus, more than 75% of the maximum amount is usually adsorbed within minutes of contacting the clay mineral with the protein solution, after which there is a slow increase in uptake until equilibrium is attained (Lee et al., 2003; McLaren, 1954b). Also, the isotherms are often of the H-type with adsorption levelling off to a plateau whose value depends on the ambient solution pH. Likewise, the extent of adsorption tends to reach a maximum near the isoelectric point (pI) of the protein (Alkan et al., 2006; Barral et al., 2008; Claus and Filip, 1988; McLaren, 1954a; Morgan and Corke, 1976; Vettori et al., 1999). The maximum in the pH–uptake curve, however, is often broader and less well-defined than for montmorillonite as shown in Figure 8.5 for the adsorption of lysozyme (pI11) from an equilibrium concentration of 0.5 mg/mL (McLaren, 1954a; McLaren et al., 1958). This behaviour may largely be

8

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Proteins and Enzymes

FIGURE 8.5 Relationship between maximum adsorption and suspension pH for lysozyme adsorbing to kaolinite from universal buffer. The vertical dashed line indicates the isoelectric point of lysozyme, while the shaded points refer to two different kaolinite samples. From McLaren (1954a).

24

20 Amount adsorbed (mg/g)

Chapter

16

12

8

4

0

2

4

6

8

10

12

pH

explained in terms of electrostatic interactions between solute and surface and the pH-dependent charge characteristics of the reactants. As Bolland et al. (1976) have indicated, the positive surface charge of kaolinite tends to decrease as suspension pH increases from 3 to 7, a tendency that may continue up to pH 9–10. Since the (small) permanent negative charge due to isomorphous substitution remains constant over this pH range, the net negative charge would effectively increase. Thus, (plateau) adsorption would gradually rise as suspension pH increased from 3 to 7. At pH 7–8, the amount adsorbed together with the estimated net positive charge on lysozyme agreed well with the cation exchange capacity of the kaolinite used. This was evidence for a cation exchange mechanism, although a strong ‘cover-up’ effect was operative. Above pH 7, the kaolinite (edge) surface progressively acquired negative charges due to dissociation of exposed aluminol and silanol groups (cf. Figure 1.6), whereas the positive charge on lysozyme (pI11) continued to diminish. A decline in adsorption might therefore be expected as the suspension pH was raised from 7 to 11.5 and this accorded with the experimental data. Only at pH 12 and beyond, when both mineral and protein were negatively charged, did adsorption fall off sharply. The extent of protein/enzyme adsorption by kaolinite (expressed on a w/w basis), however, is generally much smaller than that by montmorillonite (Albert and Harter, 1973; Claus and Filip, 1988; Fiorito et al., 2008; Fusi et al., 1989; Gianfreda and Bollag, 1994; Haska˚, 1975; Helassa et al., 2009; Hughes and Simpson, 1978; Jaynes et al., 2005; Lai and Tabatabai, 1992; Lee et al., 2003; Morgan and Corke, 1976; Quiquampoix, 1987; Shindo et al., 2002; Venkateswerlu and Stotzky, 1992; Vettori et al., 1999; Zhou

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Formation and Properties of Clay-Polymer Complexes

et al., 2008). This is because adsorption by kaolinite is restricted to external particle surfaces, and the external surface area of kaolinite is smaller than that of montmorillonite. McLaren (1954a), for example, noted that the maximum adsorption of lysozyme (26–35 mg/g) corresponded reasonably well to the magnitude of the (external) surface area of the kaolinite samples used, on the assumption that 1 mg of protein occupied 1 m2 of accessible surface. Helassa et al. (2009) have observed similarly for the adsorption of (monomeric) insecticidal protein from B. thuringiensis to the external surface of kaolinite and montmorillonite. As compared with kaolinite, less information is available on the reactions of proteins with halloysite. There has been a recent upsurge of interest, however, in using hollow tubular forms of halloysite as encapsulants of bioactive molecules, supports for protein and enzyme immobilization, and nanoscale fillers in polymer–clay nanocomposites (Du et al., 2010; Lvov et al., 2008; Sun et al., 2010; Xie et al., 2011; Zhai et al., 2010). Natural halloysite nanotubules (0.5–10 mm length; 50 nm external diameter; 15–20 nm internal pore diameter) have the advantage over synthetic carbon nanotubes in being abundant, inexpensive and biocompatible (Lvov and Price, 2008; Vergaro et al., 2010). Although halloysite can intercalate a single layer of water molecules (cf. Figure 1.7) and a range of small polar organic molecules (Churchman and Theng, 1984; Joussein et al., 2005), the formation of interlayer halloysite– protein complexes has yet to be demonstrated. On the other hand, the cylindrical pore space of tubular halloysite is accessible to a variety of proteins and enzymes (Lvov and Price, 2008). Zhai et al. (2010), for example, have successfully accommodated a-amylase and urease in the internal pore space (‘lumen’) of halloysite nanotubules. They suggested that the enzymes (at pH 6) were attached to the positively charged pore surface through electrostatic interactions. With an isoelectric point of 4.6, a-amylase would be negatively charged at pH 6, while urease (pI¼6) would have a zero net charge. It seems likely, therefore, that entropy effects associated with conformational changes (in the adsorbed enzymes) and van der Waals interactions are as important as, if not more so than, coulombic attraction. Enzymes entrapped within halloysite tubules are resistant to water washing and show a high thermal and storage stability (Lvov and Price, 2008; Zhai et al., 2010). Similar improvements in reusability and storage stability have been reported by Sanjay and Sugunan (2008) for a number of enzymes intercalated into acid-activated (K10) montmorillonite. Proteins (and enzymes) would be expected to interact with external surfaces of halloysite before entering the cylindrical pore space. Indeed, appreciable adsorption would occur on external particle surfaces because of their relative accessibility to extraneous solutes. The mode of interaction, however, would differ since halloysite has an external siloxane surface, while the internal (pore) surface consists of aluminol groups (Theng et al., 1982). These points need to be clarified.

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Mills and Creamer (1971) made an early attempt at investigating the interactions of halloysite with proteins. A rod-shaped halloysite with a cation exchange capacity of 10 cmol/kg was selected. Using casein and its as1-, b-, g- and k-components, they noted that the rate of uptake was initially rapid but this fell off after the first hour of contacting the protein with the clay suspension. For whole casein at various protein/clay ratios, adsorption declined rather sharply as the ambient pH increased from 6 to 8.5, and then levelled off with a further increase to pH 10 (Figure 8.6). The pH range of 6–10 was selected on the basis of the limited solubility of casein below pH 6 and its susceptibility to hydrolysis above pH 10. Since the isoelectric point (pI) of casein and its components is close to pH 5, the curves in Figure 8.6 represent the ‘right-hand’ branch of the pH–uptake relationship (Figure 8.2). The (expected) maximum adsorption at pHpI is therefore not observed in this instance. Between pH 6 and 10, the protein carries a net negative charge, the magnitude of which increases with rising pH. This effect together with increased competition by Naþ ions for exchange sites at the clay surface would partly account for the observed decrease in uptake. That casein was at all times positively adsorbed rather than repelled by the negatively charged halloysite surface would indicate that electrostatic interactions, as proposed by Mills and Creamer (1971), are not important.

50

Amount adsorbed (mg/g)

40

30

20

10

0 5.5

6.0

6.5

7.0

7.5

8.0 pH

8.5

9.0

9.5

10.0

FIGURE 8.6 Effect of suspension pH on the adsorption of whole casein by halloysite. The data were obtained using a 6% w/v clay suspension and at different protein/clay ratios (g protein/g clay): (○), 0.01 g/g; (●), 0.02 g/g; (D), 0.04 g/g; (▲), 0.06 g/g; (□), 0.10 g/g. Adsorption was measured after shaking the reactants for 20 h at 298 K. From Mills and Creamer (1971).

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Formation and Properties of Clay-Polymer Complexes

Further, the observed order of adsorption: g-casein>b-casein>a-casein is also the sequence of increasing net negative charge on the protein. We rather suspect that casein, being a ‘soft’ protein, could undergo structural changes (decreased a-helix content) on contact with the mineral surface. The resultant gain in conformational entropy would thus provide the driving force for adsorption, possibly augmented by van der Waals and hydrogen bonding interactions (Norde et al., 2008; Table. 8.1). Research into the interactions of proteins with palygorskite (‘attapulgite’) and sepiolite has mainly been directed at using these fibrous phyllosilicates (cf. Chapter 1) as supports for enzyme immobilization (Carrasco et al., 1995; de Fuentes et al., 2001; Hamzehi and Pflug, 1981; Huang et al., 2008; Sedaghat et al., 2009b). Since the dimension of the channels (pores) in both palygorskite and sepiolite is much smaller than the size of protein molecules, adsorption is restricted to external particle surfaces. Hamzehi and Pflug (1981), for example, observed that enzyme uptake decreased in the order montmorillonite>palygorskite>kaolinite, although the specific surface area of montmorillonite (measured by adsorption of nitrogen or small polar organic molecules) is smaller than that of palygorskite. Much of this area in palygorskite, however, is contained in the channel structure which is inaccessible to proteins and enzymes. Perderiset et al. (1988) obtained H-type isotherms for the adsorption of BSA by palygorskite. The amount adsorbed at pH 7.4 (70 mg/g) represented about 1/3 of the specific surface area, indicating that 2/3 of the area was inaccessible. The extent of adsorption at pH 4 (when the protein was positively charged) was about twice that at pH 7.4, indicating the importance of electrostatic interactions in the process. Being a ‘soft’ protein, BSA would also have undergone conformational changes on adsorption as we have suggested for casein. In view of the propensity of allophane-rich soils for retaining and stabilizing large amounts (up to 20%) of organic matter (Dahlgren et al., 2004; Hiradate et al., 2004; Parfitt, 2009; Parfitt et al., 1997; Wada, 1989), surprisingly little is known about the reactions of allophane with proteins and enzymes. This lack of information is partly related to the problem of characterizing allophane by X-ray diffraction and partly to the restricted geographical distribution of allophane-rich soils as compared with non-volcanic soils. Early work by Aomine and Kodama (1956) and Harada (1959) indicated that the rate of albumin decomposition in allophanic soils was much lower as compared with soils containing layer silicate minerals. Similarly, Zunino et al. (1982) found that the decomposition of some simple and polymeric organic compounds in volcanic ash-derived soils was substantially reduced in comparison with non-allophanic soils. Low rates of carbon turnover in allophane-rich soils have also been reported by Torn et al. (1997) and Rasmussen et al (2006). Early on, Jackman (1955) suggested that enzymes were partially inactivated when they adsorbed to allophane surfaces. The results of

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recent investigations (Allison, 2006; Rosas et al., 2008; Shahriari et al., 2010), however, indicate that enzyme activity is often stimulated in the presence of allophane (Table 8.4). Milestone (1971) made an early attempt at assessing the interactions of proteins (BSA, gelatin, ovalbumin) with allophane. The rate of uptake was low with a half-time of 90 min or longer, probably because the proteins underwent a slow conformational change (unfolding) on contact with the allophane surface. Alternatively, the low adsorption rate was due to the slow diffusion of solute molecules into the pores of allophane aggregates as Theng (1972) has suggested for the interaction of alkylammonium salts with allophane. In keeping with this view, the adsorbed protein resisted elution by a 10
2 M solution of NaOH or Na2CO3. The adsorption isotherms generally conform to the L-type (Figure 8.7). The indicated plateau values, however, were more apparent than real in that a second rise frequently occurred when the solute concentration was raised beyond the specified range. Again assuming that 1 mg protein covers 1 m2 of surface, the apparent extent of adsorption would correspond to a surface coverage of only 25–35%, a value which probably represents the surface contained in the readily accessible (interparticle) pores. For a comparable concentration of protein in solution, adsorption generally attained a maximum between pH 5 and 6. In this pH range, the net charge on the allophane was slightly positive or zero (Theng et al., 1982), whereas that on the proteins

100 pH 6

Amount adsorbed (mg/g)

80 pH 7 60

40

20

0

0

0.2

0.8 0.4 0.6 Final concentration (mg/ml)

1.0

FIGURE 8.7 Isotherms for the adsorption of bovine serum albumin by allophane at two pH values. From Milestone (1971).

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Formation and Properties of Clay-Polymer Complexes

(pI¼4.6–4.9) was slightly negative. Above pH 6, both solute and surface were negatively charged causing a marked decline in adsorption. Similarly, charge–charge repulsion would partly account for the reduction in uptake below pH 5, since then the net charge on both the protein and the mineral was positive. That positive adsorption was observed at pH 7 may be ascribed to an increase in conformational entropy resulting from structural changes in the adsorbed proteins, possibly supplemented by van der Waals and H-bonding interactions. The same explanation would apply to the interaction at pH 6–9 of slaughterhouse effluent proteins (pI4.5) with allophane-rich soils (Russell, 1982). To round off this section, we will briefly describe the interactions of prion proteins with clay and soil minerals, and the layer-by-layer formation of clay– protein complexes. Prion diseases or transmissible spongiform encephalopathies (TSEs) are a class of neurodegenerative disorders that affect sheep and goats (scrapie), cattle (‘mad cow’ disease) and humans (Creutzfeldt–Jakob disease and koru). The infectious agent is a misfolded isoform (PrPSc) of the normal cellular prion protein (PrPC) (Prusiner, 1998). Burial of TSE-infected animal carcasses can introduce prions into soils which can then serve as a reservoir of the infectious agent (Leita et al., 2006; Schramm et al., 2006). The persistence and mobility of PrPSc in soil are not well understood. Like the insecticidal proteins from B. thuringiensis (Saxena et al., 2002), however, clay-adsorbed prions may be transported to surface and ground water. As might be expected, the interactions of prions with clay and soil minerals have features in common with those of ‘ordinary’ proteins. Thus, Ma et al. (2007) found that adsorption of PrPSc (aggregates) to quartz was highest near the isoelectric point of the protein and increased with the ionic strength of the ambient solution. Johnson et al. (2006) reported much larger uptake of purified PrPSc by montmorillonite than by kaolinite. Similar results were obtained for a recombinant murine prion protein by Polano et al (2008) who also noted that more protein was adsorbed by Naþ– than by Ca2þ–montmorillonite. In the absence of basal spacing data, we can only surmise that some intercalation occurred with Naþ–montmorillonite but not with the calcium-exchanged clay sample. Using Fourier-transform IR spectroscopy, Revault et al. (2005) found evidence to show that adsorption of non-infectious recombinant ovine PrP (recPrP) to montmorillonite led to an increase in its b-sheet content. This conformational change, however, was significantly different from the a-helix to b-sheet conversion that occurs when recPrP transforms to its pathogenic counterpart. It seems unlikely, therefore, that adsorption to clay minerals in soil would convert PrPC into the infectious PrPSc form. Rigou et al. (2006) observed that recPrP was strongly adsorbed to montmorillonite through the positively charged N-terminal domain of the protein. As a result, little, if any, desorption occurred when the complex was incubated

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273

with standard buffers and detergents. Using soil columns, Cooke and Shaw (2007) similarly found that recPrP has limited mobility in loamy sand and clay loam, indicative of strong mineral–solute interaction. Likewise, Johnson et al. (2006) reported strong adsorption of purified PrPSc by montmorillonite, causing a cleavage at the N-terminal site of the protein. Nevertheless, the bound protein remained infectious. Polano et al. (2008) have suggested that prion proteins can attach to humic substances through electrostatic and H-bonding interactions, begging the question whether complex formation with soil organic matter would affect prion infectivity. The formation of multi-layered smectite–polycation complexes through layer-by-layer deposition has been described in Chapter 5 (cf. Figure 5.7). Using a similar technique, Lvov et al. (1995) were able to assemble multilayer films consisting of alternating montmorillonite and protein layers with positively charged polyethylenimine acting as a ‘glue’. Likewise, Liu et al. (2005) obtained ‘nanocluster’ films by depositing positively charged haemoglobin (Hb) and negatively charged poly(styrenesulfonate) (PSS) on montmorillonite. In a variant approach, Lin et al. (2007a,b) intercalated (positively charged) poly(oxyalkylene)-amine into Naþ–montmorillonite or a synthetic fluorinated mica, and then replaced the polycation with BSA. This allowed BSA to be intercalated in its uncompressed, globular form, giving rise to basal spacings of 6–7 nm. The formation of clay mineral–protein films through a layer-by-layer technique has been systematically investigated by Szabo´ et al. (2007), using ultraviolet and attenuated total reflectance Fourier-transform IR spectroscopy and atomic force microscopy. They were able to construct films comprising up to 15–15 alternating layers of saponite and protein (lysozyme, papain, protamine) on glass, quartz and ZnSe supports. X-ray diffraction analysis indicated the presence of protein monolayers between the silicate layers. As already mentioned, electrostatic interactions between the negatively charged clay surface and positively charged protein provide the driving force for film formation. Hu and co-workers (Li and Hu, 2003; Liu et al., 2005; Zhou et al., 2002a,b) prepared stable films of montmorillonite with myoglobin, haemoglobin and horseradish peroxidase on pyrolytic graphite electrodes. Ultraviolet–visible and reflectance absorption IR spectroscopy indicated that the proteins retained their secondary structure. The clay/protein film electrodes could catalyze the reduction of oxygen, hydrogen peroxide and nitrite. The synthesis and applications of clay–enzyme electrodes as biosensors have recently been reviewed by Mousty (2010).

8.4. COMPLEX FORMATION AND ENZYME ACTIVITY As mentioned at the beginning of this chapter, enzymes associated or complexed with clay minerals operate in a structurally restricted system. Accordingly, the behaviour of such enzymes towards their respective substrates is different from that of their ‘free’ counterparts in solution.

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Formation and Properties of Clay-Polymer Complexes

A generally observable effect, imposed by the negatively charged (polyanionic) environment of the solid matrix such as clay minerals, is the shift to higher pH values of the pH-activity profile for adsorbed enzymes. This displacement in pH at which the enzyme shows optimal activity has been well documented for a number of enzymes attached to clay minerals and synthetic polyanions (Baron et al., 1999; Katchalski et al., 1971; Ladd and Butler, 1975; McLaren and Packer, 1970; Pflug, 1982; Ruggiero et al., 1996; Skujin¸sˇ et al., 1974). The extent of displacement is dependent on the surface charge density of the solid matrix. It is also influenced by the ionic strength (I) of the suspending medium, becoming effectively zero for I1. Another effect of the clay–enzyme interaction is the apparent enhancement of the Michaelis–Menten constant (Km) for the reaction rate of adsorbed enzymes as compared with their counterparts in solution. The pH effect may be explained in terms of the influence that the negatively charged surface exerts on the distribution of protons and positively charged substrates in its vicinity. Since cationic species, in general, tend to distribute themselves in a diffuse ionic double layer (cf. Figure 1.19), their concentration at the solid/solution interface is greater than that in the surrounding (bulk) solution (van Olphen, 1977). In terms of pH, the distribution of protons at the interface may be written as pHs ¼ pHb þ 0:43 zec=kT

ð8:1Þ

where the subscripts s and b refer to the surface and bulk phases, respectively; z is the valency of the cationic species which, in this case, equals unity; e is the electronic charge; c is the local electric potential, k is the Boltzmann constant and T the absolute temperature. For practical purposes, the following approximation can be applied (McLaren and Packer, 1970)
DpH ¼ pHs
pHb ¼ z=60

ð8:2Þ

where z is the zeta potential which can be estimated from the electrophoretic mobility (m) of the suspended particle, using the Smoluchowski equation m ¼ zє=4p

ð8:3Þ

m is considered to be negative for particles which move towards the anode; є and  are the dielectric constant and viscosity of the suspending medium, respectively. For particles with an equivalent spherical diameter (e.s.d.) 610
4 m, suspended in distilled water at 298 K, combination of Equations (8.2) and (8.3) after inserting the appropriate numerical values yields the relationship
DpH ¼ pHs
pHb ¼ 0:217m:

ð8:4Þ

For small clay particles (e.s.d.<10
6 m), the equation proposed by Hartley and Roe (1940):

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Proteins and Enzymes


DpH ¼ pHs
pHb ¼ 0:325m

275

ð8:5Þ

is more appropriate. Electrophoretic mobility values for some representative 2:1 and 1:1 type layer silicates (in the sodium form) range from
3.15 to
4.34 mms
1/V cm
1 (Marshall, 1964). Application of Equation (8.5) gives DpH values between 1.02 and 1.41, indicating that the mineral ‘surface’ is at least 10 times more acid than the bulk solution (suspension). The term ‘surface’ refers to the plane of shear at a distance of 0.5 nm from the basal (oxygen or hydroxyl plane) of the silicate layer where the potential is zeta (cf. Figure 1.20). Using IR spectroscopy, Harter and Ahlrichs (1967) derived a value of DpH2 for montmorillonite, suspended in a solution at pHb7. As Mortland (1970) has pointed out, the magnitude of the pH shift is influenced by the water content and nature of the exchangeable cation. The ionic strength of the system also affects DpH since the zeta potential (Equation 8.2) decreases as the electrolyte concentration of the ambient solution increases. The presence of buffers would also increase proton transport, shifting the pH-activity profile to more alkaline pH (Tischer and Kasche, 1999). It is difficult, however, to compare enzyme activity at different values of ionic strength because electrolyte addition would affect protein adsorption. A useful approach is to estimate pHs at a given value of pHb and ionic strength (I) by means of Equation (8.5). Using Equation (8.5), McLaren (1960) derived a DpH value of about
1.35 for lysozyme and 1.05 for kaolinite in solution at pHb¼8.05. This may be compared with the value of DpH¼0.54 for lysozyme adsorbed to kaolinite at the same ionic strength (I¼0.05). Thus, the pH at the ‘surface’ of lysozyme in solution is about two units higher than that of its counterpart at the kaolinite surface. Since enzyme activity is controlled by the pH environment in the vicinity of the substrate surface, the pH optimum for an enzyme acting on lysozyme in solution would be expected to be two pH units less than that in a system where lysozyme (substrate) is bound to kaolinite. In other words, the apparent pH optimum of the kaolinite-adsorbed enzyme would increase by DpH. This expectation is borne out by McLaren and Estermann’s (1957) experiment on the digestion by chymotrypsin of heat-denatured lysozyme, adsorbed to kaolinite (Figure 8.8). Table 8.2 lists some examples illustrating this effect for various enzymes acting on their respective substrates in the presence of clay minerals, polymers and other solid carriers. As expected, DpH is usually positive for systems where clay minerals act as a carrier. On the other hand, for systems involving polycationic supports, such as the DEAE–cellulose–invertase–sucrose assemblage, DpH is negative, while for systems at high ionic strength or when both carrier and substrate are negatively charged (e.g. ribonuclease–cation exchange resin–RNA), this parameter is sensibly zero.

276

Formation and Properties of Clay-Polymer Complexes

100

Relative activity (%)

80

60

40

20

5

6 7 8 9 pH (solution, suspension)

10

FIGURE 8.8 Diagram showing the relative activity of chymotrypsin in digesting denatured lysozyme in solution (open circles) and when adsorbed to kaolinite (half-shaded circles for universal buffer; open squares for ethylammonium buffer) as a function of pH. Dashed curve refers to data by Northrop (1922) for casein. From McLaren and Estermann (1957).

TABLE 8.2 Observed DpH Values for Some Carrier-Bound Enzymes Acting on Their Respective Substrates. Enzyme

Carrier

Substrate

Acid phosphatase

Kaolinite

p-Nitrophenyl phosphate

0

Huang et al. (2005)

Alkaline phosphatase

Bentonite

p-Nitrophenyl phosphate

0

Ghiaci et al. (2009)

a-Amylase

Bentonite

Starch

0

Sedaghat et al. (2009a)

Amyloglucosidase

Activated charcoal

Dextrin

þ1.0

Rani et al. (2000)

Catalase

Kaolinite

Hydrogen peroxide

þ0.65

Aliev et al. (1976)

Catalase

Montmorillonite

Hydrogen peroxide

þ1.0

Aliev et al. (1976)

Cellulase

Montmorillonite

Cellulose

þ1.2

Pflug (1982)

Chitinase

Kaolinite

Chitin

þ1.1

Skujin¸sˇ et al. (1974)

DpH

Reference

Continued

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277

Proteins and Enzymes

Chymotrypsin

Kaolinite

Lysozyme

þ2.0

McLaren and Estermann (1957)

Chymotrypsin

Montmorillonite

Lysozyme

þ2.0

McLaren and Peterson (1965)

Chymotrypsin

Carboxymethyl cellulose

Acetyltyrosine

Diastase

Acid-treated bentonite

Starch

þ1.7

Bajpai and Sachdeva (2002)

Ficin

Carboxymethyl cellulose

Benzoylarginine ethyl ester

þ0.3

Hornby et al. (1966)

Invertase

DEAE-cellulose

Sucrose


2.0

Suzuki et al. (1966)

Invertase

Acid-activated montmorillonite

Sucrose

þ1.0

Sanjay and Sugunan (2006)

Papain

Kaolinite

Benzoylarginine ethyl ester

þ0.9

Benesi and McLaren (1975)

Phosphatase

Kaolinite

bNaphthylphosphate

þ0.7

RamirezMartinez and McLaren (1966)

Ribonuclease

Cation exchange resin

Ribonucleic acid

Urease

Montmorillonite

Urea

þ0.6

Durand (1964)

Urease

Montmorillonite

Urea

þ0.1

Gianfreda et al. (1992)

0

0

Mitz and Summaria (1961)

Barnett and Bull (1959)

Quiquampoix and co-workers (Baron et al., 1999; Leprince and Quiquampoix, 1996; Quiquampoix, 1987; Quiquampoix et al., 1993; Staunton and Quiquampoix, 1994) have pointed out that the ‘DpH effect’ cannot adequately account for the observed upward shift of the pH-activity profile of clay-bound enzymes. Thus, when catalytic activity is expressed in absolute terms—rather than normalized to the maximum value measured for both free and bound enzymes—the activity of adsorbed enzymes falls within the range shown by their

278

Formation and Properties of Clay-Polymer Complexes

counterparts in solution. Further, DpH values are derived on the assumption that adsorption does not affect enzyme conformation. As we have seen, enzymes commonly undergo conformational changes when they interact with minerals. In summarizing the experimental results, Quiquampoix (2008a) has proposed that the apparent displacement in the pH optimum of clay-bound enzymes may alternatively be explained in terms of a pH-dependent modification of enzyme conformation (due to electrostatic interactions with the mineral surface) together with a pH-dependent change in orientation of the active (catalytic) site on the adsorbed enzyme. The DpH effect may be ascribed to the development of a ‘static’ proton gradient (between the bulk solution and the mineral–enzyme surface). Reactions catalyzed by clay-bound enzymes, such as hydrolysis of esters and amides, would also liberate protons. The resultant ‘dynamic’ proton gradient can also contribute to pH shifts. Enzyme activity is also influenced by the concentration of substrate and its diffusion from the bulk solution to the mineral surface (Tischer and Kasche, 1999). These points are diagrammatically illustrated in Figure 8.9 (Quiquampoix and Burns, 2007). H+

H+ Interfacial pH P

S S

Diffusion limitation

Orientation

Structural modification

FIGURE 8.9 Diagram showing the various factors (represented by the bottom rectangles) influencing the activity of enzymes adsorbed to clay mineral surfaces. Polygonal box represents enzyme with (indented) catalytically active site. Thin darkish rectangles depict mineral surface. S, enzyme substrate; P, reaction product. From Quiquampoix and Burns (2007).

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279

Many of the examples listed in Table 8.2 refer to systems in which carrierbound enzymes act on their substrates in solution. Such systems have received a considerable amount of attention in view of their importance and relevance to biochemical processes occurring in the living cell (Katchalski et al., 1971; McLaren and Packer, 1970). In soils, however, both substrates and enzymes are likely to be closely associated with the mineral (carrier) surface, making it difficult to distinguish between enzyme- and substrate-immobilized systems. This is especially true when the substrate on which the enzyme acts is itself a protein. The kaolinite–lysozyme–chymotrypsin assemblage (Figure 8.8) is a good example of a system in which the added enzyme (chymotrypsin) is adsorbed by the clay–lysozyme complex. Indeed, the formation of a clay–substrate–enzyme complex appears to be a prerequisite for proteolysis to occur (McLaren and Estermann, 1956). The amount of chymotrypsin adsorbed by the kaolinite–lysozyme complex (KHLC) was less than that associated with the parent, lysozyme-free mineral. The rapid decrease in enzyme activity at pH >9.6 coincided with the sharp decline in the amount of chymotrypsin adsorbed by KHLC since this pH range was far above the isoelectric point (pI8.6) of the enzyme. At pH 9.0, for example, about 95% of the added enzyme was adsorbed by KHLC, increasing to 100% at pH 8.3 (in universal buffer). The close correspondence between adsorption and activity of chymotrypsin lends further support to the notion that a clay–enzyme–substrate complex is formed before substrate decomposition occurs. In agreement with previous investigators (Ensminger and Gieseking, 1942; Lynch and Cotnoir, 1956; Mortland and Gieseking, 1952), Estermann et al. (1959) found that lysozyme complexed with montmorillonite was less susceptible to enzymatic hydrolysis (by chymotrypsin) than its kaolinite-adsorbed counterpart. This might be because most of the substrate in montmorillonite is present in the interlayer space whereas that in kaolinite is associated with external particle surfaces. Although the interlayer substrate would partly be accessible to extracellular enzymes, intercalation would impose steric constraints on the formation of an enzyme–substrate complex. In addition, the diffusion of substrates into, and of reaction products out of, the interlayer space would be retarded (Figure 8.9). This interpretation accords with experiment, at least for the system being considered here. Thus, the (initial) basal spacing of the montmorillonite– lysozyme complex was 4.6 nm, corresponding to an interlayer separation of 3.6 nm that would be sufficiently wide for the enzyme to gain entry into the interlayer space. As digestion (by chymotrypsin) progressed, this spacing decreased until at the end of the incubation period (10days) a spacing of 1.7 nm was recorded, beyond which stage little, if any, further decomposition would have occurred. Lysozyme adsorbed to kaolinite and montmorillonite surfaces was equally susceptible to being hydrolysed by cultures of microorganisms and by soil

280

Formation and Properties of Clay-Polymer Complexes

(Estermann and McLaren, 1959). Apparently, the agents responsible in this instance were a variety of exoenzymes secreted by the cultures or present in the soil. Indeed, the rate of hydrolysis was enhanced as compared with the clay-free system. The stimulating effect occurs whether or not the microorganisms were attached to the clay minerals. Thus, treatments that would dislodge the bacteria from the KHLC, such as shaking the system, did not sensibly change the rate of hydrolysis as measured by the amount of ammonia produced. On the other hand, no rate enhancement was observed with a non-adsorbed substrate, such as (denatured) lysozyme that had previously been digested by chymotrypsin. In being able to serve as a concentrating agent for both substrate and enzyme, clay minerals can accelerate substrate decomposition. In an attempt to simulate soil systems, McLaren and Estermann (1956) have examined the behaviour of chymotrypsin towards KHLC in paste form. Some of the results are shown in Figure 8.10 from which the following inferences may be drawn. Like its counterpart in suspension, adsorbed lysozyme in a paste of KHLC was susceptible to digestion by chymotrypsin at a rate that was smaller than that observed for the unadsorbed protein. For a given period of digestion, however, the amount of lysozyme hydrolysed was less in the paste than in suspension but this difference in rate tended to vanish when the initial surface coverage by the substrate (lysozyme) was smaller than the ‘monolayer’ capacity.

Increase in D280 (%)

50

63 mg

40

63 mg 1/ 2

30 61 mg 20

1/

3

65 mg

10 0

/1

Maximum D280 possible

1

60

0

20

40

100 80 60 Digestion time (min)

120

140

FIGURE 8.10 Digestion of KHLC in suspension (○) and in a paste (●) by chymotrypsin, corrected for digestion of eluted substrate. Horizontal broken lines indicate maximum possible absorbance at 280 nm (D280) for the digestion of lysozyme adsorbed to kaolinite at 1/3, 1/2 and 1/1 coverage by substrate. The amount of chymotrypsin used (in mg) is indicated on the curves. From McLaren and Estermann (1956).

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281

We might add that the mobility of chymotrypsin (over the KHLC surface) in the paste is probably more restricted than that in suspension. Likewise, the diffusion of hydrolysis products out of the complex would be faster in the paste than in suspension. As a result, there would be a rapid build-up of surface vacancies for the enzyme to occupy as compared with the system in paste form. Since the rate of hydrolysis is primarily controlled by the amount of adsorbed enzyme, a low surface coverage by the substrate would favour a more rapid breakdown of lysozyme in the complex. By the same token, the difference in hydrolysis rate between KHLC in suspension and that in paste form would diminish as the initial surface coverage by lysozyme decreases. Irrespective of coverage, however, the rate at which adsorbed proteins are digested by exoenzymes tends to reach a limiting value that is appreciably lower than the maximum attainable. This would suggest that after a given period of contact, the enzyme may ‘strike’ a strongly adsorbing site at the clay surface. At this point, the enzyme becomes more or less immobilized, leaving patches of undigested substrate. An analogous situation has been described by Morgan and Corke (1976) who noted that the specific activity of glucose oxidase in complexes with different clay minerals was influenced by the extent of enzyme adsorption. At low levels of uptake, enzyme activity was markedly suppressed as compared with that in solution. As the amount (of enzyme) adsorbed increased, its specific activity rose until at maximum adsorption it approached that of the free enzyme. In comparing the activity of catalase and pronase in the presence of kaolinite, montmorillonite and some soils, Aliev and Zvyagintsev (1974) also found that the activity of clay-adsorbed enzymes was about half that of their counterparts in solution with montmorillonite being more effective than kaolinite in suppressing enzyme activity. Like jackbean urease, adsorbed catalase could be hydrolysed by pronase but at a much lower rate than the nonadsorbed form. Since the adsorption of catalase was presumably confined to external particle surfaces (Harter and Stotzky, 1973), the observed reduction in hydrolysis rate was probably due more to conformational and surface (mobility, coverage) effects than to steric and accessibility factors. An interesting feature of clay–enzyme–substrate systems is that the order in which enzyme and substrate are added to the clay mineral can markedly influence the properties of the resultant complex. Thus, for the kaolinite– chymotrypsin–lysozyme system, referred to above, the rate of substrate hydrolysis is much lower if the addition of chymotrypsin (the enzyme) precedes that of lysozyme (the substrate) than if the sequence of addition is reversed (McLaren and Estermann, 1956). This effect lends further support to the concept that the adsorbing sites on the clay surface are non-uniform in strength and affinity towards proteins. By the same token, chymotrypsin added to the substrate-free clay shows a relatively low activity towards, and is not displaced by, lysozyme presumably because the enzyme would first occupy the

282

Formation and Properties of Clay-Polymer Complexes

TABLE 8.3 The Effect of Sequence of Enzyme and Substrate Addition on the Activity of Chitinase, in the Presence and Absence of Kaolinite. Sequence of Addition

Relative Activity (%)

(A)

Chitinþchitinaseþbuffer (control)

100

(B)

Chitinþkaoliniteþchitinaseþbuffer

15.5

(C)

Chitinaseþkaoliniteþchitinþbuffer

5.5

(D)

50% of (B)þ50% of (C)

10.6

The measurements were made under the following conditions: time, 30 min; temperature, 310 K; chitinase, 14 mg/mL; buffer, 0.03 M sodium acetate at pH 4.5; kaolinite, 7 mg/mL; chitin, 1 mg/mL. After adding the clay, the mixture was shaken end-over-end for 10 min. The structural formula of chitin is shown in the Appendix. From Skujin¸sˇ et al. (1974).

most strongly adsorbing sites. Skujin¸sˇ et al. (1974) have found similarly for the system consisting of kaolinite, chitinase and chitin (Table 8.3). It seems clear from what we have seen that the formation of an enzyme– substrate complex (at the clay mineral surface) is an essential prerequisite for substrate decomposition. In other words, the enzyme-catalysed decomposition of clay-adsorbed substrates occurs at the mineral/solution interface rather than in the surrounding bulk phase. With reference to the kaolinite–chymotrypsin– lysozyme system of McLaren and Estermann (1956), for example, only about 10% of the total enzyme activity could be ascribed to the free enzyme acting on lysozyme released from KHLC into the bulk solution. On the other hand, Pinck and Allison’s (1961) experiments would indicate that urease complexed with kaolinite and montmorillonite must be detached from the mineral surface before it could act on urea in solution. These workers, however, used a commercial, impure sample of urease, and furthermore, the ammonia produced by the reaction increased the pH of the system to levels where little urease would have been retained by the clay. Indeed, most of the enzyme initially adsorbed eventually became detached from its complex. The retention of essential, albeit modified, activity by adsorbed enzymes is, of course, fundamental to the synthesis and application of immobilized enzymes. The rate of enzyme-catalyzed reactions is frequently, but by no means invariably, reduced by the presence of clay minerals in the system. Although adsorption of either enzyme or substrate does not necessarily influence the frequency of enzyme–substrate encounter, it does undoubtedly affect the mobility and diffusion of the reactants. Thus, a decrease in reaction rate may partly be ascribed to diminished substrate accessibility to the active sites on the enzyme due to steric and conformational effects and partly to variations in orientation of mineral-bound enzymes (Doonan, 1969; Quiquampoix and Burns, 2007; Zimmerman and Ahn, 2010) as indicated in Figure 8.9.

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Proteins and Enzymes

Phosphorus released (mg/l)

100 A

80

60 B 40

20 C 0

0

0.25

0.50 0.75 Amount of clay (g)

1.0

FIGURE 8.11 Diagram showing the effect of adding different amounts of three clay mineral species on the activity of phosphatase. A, kaolinite; B, illite; and C, montmorillonite. From Mortland and Gieseking (1952).

Enzyme activity may be further reduced if the active sites (on the enzyme) are involved in the adsorption process, and if either the substrate or the enzyme is capable of penetrating the interlayer space of the clay. Because of its large surface area and exchange capacity, as well as its ability to intercalate organic compounds, montmorillonite is generally more effective in suppressing enzyme activity than kaolinite (Ensminger and Gieseking, 1941; Makboul and Ottow, 1979a–c; Mortland and Gieseking, 1952; Quiquampoix, 1987; Ross and McNeilly, 1972). Besides being influenced by the type of clay mineral, the extent of activity inhibition is dependent on the amount of mineral added (Aomine and Kobayashi, 1964a; Makboul and Ottow, 1979a–c; Mortland and Gieseking, 1952). An example of this relationship is given in Figure 8.11 and Table 8.5. Because of the many factors involved, it is difficult to compare published data on the part played by clay minerals in enzyme-catalysed reactions, unless the conditions and methods used to assess them are identical. Table 8.4 shows that the magnitude of clay-induced changes in enzyme activity can vary widely even for the same clay mineral species. Ross and McNeilly (1972), for example, observed that kaolinite had a slight depressive effect on the activity of glucose oxidase, whereas Zvyagintsev and Velikanov (1968) reported differently for a similar system. Early on, Aomine and Kobayashi (1964a,b, 1966) and Kobayashi and Aomine (1967) reported that protease activity was inhibited by both allophane and montmorillonite but more so in the presence of allophane, presumably because allophane has a larger surface area than montmorillonite (Hall et al., 1985). The extent of enzyme inactivation, however, was not

284

Formation and Properties of Clay-Polymer Complexes

TABLE 8.4 Changes in Activity of Enzymes Resulting from Their Interaction with Clay Minerals. Enzyme

Clay Mineral

Activity Change (%)

Reference

Acid phosphatase

Kaolinite


75

Ramirez-Martinez and McLaren (1966)

Acid phosphatase

Kaolinite


64

Gianfreda and Bollag (1994)

Acid phosphatase

Montmorillonite


68

Gianfreda and Bollag (1994)

Acid phosphatase

Na– montmorillonite


80

Rao et al. (2000)

Acid phosphatase

Al(OH)x– montmorillonite


42

Rao et al. (2000)

Acid phosphatase

Allophane

þ50

Allison (2006)

Acid phosphatase

Allophane

þ33

Rosas et al. (2008)

Alkaline phosphatase

Montmorillonite


67

Tietjen and Wetzel (2003)

Arylsulphatase

Kaolinite


18

Hughes and Simpson (1978)

Arylsulphatase

Ca– montmorillonite


52

Hughes and Simpson (1978)

Aspartase

Ca– montmorillonite


20

Naidja and Huang (1996)

Catalase

Ca– montmorillonite


81 to
99

Calamai et al. (2000)

Chitinase

Kaolinite


95

Skujin¸sˇ et al. (1974)

Glucose oxidase

Allophane


52

Ross and McNeilly (1972)

Glucose oxidase

Kaolinite


17

Ross and McNeilly (1972)

Glucose oxidase

Illite


21

Ross and McNeilly (1972)

Glucose oxidase

Ca– montmorillonite


77

Ross and McNeilly (1972)

Glucose oxidase

Na– montmorillonite


90

Garwood et al. (1983)

Glucose oxidase

Ca– montmorillonite


57 to
96

Morgan and Corke (1976)

Glucosidase

Kaolinite


13 to
100

Quiquampoix (1987) Continued

Chapter

8

285

Proteins and Enzymes

Glucosidase

Na– montmorillonite


35 to
100

Quiquampoix (1987)

b-Glucuronidase

Kaolinite

þ81

Fiorito et al. (2008)

b-Glucuronidase

Montmorillonite

þ72

Fiorito et al. (2008)

Invertase

Na– montmorillonite


88 to
96

Gianfreda et al. (1991)

Invertase

Al(OH)x– montmorillonite


89 to
95

Gianfreda et al. (1991)

Laccase

Bentonite


11 to
100

Claus and Filip (1988)

Laccase

Kaolinite


30 to
83

Claus and Filip (1988)

Laccase

Kaolinite


14

Gianfreda and Bollag (1994)

Laccase

Montmorillonite

0

Gianfreda and Bollag (1994)

Mixed endopeptidases

Montmorillonite


66

Haska˚ (1981)

N-acetylglucosaminidase

Allophane

þ40

Allison (2006)

Polyphenoloxidase

Allophane

þ55

Allison (2006)

Peroxidase

Ca– montmorillonite

0 to
69

Lozzi et al. (2001)

Peroxidase

Na– montmorillonite


88 to
99

Lozzi et al. (2001)

Protease

Montmorillonite


100

Tietjen and Wetzel (2003)

Tyrosinase

Bentonite


84 to
100

Claus and Filip (1988)

Tyrosinase

Kaolinite

0 to
75

Claus and Filip (1988)

Tyrosinase

Al(OH)x– montmorillonite


24 to
62

Naidja et al. (1997)

Urease

Na– montmorillonite


41

Gianfreda et al. (1992)

Urease

Al(OH)x– montmorillonite


49 to
67

Gianfreda et al. (1992)

Modified from Zimmerman and Ahn (2010).

286

Formation and Properties of Clay-Polymer Complexes

proportional to the amount adsorbed, reflecting reduced accessibility of the active sites on the enzyme to the respective substrate. The discrepancy between the amount adsorbed and the extent of inhibition was particularly marked for b-amylase. The reason for this behaviour was not clear. Since the b-amylase preparation used was apparently polydisperse, the enzymically active fraction might have been preferentially adsorbed by the clay minerals. This suggestion may also explain why the activity of b-amylase, unlike that of protease, was more strongly inhibited by montmorillonite than by allophane (Table 8.5). In common with the kaolinite–lysozyme–chymotrypsin and kaolinite– chitin–chitinase systems (Table 8.3), the activity of b-amylase in the presence of clay minerals relative to that of the enzyme in solution, depended on the order in which the enzyme and the substrate were added to the clay (Aomine and Kobayashi, 1966). In this instance, the activity was less inhibited when the substrate (starch) was added to the clay before the enzyme than when the sequence of addition was reversed. This is another illustration of the influence of enzyme and substrate affinity for the mineral surface, and their relative surface requirement, on reaction rates. Similarly, Ross and McNeilly (1972) reported that the activity of glucose oxidase was much more depressed by montmorillonite than by allophane. Morgan and Corke (1976) subsequently noted that the specific activity of glucose oxidase, attached to montmorillonite, increased with the amount adsorbed, approaching that of the free enzyme at maximum uptake. This is perhaps hardly surprising since enzyme–substrate complex formation, as we will see later, must precede substrate decomposition (product formation). We may thus include surface concentration of enzyme in the list of factors affecting the activity of immobilized enzymes. Allison (2006) and Rosas et al. (2008), on the other hand, found that the activity of some enzymes (e.g. acid phosphatase) was enhanced—rather than depressed—by adsorption to allophane (Table 8.4). This would suggest a favourable orientation of active sites on the enzyme and a lessening of substrate and product diffusion limitation (Zimmerman and Ahn, 2010). Alternatively, the high propensity of allophane for adsorbing and stabilizing both enzyme and substrate would facilitate enzyme–substrate complex formation. This is especially true of phosphate-containing substrates since such compounds are strongly adsorbed by ligand-exchange involving amphoteric Al (OH)(H2O) groups, exposed at defect sites along the walls of nanosize allophane spherules (Hashizume and Theng, 2007; Theng et al., 1982; cf. Figure 1.18). The reaction kinetics of adsorbed enzymes may not conform to the Michaelis–Menten type in that the initial rate is not linearly related to the total enzyme concentration in the system. Rather, it is proportional to the surface area of the (solid) carrier in contact with the solvent. For somewhat different reasons, described below, a departure from Michaelis–Menten kinetics has

a

a-Amylase

Haemoglobin

b

b

b-Amylase

b

Cellulase

Amylase

Soluble starch

þ

Na -carboxy-methylcellulose

Enzyme Added (mg)

Clay Added (mg)

Relative Activity (%) Allophane

Halloysite

Montmorillonite

0.1

0.3

62.9

93.8

81.0

0.1

0.6

51.7

0.1

0.9

44.3

0.1

1.2

42.0

0.05

0.08

86.2

88.1

87.7

0.1

0.3

76.8

78.2

78.9

0.1

0.3

27.3

64.6

15.5

0.1

1.5

13.2

45.7

8.7

0.1

0.3

85.6

99.3

97.2

0.2

3.0

83.9

98.3

88.6

Proteins and Enzymes

Protease

Substrate

8

Enzyme

Chapter

TABLE 8.5 Influence of Some Soil Clays on the Relative Activity of Different Enzymes, Taking the Activity of the Clay-Free System as 100%.

a

pH 5.0. pH 5.6.

b

From Aomine and Kobayashi (1964a).

287

288

Formation and Properties of Clay-Polymer Complexes

been reported by Irving and Cosgrove (1976) for soil acid phosphatase acting on p-nitrophenylphosphate. In most instances, however, enzyme-catalyzed reactions involving clay minerals, soils and organic matter appear to follow Michaelis–Menten kinetics. The (apparent) Michaelis constant (Km) is indicative of enzyme–substrate affinity in that the higher the value the lower is the affinity, while the maximum velocity (Vmax) of the reaction describes the rate of substrate conversion at saturation of the enzyme’s active sites by the substrate. The data of Table 8.6 show that Km values for reactions catalyzed by clay-adsorbed enzymes are commonly higher, while Vmax values are lower,

TABLE 8.6 Changes in Michaelis Constant (Km) and Maximum Reaction Velocity (Vmax) of Some Enzymes in the Presence of Different Clay Minerals. Enzyme

Clay Mineral

Kma

Vmaxa

Reference

Acid phosphatase

Montmorillonite

þ5779


46

Makboul and Ottow (1979a)

Acid phosphatase

Illite

þ500


56

Makboul and Ottow (1979a)

Acid phosphatase

Kaolinite

þ443


69

Makboul and Ottow (1979a)

Acid phosphatase

Montmorillonite

0


38 to
67

Dick and Tabatabai (1987)

Acid phosphatase

Illite

0


21 to
46

Dick and Tabatabai (1987)

Acid phosphatase

Kaolinite

þ68 to þ307

0

Dick and Tabatabai (1987)

Acid phosphatase

Allophane


14

þ42

Rosas et al. (2008)

Alkaline phosphatase

Montmorillonite

þ85

þ9

Makboul and Ottow (1979b)

Alkaline phosphatase

Illite


8


62

Makboul and Ottow (1979b)

Alkaline phosphatase

Kaolinite

þ42


3

Makboul and Ottow (1979b)

a-Amylase

Bentonite

þ400


620

Sedaghat et al. (2009a)

b-Glucosidase

Bentonite

þ45


98

Sarkar and Burns (1984) Continued

Chapter

8

289

Proteins and Enzymes

b-Glucosidase

Al(OH)x– montmorillonite


48


36

Sarkar and Burns (1984)

b-Glucosidase

Montmorillonite


60


3

Serefoglou et al. (2008)

Protease

Montmorillonite


55


66

Kobayashi and Aomine (1967)

Protease

Allophane


4


56

Kobayashi and Aomine (1967)

Pyrophosphatase

Montmorillonite

0


39 to
48

Dick and Tabatabai (1987)

Pyrophosphatase

Illite

0


18 to
41

Dick and Tabatabai (1987)

Pyrophosphatase

Kaolinite

þ71 to þ138

0

Dick and Tabatabai (1987)

Urease

Montmorillonite

þ191


49

Makboul and Ottow (1979c)

Urease

Illite

þ88


50

Makboul and Ottow (1979c)

Urease

Kaolinite

þ161


48

Makboul and Ottow (1979c)

a

Expressed as percent change relative to the free enzyme.

Modified from Zimmerman and Ahn (2010).

as compared with the corresponding free enzymes (in a homogeneous solution). In its simplest form, the classical model for an enzyme-catalysed reaction may be presented by the following scheme k1

k3

E þ S Ð ES ! E þ P k2

ð8:6Þ

where E, S, ES and P refer to the enzyme, substrate, enzyme–substrate complex, and reaction products, respectively; k1, k2 and k3 are the corresponding velocity constants of the assumed processes. Km is equated with the ratio k2þk3/k1, but for k2>>k3, Km k2/k1 and represents the thermodynamic dissociation constant of ES. The relationship between Km, E and S, expressed in terms of reaction velocities, is given by the Michaelis–Menten equation u ¼ V max ðSÞ=Km þ ðSÞ;

ð8:7Þ

where u and Vmax are the measured initial and maximal velocity, respectively, and (S) is the substrate concentration. Since for u ¼ ½Vmax, Km is numerically

290

Formation and Properties of Clay-Polymer Complexes

equal to (S), Km has the dimension of concentration. A number of linear transformations of Equation (8.7) have been proposed to facilitate determination of Vmax and Km (Dowd and Riggs, 1965). Among these, the double reciprocal form due to Lineweaver and Burk (1934) 1=u ¼ ðKm =Vmax Þ  ð1=ðSÞÞ þ ð1=Vmax Þ

ð8:8Þ

and the single reciprocal form due to Eadie and Hofstee u ¼ ðVmax
Km Þðu=ðSÞÞ

ð8:9Þ

have been widely used. More often than not, the apparent Michaelis constant of a carrier-bound 0 ) acting on a (charged) substrate differs from the value of the enzyme (Km corresponding enzyme in a homogeneous solution (Km) due to the unequal distribution of substrate species between the polyelectrolyte surface and the bulk solution (e.g. Katchalski et al., 1971). When the charge on the carrier–enzyme complex is opposite to that on the substrate, the concentration of substrate near the surface, (S)s, is greater than that in the bulk solution, (S)b. Since Vmax is then obtained at a lower (S)b value as compared with the carrier-free system, Km0 Km. the substrate is of the same sign, (S)s<(S)b, and hence Km 0 The relationship between Km , Km and the electrostatic potential, c, is completely analogous to Equation (8.1) relating pH to c; that is, 0 DpKm ¼ pKm
pKm ¼ 0:43zec=kT

ð8:10Þ

Clearly, Equation (8.10) does not apply to a situation where either the carrier or the substrate is uncharged, in which case Km0 ¼Km, provided that diffusion effects are negligible. However, if the diffusion of substrate to the carrier– enzyme/solution interface is retarded so that (S)s is effectively less than (S)b, the following empirical relationship is applicable 0 Km ¼ Km þ Vmax ðt=DÞ;

ð8:11Þ

where t is the diffusion layer thickness and D the diffusion coefficient. Diffusion effects may be important with high molecular weight substrates. The validity of the above analyses has been substantiated by experimental measurements using enzymes attached to polymer matrices, acting on their respective substrates (Hornby et al., 1966, 1968; Ladd and Butler, 1975). Since clay–enzyme complexes are often negatively charged, the direction 0 will change with respect of Km can, in principle, be predicted in which Km from the charge characteristics of the substrate. In this connection, it is relevant to mention Benesi and McLaren’s (1975) data comparing the activity of papain in solution with that of the kaolinite-bound enzyme. Requiring a free thiol (SH) group, the activity of papain depends on the redox potential of the system and, hence, on the disulphide/thiol ratio, r. For a redox pair, such as þR

S

S

Rþ and þR

SH, the former would be more strongly

Chapter

8

291

Proteins and Enzymes

attracted to the negatively charged clay surface. The approach used was to determine enzyme activity (under nitrogen) for papain in solution at pHb 6.1 and when adsorbed to kaolinite at pHb 6.9, in the presence of different disulphide–thiol mixtures at various values of r. One mixture (A) consisted, for example, of dithiodiglycol and b-mercaptoethanol, both of which were uncharged; a second mixture (B) of dithioglycol and b-aminoethylmercaptan (monovalent cation); a third (C) of cystine ethyl ester (divalent cation, disulphide) and cysteine ethyl ester (monovalent cation, thiol). Figure 8.12 shows that in the presence of pair (A), the curve for papain in solution coincided with that of the adsorbed enzyme. This would indicate that although the amount of active, reduced SH-enzyme clearly depended on r, this ratio was apparently the same in solution as near the clay–enzyme complex surface. On the other hand, the relative activity of the kaolinitebound enzyme for pair (B) was markedly reduced as compared with the corresponding free enzyme. This difference was more pronounced for pair (B) because the concentration of þR

S

S

Rþ near the surface was increased over and above that of þR

SH. It must be borne in mind, however, that enzyme reactions at clay and soil mineral surfaces are influenced by factors other than electrostatic (charge– charge) interactions. Indeed, steric, conformational and diffusion effects 100 A 75 50 25 Relative activity (%)

FIGURE 8.12 Relative activity of papain (with benzoylarginine ethyl ester) as a function of the oxidizing disulphide/reducing thiol ratio. (○), Papain in solution (pHb¼6.1, ionic strength0.07); (●), papain adsorbed to kaolinite (pHb¼6.9, ionic strength0.07). (A) Dithiodiglycol and b-mercaptoethanol. (B) Dithiodiglycol and b-aminoethylmercaptan. From Benesi and McLaren (1975).

0

20

40

60

80

¥

100

100 B 75 50 25 0

20

40 60 80 100 R = [disulphide]/[thiol]

¥

292

Formation and Properties of Clay-Polymer Complexes

may be of overriding importance. We have already mentioned that the hydrolysis of lysozyme, complexed with kaolinite, by bacterial exoenzymes was stimulated rather than retarded as compared with the clay-free system. An early attempt at measuring reaction rates and apparent Michaelis constants of enzymes in the presence of clay minerals is that by Kobayashi and Aomine (1967) who reacted allophane and montmorillonite suspended (in a buffer solution) with the enzyme before adding the appropriate substrate to the clay–enzyme ‘mixture’ or the clay–enzyme ‘complex’. The latter referred to the material after centrifuging the mixture and washing the precipitate with sodium acetate buffer (pH 5) to remove any unadsorbed and weakly attached enzyme. Figure 8.13 shows the results for pronase (a protease extracted from Streptomyces griseus) acting on a dipeptide (carboxybenzyl-L-glutamyl-Ltyrosine (CBGT)), in the absence and presence of clay. As might be expected, the adsorbed enzyme obtained maximal velocity at a lower substrate concentration as compared with its free counterpart in solution. The effect of montmorillonite on the reaction velocity was greater than that of allophane. The average Vmax and Km values derived from Figure 8.13 are given in Table 8.6. These refer to the hydrolysis of CBGT by the clay–enzyme ‘mixture’ and hence would include the contribution from any unadsorbed enzyme. Interestingly, the montmorillonite–pronase ‘complex’ (washed) showed almost no enzymic activity towards CBGT whereas the corresponding ‘complex’ with allophane retained most of its initial capacity to hydrolyse the dipeptide. On this basis, Kobayashi and Aomine (1967) inferred that the activity of the unwashed montmorillonite–enzyme system was due to pronase adsorbed to external particle surfaces, and to some extent, to the free enzyme both of which were presumably removed during washing. If a proportion of the adsorbed pronase were present in the interlayer space, the intercalated enzyme was evidently incapable of forming an enzyme–substrate complex. FIGURE 8.13 Relationship between initial reaction rate (v) and substrate concentration in the bulk solution (Sb) for the hydrolysis of carbobenzoxyL-glutamyl-L-tyrosine by protease. Curve A: hydrolysis reaction in solution (control). Curve B: reaction in the presence of allophone. Curve C: reaction in the presence of montmorillonite. From Kobayashi and Aomine (1967).

5.0

v ´ 106 (mol/min)

A B 2.5 C

0

0

2.5 (S)b ´ 103 (mol/l)

5.0

Chapter

8

Proteins and Enzymes

293

On the other hand, most of the allophane-bound pronase was retained against washing, in keeping with Milestone’s (1971) finding that proteins are tenaciously held by allophane. The influence of diffusion effects on Km (cf. Equation 8.11) may be illustrated by the measurements of Usame and Inoue (1974). Using glucoamylase attached to an acid clay and soluble starch as substrate, these workers found 0 was 5–6 times greater than Km for a high molecular weight (8103 that Km Da) substrate. On the other hand, the apparent Michaelis constant for the bound enzyme, acting on a low molecular weight starch, was similar in magnitude to that of its free counterpart. Steric effects may further reduce the rate of enzyme–substrate complex formation for high molecular weight (uncharged) substrates. Kinetic studies of enzyme action in the presence of clays (and soils) thus lend further support to the concept that substrate adsorption by the clay–enzyme derivative precedes its hydrolysis or digestion. An example where the apparent Michaelis constant of an enzyme (soil acid phosphatase) acting on its substrate is measured in conjunction with substrate (p-nitrophenyl phosphate) adsorption by the soil–enzyme system has been described by Cervelli et al. (1973). To this end, they took whole soils (rather than the respective clay fractions) and further assumed that only the fraction of the total substrate left in solution was available to the enzyme. For all four 0 kaolinite>illite, whereas the reduction in Vmax values was similar for all three minerals (Table 8.6). Likewise, Parks (1974) found that for an alkaline phosphatase (from Escherichia coli) acting on p-nitrophenyl phosphate, the apparent Michaelis

294

Formation and Properties of Clay-Polymer Complexes

constant for the clay–enzyme system was generally higher than for the corresponding (free) enzyme in solution. The clay minerals used were unfractionated, raw samples of a kaolinite, an illite, a montmorillonite and a palygorskite (‘attapulgite’). Reaction rates were measured in the presence of 1 M Tris buffer at pH 8 coinciding with the pH optimum of the enzyme. Under these conditions, the concentration of substrate near the surface, (S)s, was presumably less than that in the bulk solution, (S)b, so that the adsorbed alkaline phosphatase reached maximal velocity at a higher bulk substrate concentration than the enzyme in a homogeneous solution. Because of the many variables that come into play when clay-adsorbed enzymes act on their respective substrates, it is not surprising that the change in the apparent Michaelis constants and maximum reaction velocities of clayadsorbed enzyme reactions varies widely even for the same clay mineral species. For the same reason, it is difficult to predict the direction to which Km and Vmax will be shifted with respect to the corresponding values for the free enzymes (Table 8.6).

8.5. ENZYME BEHAVIOUR IN SOIL To round off our discussion on the clay–enzyme interaction, we wish to make a few comments about the behaviour of enzymes in soil so as to establish a link between the abiotic chemically definable system, described in the preceding section, and the ‘living’ system that soil essentially is. Although clay minerals play an active role in degradative and synthetic reactions involving organic substances in soil (Huang, 1990, 2008), there is compelling evidence to indicate that decomposition processes in soil are largely catalyzed by extracellular enzymes (Aliev et al., 1976; Allison, 2006; Hofmann and Hoffmann, 1966; Kiss et al., 1975; Skujin¸sˇ, 1967, 1976; Voets and Dedeken, 1966; Zimmerman and Ahn, 2010). Since extracellular enzymes, as indeed the microorganisms themselves, are closely associated with the inorganic colloidal fraction, their activity is often related to the type of clay mineral that is dominantly present in the soil system (Burns, 1990; Durand, 1965; Estermann and McLaren, 1959; Marshall, 1975; Skujin¸sˇ, 1967; Stotzky, 1972; Theng and Orchard, 1995). It is generally accepted that microorganisms, both living (proliferating) and dead, are the primary source of extracellular enzymes in soil whose distribution is far from uniform. Conventional biochemical techniques for measuring enzyme activities in soil, however, do not normally discriminate between extra- and intracellular species. Nevertheless, we may reasonably expect to find a positive correlation between enzyme activity and microbial numbers/ activity. Other data, however, tend to support the view that enzyme activity bears little, if any, relationship to either microbial population as a whole or to the number of microbes in each of the main groups of microorganisms. Similarly, experimental results on the extent to which seasonal factors affect

Chapter

8

Proteins and Enzymes

295

the dynamics and accumulation of soil enzymes are not always concordant. The same can be said about the effect of clay mineral addition on enzyme activity in soil (Table 8.4). These and other apparent inconsistencies in the literature on enzyme behaviour in soil are perhaps to be expected since microbial numbers are influenced by climatic and soil conditions, and in the case of cultivated soils, by management practices as well (Ajwa et al., 1999; Burns, 1982; Frankenberger and Dick, 1983; Kandeler et al., 1999; Marx et al., 2005; Miller and Dick, 1995; Paulson and Kurtz, 1969; Perez Mateos and Gonzalez Carcedo, 1985; Skujin¸sˇ, 1976; Taylor et al., 2002). The addition of substrates to soil tends to stimulate production of the appropriate enzymes. Thus, the microbial synthesis of amylase (Drobnik, 1955), dextranase (Dra˘gan-Bularda and Kiss, 1972), levan sucrase (Kiss, 1961) and xylanase (Srensen, 1955) in soil has been shown to increase in the presence of their corresponding substrates. Interestingly, Kiss (1958a) found that the addition of kaolinite and montmorillonite to soil did not affect the soil’s natural invertase activity. When these minerals were added together with sucrose, however, there was a marked stimulation of enzyme activity (Kiss, 1958b). This would suggest that sucrose induced the microbial synthesis of invertase, the secreted extracellular portion of which was stabilized by adsorption to the (added) clay. Similarly, Srensen (1969, 1972) reported that the simultaneous addition of montmorillonite and carbohydrates to soil led to an appreciable increase in activity of the appropriate enzymes. Using 14C-labelled substrates, he was able to recover an appreciable proportion (6–12%) of the carbohydrate carbon in the form of amino acids during the first 10–30 days of incubation. In the presence of montmorillonite (5% w/w), the amount of recoverable amino acids was substantially increased, indicating that part of the added carbon was rapidly transformed into enzyme proteins that were then protected from proteolysis by adsorption to the mineral surface. Because of their intimate association with mineral and humus nanoparticles in soil (Theng and Yuan, 2008), enzymes are difficult to extract from the soil mass. Thus, only a few enzymes have been isolated from soils and then mostly in combination with inorganic and organic colloids. Briggs and Segal (1963), for example, separated a mixture of proteins with urease activity, and Martin-Smith (1963) one that was active towards uric acid. Subsequently, Bartha and Bordeleau (1969) obtained a preparation with oxidase and peroxidase activities towards aniline and o-anisidine, respectively, while Chalvignac and Mayaudon (1971) reported the extraction of an enzyme system capable of converting tryptophan into indole acetic acid. Similarly, Ladd (1972) succeeded in extracting a peptidase from soil, and Satyanarayana and Getzin (1973) an esterase that decomposed malathion. There is evidence to suggest that extracellular enzymes in soil are associated more with the organic matter (humus) fraction than with mineral colloids (Boyd and Mortland, 1990; Burns, 1986; Cacco and Maggioni, 1976;

296

Formation and Properties of Clay-Polymer Complexes

Chalvignac and Mayaudon, 1971; Ladd, 1972; Ladd and Butler, 1975; Pettit et al., 1976). Thus, the urease-active material extracted from soil by Burns et al. (1972a,b) and by McLaren et al. (1975) was identified with an enzyme–humus conjugate, having 20–40% of the total activity of the soil. Similar yields were reported by Nannipieri et al. (1974) using a soil which, incidentally, contained the same number of microorganisms after extraction. The association between urease and humus, however, was apparently nonuniform in that some fractions of soil organic matter displayed greater urease activity than others (McLaren et al., 1975). Unlike its free counterpart, the humus-bound urease was resistant to proteolysis by pronase (McLaren and Puk¸ite, 1975) presumably because the proteolytic enzyme was, on steric grounds, prevented from entering the pores of the humic material. The association of extracellular urease with organic matter in an alpine humus soil was also suggested by Lloyd (1975). Likewise, the diphenol oxidases extracted by Mayaudon et al. (1973a,b) and Mayaudon and Sarkar (1974) were largely complexed with humus, while the esterase obtained by Satyanarayana and Getzin (1973) appeared to be a carbohydrate–protein derivative. This is not to say, however, that the mineral fraction in soil plays only a minor part in complex formation with extracellular proteins and enzymes. Even if most of the enzymatically active preparations extracted from soil are complexed with humus, such materials would be indirectly bound to the mineral constituents since humic substances in soil are closely associated with the clay fraction (Oades, 1989; Tate and Theng, 1980; cf. Chapter 12). In the above-mentioned instances, the clay–humus association might be weak as compared with that between enzyme and humus. The methods used to extract enzymatically active substances from soil could conceivably release the humus–enzyme conjugates from their respective mineral (clay) complexes, as Mayaudon et al. (1973a,b) have suggested. Another piece of evidence for the involvement of the inorganic components of soils, notably the clay fraction, in binding (soil) enzymes, is provided by soil fractionation studies. Quite early on, Haig (1955) found that in a fine sandy loam, esterase activity, as measured by its effect on phenylacetate, decreased in the order clay>silt>>sand. Indeed, the enzyme was so strongly attached to the (montmorillonitic) clay fraction that it effectively resisted elution by organic bases and phosphate or separation from its clay complex by sonic treatment. Hoffmann (1959) reported similarly for some carbohydrates and urease. The virtual absence of microorganisms from the clay fraction further indicated that the clay-bound enzymes were largely extracellular. Likewise, Galstyan et al. (1968) and Galstyan and Havoundjian (1970) showed that the activity of various enzymes in the surface horizon of two genetically different soils decreased with an increase in mean particle size, being concentrated in the silt and clay fractions and absent from sand particles. They also noted that enzyme activity was positively correlated with the humus content of the respective particle-size fractions. Stemmer et al.

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(1998) concluded similarly for invertase activity in four different soils, while Acosta-Martı´nez and Tabatabai (2001) found that arylamidase activity in 26 surface soils was significantly correlated with organic carbon, total nitrogen and clay contents. In fractionating a series of soils by ultrasonic dispersion in a dense liquid, Ross (1975) and Speir (1977) observed that although the ‘light’ fraction was enriched in a number of enzymes, most of the activity was in the residue. This was in accord with the results of Aliev et al. (1976) who found large amounts of (soil) particle-bound extracellular enzymes which could not be released into solution by ultrasonic treatment. The strong attachment of enzymes to the inorganic soil colloids was also inferred by Dalal (1975) and Khaziev (1975) from measurements of the enthalpies and entropies of activation. Kuprevich and Shcherbakova (1971) have suggested that the total enzyme activity of a soil may be characteristic of soil type. This concept has important implications for both soil classification as well as forensic science since the detection limits of enzymes are much lower than of any other organic compound in soil. The forensic aspect of soil enzymology has been examined by Thornton et al. (1975) who found that not only were enzyme activity levels diagnostic of geographical locality but soils collected from adjacent sites within a given location could be characterized by their individual enzyme pattern. The value of enzyme assay for forensic purposes may be considerably enhanced by determining the apparent Michaelis constant, Km, of the enzyme-catalyzed reaction since this parameter is independent of sample size. When the sensitivity of the assay is further increased by using radioactive substrates, Km ratios for various substrates can serve as soil markers (‘fingerprints’). The forensic comparison of soils, based on enzyme activity measurements, however, has been superseded by bacterial DNA profiling (Heath and Saunders, 2006; Horswell et al., 2002). We have already seen that clay minerals may either inhibit, enhance or exert no measurable influence on enzyme activity although the inhibitory effect is the more common (Table 8.4). This observation also extends to soils but the interpretation of experimental data is more difficult and equivocal than for the corresponding clay–enzyme systems because biochemical processes in soil are dynamic and consecutive, rather than steady state, in nature. The Michaelis constant for an enzyme-catalyzed reaction in a dynamic column is often appreciably higher than in a static batch situation because of diffusion and surface charge effects (McLaren, 1972, 1975). During the infiltration of a soil column by a substrate solution, for example, the ratio of substrate-to-product concentration (cf. Equation 8.6) depends on the position within, and the time of passage through, the column. In other words, the process has a vectorial as well as a temporal component (McLaren, 1970). In addition, a particular compound present in, or added to, soil may undergo a series of consecutive transformations not all of which may be mediated by extracellular enzymes, as exemplified by the sequence

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urea!NH4þ!NO2
!NO3
!N2O!N2. The denitrification step, by which nitrate is transformed into nitrous oxide and then dinitrogen, has received particular attention in terms of both experimental and theoretical considerations because nitrous oxide is a potent greenhouse gas (Ardakani et al., 1973, 1974, 1975; de Klein et al., 2003; Groffman and Tiedje, 1989; McLaren, 1969; McLaren and Ardakani, 1972; Zumft, 1997). Finally, we wish to briefly mention the use of clay-immobilized enzymes in converting organic pollutants in soil into less toxic or non-toxic metabolic products. Although there are advantages in using cell-free enzymes for bioremediation, such materials are costly to isolate and purify, have short-term stability, and are easily denatured in the ‘hostile’ soil environment. On the other hand, enzymes immobilized on clay and mineral supports are resistant to denaturation, high temperature, pH and prolonged storage. More importantly, many immobilized enzymes retain their activity, and can be reused, after several cycles of operation (Dura´n and Esposito, 2000; Gianfreda and Rao, 2004; Shen et al., 2002). Sarkar et al. (1989), for example, found that soluble laccase and glucose oxidase were essentially inactivated after 15 days contact with soil (suspensions), whereas their clay- and soil-immobilized counterparts remained active under the same conditions. Ruggiero et al. (1989) also observed that the efficiency of a laccase (from the fungus Trametes versicolor), immobilized on kaolinite, in removing 2,4-dichlorophenol (2,4-DCP) from solution was similar to that of the free enzyme and greater than that of the montmorillonite– enzyme complex. Further, the immobilized laccase could be separated from the reaction mixture and reused without appreciable loss on activity. Similarly, Gianfreda and Bollag (1994) reported that laccase and horseradish peroxidase, immobilized on montmorillonite, retained much of its activity towards the removal (oxidation) of 2,4-DCP over several cycles of incubation, both in the presence and absence of soil (Figure 8.14). Like laccase, tyrosinase, immobilized on clay minerals (through adsorption or covalent bonding), could oxidize a range of phenols and amines (Claus and Filip, 1990; Sarkar et al., 1989). We might also add that the substrate (e.g. 2,4DCP) and the reactive intermediates formed during the oxidation process may be detoxified by polymerization or incorporation into humic substances (Ahn, et al., 2002; Lassen et al., 1994; Ruggiero et al., 1996; Sarkar et al., 1988). The environmental and industrial applications of laccases and tyrosinases, immobilized on various supports, have been reviewed by Dura´n et al. (2002). Laccase can also catalyze the (indirect) oxidation of compounds other than phenols and amines in the presence of a mediator such as 2,2-azino-bis(3ethylbenzothiazoline-6-sulphonic acid) (ABTS) acting as an ‘electron shuttle’ between enzyme and substrate. Thus, Dodor et al. (2004) observed that the kaolinite-immobilized laccase–ABTS system was capable of oxidizing more than 80% of added anthracene and benzo[a]pyrene after 24h of incubation, retaining the same activity over several cycles of operation. More recently,

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FIGURE 8.14 Diagrams showing the removal of 2,4-dichlorophenol (2,4-DCP) by montmorilloniteimmobilized laccase and peroxidase in different buffer solutions with and without addition of 100 mg soil material. Laccase: 24 h cycle for 1 mM 2,4-DCP in 0.1 M citrate-phosphate or phosphate buffer at pH 6. Peroxidase: 2 h cycle for 1 mM 2,4-DCPþ1 mM H2O2 in 0.1 M citrate-phosphate or phosphate buffer at pH 6. From Gianfreda and Bollag (1994).

100 Montmorillonite-Laccase 2,4-DCP removal (%)

Chapter

80 60 40 20 0

1

2

3

4

5

6

7

100 2,4-DCP removal (%)

Montmorillonite-Peroxidase 80

Phosphate buffer Citrate-phosphate buffer

60

Phosphate buffer + soil Citrate-phosphate buffer + soil

40 20 0

1

2

3

4

5

6

Cycles

Acevedo et al. (2010) reported that manganese peroxidase (from Anthracophyllum discolor), immobilized on allophane nanoparticles, was effective in degrading polycyclic aromatic hydrocarbons. Indeed, the immobilized enzyme was more active than its free counterpart in the transformation of anthracene in soil.

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