ARTICLE IN PRESS

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doi:10.1016/j.so

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Soil Biology & Biochemistry 39 (2007) 493–504

www.elsevier.com/locate/soilbio

Fate of prions in soil: Interaction of a recombinant ovine prion proteinwith synthetic humic-like mineral complexes

Maria A. Raoa,�, Fabio Russoa, Vincenzo Granatab, Rita Berisioc,Adriana Zagarib, Liliana Gianfredaa

aDipartimento di Scienze del Suolo, della Pianta e dell’Ambiente, Universita di Napoli Federico II, Via Universita 100, I-80055 Portici, ItalybDipartimento delle Scienze Biologiche, and CNISM, Universita di Napoli Federico II, Via Mezzocannone 16, I-80134 Napoli, Italy

cIstituto di Biostrutture e Bioimmagini, C.N.R., Via Mezzocannone 16, I-80134 Napoli, Italy

Received 31 May 2006; received in revised form 4 August 2006; accepted 18 August 2006

Available online 2 October 2006

Abstract

Prion proteins are regarded as the main agents of transmissible spongiform encephalopathies. Understanding their fate in soil may be

crucial to elucidate the dissemination of the prion in the environment, associated with a possible transmission of infectivity. Studies were

performed with simplified model systems, derived by the birnessite-mediated oxidative polymerization of catechol, which simulate

processes naturally occurring in soil. A benign full-length recombinant purified ovine protein (PrP) (residues 23–234), as well as a

truncated-form ovPrP (tPrP) (103–234) were utilized. Catechol and prion protein interacted under experimental conditions that

reproduced the interaction of PrP with soluble organic matter or with insoluble organic matter during or after formation of catechol

polymers. PrP stability in all buffers and chemicals was preliminarily monitored by circular dicroism (CD) measurements. The

disappearance of protein molecules from the solution, the decrease of UV–Visible absorbance of supernatants, and the FT-IR spectra

and the elemental analyses of solid-phase residues indicated that both PrP and tPrP were involved in catechol polymerization by

birnessite. Furthermore, a clear flocculation of soluble catechol–protein polymeric products in solid aggregates was observed when PrP

was added to the supernatants. Different kinds of extracting agents were not able to desorb/extract PrP as well as tPrP from the formed

solid aggregates, thereby highlighting the high stability of protein–organic and –organo-mineral complexes.

r 2006 Elsevier Ltd. All rights reserved.

Keywords: Prions; Catechol; Birnessite; Oxidative polymerization; Humic substances; Adsorption; Entrapment; Extraction; Desorption

1. Introduction

Recently, a growing concern is dedicated to the fate ofprion proteins in soil (Revault et al., 2005; Vasina et al.,2005; Johnson et al., 2006; Leita et al., 2006; Rigou et al.,2006). According to ‘‘the protein-only hypothesis’’ byPrusiner (1991, 1998) prion protein, a solely endogenouscellular protein, may be considered the main ‘‘infectiousagent’’ of a group of transmissible spongiform encephalo-pathies (TSE). In the pathogenesis the protein is alteredfrom the a-helix-rich host prion protein (PrPC) into apathogenic isoform (PrPSc), insoluble, partially resistant to

e front matter r 2006 Elsevier Ltd. All rights reserved.

ilbio.2006.08.020

ing author. Tel.: +390812539179; fax: +39 0812539186.

ess: mariarao@unina.it (M.A. Rao).

K-protease and with a high content in b-sheet structure(Prusiner, 1991, 1998).Soil is a natural sink of several contaminants which may

also include prion proteins. Dispersion of prion proteins insoil occurs from meat and bone meal storage plants, use offertilizers augmented with meat and bone, decompositionof TSE-contaminated animal carcasses buried in soil, TSE-infected tissues, liquid and solid waste fragments fromabattoirs, central sterilization units, operating theatres andwastes from TSE-infected animals (Woolhouse et al., 1998;Andreoletti et al., 2002). Therefore, understanding theretention and/or the dissemination of prion proteins insoils may be important for dealing with the disseminationof TSE infectivity. A study by Brown and Gajdusek (1991)demonstrated that ovine prion protein conserved itsinfectivity after 3 years internment in soil.

ARTICLE IN PRESSM.A. Rao et al. / Soil Biology & Biochemistry 39 (2007) 493–504494

Once introduced into the soil a protein may remain freein solution, and as such to be biologically or chemicallydegraded, be associated with preformed organic andinorganic soil components by adsorption/entrapment, orparticipate in the formation of organic and organo-mineralsoil complexes. Relatively stable protein–soil colloidcomplexes may be formed. A strong retention andaccumulation of the protein on the formed complexes aswell as its release can occur under some specific environ-mental conditions.

As demonstrated by several studies (Ladd and Butler,1975; Burns, 1986; Boyd and Mortland, 1990; Quiquam-poix, 2000; Gianfreda et al., 2002) profound structuralalterations and conformational changes of protein mole-cules associated to soil colloids may take place withvariations of the protein functionality. Structural molecu-lar alterations may also result upon protein release fromthe soil aggregates under some environmental conditions.As a consequence, the interaction of prion proteins withsoil complexes or their release from these latter maybecome an important step in the risk of prion infectivitydissemination in soil.

Studies by Johnson et al. (2006) and Leita et al. (2006)were devoted to investigate the interaction of a scrapie-specific isoform of the prion protein or of a brainhomogenate from hamster-adapted scrapie 263K strainwith mineral soil constituents and whole soils, respectively.Moreover, studies were performed by Revault et al. (2005)and Rigou et al. (2006) on the adsorption/desorption of anovine recombinant PrP onto montmorillonite, as represen-tative of soil clays, and some sandy soils. In both studies,the authors suggested that the N-terminal part of the ovinerecombinant PrP rich in positive amino acid residues andconferring to the protein a net positive charge played amain role in the adsorption/desorption processes. Johnsonet al. (2006) also demonstrated that PrPSc stronglyinteracted with montmorillonite and cleaved at an N-terminal site upon desorption.

Little attention, if any, was, however, paid to the role ofsoil organic matter and organo-mineral complexes in theinteraction with prion protein. In soil, organo-mineralmatrices are expected to be more common than isolatedhumic or clay components. Organic matter is an importantconstituent of soils with properties markedly different frommineral constituents and contributes to the adsorption/entrapment of molecules and macromolecules entering thesoil. Polymerization of simple organic monomeric pre-cursors occurring in the presence of oxidative enzymes ormanganese and iron oxides is considered one of the mostimportant processes contributing to the formation ofhumic substances in soil (Bollag et al., 1995; Huang,1995). The process is very fast and produces a populationof polymeric products of different molecular structures,sizes, shapes and complexity (Huang, 1995). Furtherpolymerization, self-assemblage of different polymericproducts, and/or association with mineral constituentsquickly lead to the formation of more complex organic and

organo-mineral soil colloids (Huang, 1995). If activeproteins are involved in the process, protein–organic andprotein–organo–mineral complexes with different structur-al and functional properties may form. Furthermore, theinteraction of proteins with the prevalently hydrophobicsurfaces of organic matter might cause structural con-formational changes in the immobilized protein. In the caseof the prion protein, such conformational changes mayresult in the transformation from the non-pathogenic to thepathogenic form of the protein. To our knowledge, noinformation is available on the interaction of prionproteins with organic and organo-mineral soil components.In this study we have investigated the interaction of a

recombinant ovine protein (Rezaei et al., 2000), structu-rally well characterized (Eghiaian et al., 2004), with organicand organo-mineral components obtained by birnessite-mediated polymerization of catechol and resemblingorganic and organo-mineral soil colloids. The recombinantprotein is used as a model of the natural cellular prionprotein PrPC (Leclerc et al., 2001; Somerville, 2002;Legname et al., 2004). Catechol is a very common humusprecursor, is frequently present in soil, and may beinvolved in polymerization processes by biotic and abioticcatalysis (Huang, 1995; Naidja et al., 1998; Matocha et al.,2001). Birnessite (d-MnO2) is a naturally occurring, poorlycrystalline manganese oxide, abundantly present in soil,and promotes the oxidative transformation of phenols(Bollag et al., 1995; Huang, 1995; Rao et al., 1999).The properties of the deriving complexes, the allocation

and the stability of the protein in/on the complexes as wellas its potential release from the formed complexes and itspossible structural alteration were investigated. In order toevaluate the role of the N-terminal part of the protein inthe interaction with catechol and birnessite, comparativestudies were also performed with the C-terminal part of theprotein, i.e. missing the N-terminal fragment.

2. Material and methods

2.1. Chemicals

Reagent-grade catechol (Cat) (499.0% purity) andhigh-performance liquid chromatography (HPLC)-gradesolvents were purchased from Sigma Aldrich (Germany).Sarkosyl (N-lauroyl sarkosyl sodium salt) was from

Fluka (Germany). All other chemicals, reagent grade, weresupplied by Analar, BDH Ltd. (Germany), unless other-wise stated.Birnessite (d-MnO2) (Bir) was synthesized with KMnO4

and HCl according to McKenzie (1989). X-ray diffractionstudies demonstrated the presence of poorly crystallineminerals with characteristic peaks as reported in theliterature (McKenzie, 1989).A purified full-length ovine ARQ genetic variant (MM

23kDa, residues 23–234) prion protein (PrP) and its C-terminal fragment (MM 15kDa, residues 103–234), there-after indicated as tPrP, were prepared according to Rezaei

ARTICLE IN PRESSM.A. Rao et al. / Soil Biology & Biochemistry 39 (2007) 493–504 495

et al. (2000) and Eghiaian et al. (2004), respectively. Bothwere kindly furnished by Dr. Jeanne Grosclaude, from theVirologie et Immunologie Moleculaires, INRA (Jouy-en-Josas, France). The full-length protein is formed by a well-folded C-terminal domain (residues 125–234) and a ratherflexible N-terminal arm (residues 23–124) (Eghiaian et al.,2004). It has an isoelectric point (pI) of 9.2, a molarextinction coefficient of 57,930M�1 cm�1 at 280 nm, and itsstructural stability is preserved in the pH range from 4.6 to7.2 (Rezaei et al., 2000). Above and below these pH values,the protein may loose partly its a-helix conformation toachieve a b sheet character (Rezaei et al., 2000). The C-terminal fragment used in this study has a theoretical pI of8.84 and a molar extinction coefficient of 18, 005M�1 cm�1

at 280 nm (Eghiaian et al., 2004).

2.2. Circular dicroism (CD) spectroscopy measurements

The behavior of PrP in all buffers and components waschecked by CD measurements and only those whichpreserved PrP stability and solubility were successivelyused.

CD measurements were carried out in the far-UV (250–190 nm) using a Jasco J-810 spectropolarimeter equipped,with a Peltier type temperature control system (ModelPTC-423-S). CD spectra were recorded with a 0.1 cmoptical path length of the cell, a time constant of 4 s, a 2 nmbandwidth, and a scan rate of 5 nmmin�1. All spectra weresignal-averaged over at least three scans and the baselinecorrected by subtracting the buffer spectrum. Concentra-tion of PrP used for CD experiments was in the range 0.2–0.5mgml�1.

CD spectra were recorded on PrP–MnCl2 mixtures usinga 0.25mgml�1 PrP concentration, 10mM acetate buffer,pH 5.5 and containing MnCl2 amounts corresponding toMn2+:PrP molar ratios equal to 10, 40, 100, 1000, very farexceeding those possibly occurring in the polymerizationconditions. The solutions were incubated for 18 h at 20 1Cprior CD measurements.

Thermal unfolding curves were measured in the tem-perature mode at 222 nm in the range 20–80 1C using ascanning rate of 1 1Cmin�1. To control the reversibility ofthe unfolding process, PrP refolding was followed using thesame scanning rate, and the conservativeness of the meltingtemperature was checked.

In order to identify experimental conditions suitable forextraction experiments, the behavior of PrP in solutions ofCu2+ was studied by recording CD spectra at variousincubation times (10min, 30min, 18 h, 30 h) in 10mMphosphate buffer at pH 7.5. This pH was chosen to keepHis residues of the protein N-terminal arm, devoted toCu2+ binding, in their uncharged state. Solutions wereprepared using a PrP:CuCl2 ratio of 1:4, 1:10, 1:30, and1:100. Best conditions, which did not produce proteinprecipitation, were used for the extraction experiments (seeabove). Similarly, the stability of PrP in pyrophosphate

solutions was measured by recording CD spectra afterincubation times of 10 and 30min, 2 and 5 h.

2.3. Formation of catechol–birnessite–protein complexes

Catechol and birnessite mixtures were prepared byincubating 1ml of 0.1M sodium acetate buffer, pH 5.5,increasing amounts of birnessite (1–5mgml�1) and in-creasing concentrations of catechol (1–5mM) in Erlen-meyer tubes for 1, 2 and 24 h at 25 1C. After incubation, themixtures were centrifuged for 30min at 10,000 g and at4 1C, and residual catechol and soluble manganeseconcentrations in the supernatants were determined. FinalpH of the mixtures was measured as well. Appropriatecontrols with either acetate buffer, Cat or Bir only werealso carried out.The interaction of either PrP or tPrP with catechol and

birnessite was investigated using 3 and 5mM catechol,0.5mgml�1 protein (by adding suitable amounts of afreshly prepared protein solution whose concentration wasdetermined by absorbance at 280 nm) and 5mgml�1

birnessite in 1ml of 0.1M sodium acetate buffer, pH 5.5.Catechol, birnessite and PrP or tPrP were incubated indifferent trials. Two ternary systems catechol–birnessite–protein were produced: Cat–Bir–PrP or Cat–Bir–tPrPwhere the three components were mixed at the same time,and Cat–Bir+PrP or Cat–Bir+tPrP where Cat and Birwere incubated for 2 h before protein addition. Threebinary systems were also produced: Cat–Bir, Cat–PrP orCat–tPrP and Bir–PrP or Bir–tPrP. Samples with only Cat,PrP, tPrP or Bir served as controls.All mixtures were incubated for a total of 4 h at 25 1C. At

the end of incubation samples were centrifuged (30min at10,000 g and 4 1C) and the supernatants were analyzed forresidual catechol, soluble Mn concentrations, final pH andUV–Vis spectra. The presence of PrP or tPrP in thesupernatants was revealed by SDS-PAGE electrophoresisand their amounts were measured by HPLC analysis.Indeed, the presence of catechol and its polymericproducts, absorbing at 280 nm, prevented the determina-tion of protein concentration from direct UV measure-ments. Occasionally, a colorimetric method, the ProteinAssay ESL (Roche, Germany) was used (detection limit20 mg). No interference under all the experimental condi-tions was observed, thus allowing the concentration of PrPor tPrP to be determined in the presence of catechol orcatechol polymers.The precipitates were washed twice with 0.02M NaCl

and then with bi-distilled water until acetate free,lyophilized and stored at 4 1C for further analyses asreported below.Experiments of catechol polymerization were also

performed by adding catechol to birnessite that hadpreviously reacted with PrP. In 1ml of 0.1M sodiumacetate, pH 5.5, 5mg of birnessite and 0.5mg of PrP wereleft to interact for 2 h. Two test tubes were supplementedwith 0.1ml of 0.05M catechol and left to interact for

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another 2 h. Controls without catechol were produced todetermine the amount of PrP in the supernatants. At theend of incubation, the mixtures were processed aspreviously described.

2.4. Experiments with soluble catechol polymers

Experiments were performed adding 0.5mgml�1 (finalconcentration) PrP or tPrP to soluble catechol polymersobtained in the supernatant of the two binary systems Cat–Bir (3 and 5mM catechol). After 2 h incubation time at25 1C some aliquots were analyzed by UV–Vis and CDspectroscopy, and the remaining parts were centrifuged for30min at 10,000 g and at 4 1C. After removal ofprecipitates the amounts of PrP or tPrP were quantifiedby HPLC and electrophoresis analyses.

2.5. Desorption/extraction tests

Sequential extraction tests were carried out by suspend-ing suitable amounts (usually 5mg) of the solid phasesobtained from centrifugation of the ternary mixtures withPrP or tPrP in 1ml of 0.1M phosphate buffer at pHincreasing from 7.0 to 8.5. The buffered suspensions werekept under mixing at 25 1C for 1 h. Further extractionexperiments were performed at 25 1C using for 1 h in ashaker either 0.14M pyrophosphate at pH 7.0 or 0.7mMCuCl2 solution in 0.1M phosphate buffer, pH 7.5.Sequential extractions using a water solution containing10% (v/v) propanol and incubation under magnet stirringfor 24 h and further 72 h as well as treatments for 1 h withan extraction solution composed by 1% sarkosyl in 0.1Mphosphate buffer, pH 7.5 were also carried out.

After the treatment, all the samples were centrifuged for30min at 10,000 g and 4 1C and the presence of PrP or tPrPreleased in the supernatants was determined by proteindetermination assays, HPLC and electrophoresis analyses.

When measurable amounts of PrP or tPrP were detectedin the extracts, CD measurements were performed toevaluate, if any, the variation of the protein conforma-tional structures.

2.6. HPLC analyses

Catechol, PrP and tPrP concentrations were determinedby HPLC analysis with a Shimadzu instrument equippedwith a variable-wavelength absorbance detector set at 280,222 and 227 nm, respectively. A Spheri-5-RP1822 cm� 4.6mm C18-80 BrownLee (Chebios) column witha 5 mm particle size and a BrownLee Spheri-5-RP300 7 mmparticle size (4.6mm� 30mm) guard column for catechol,a Biosep Sec-S2000 300mm� 7.80mm Phenomenex col-umn and Biosep Sec-S2000 75mm� 7.80mm Phenomenexguard column for both the proteins were used. Catecholanalysis was performed in isocratic elution at a flow rate of1mlmin�1 with a mobile phase composed of acidifiedwater (3ml of 95% H3PO4 in 1 l of water, at pH 2.95 with

0.5M NaOH, low in carbonate) and acetonitrile at a ratioof 70:30 (v:v). Full-length PrP and tPrP were eluted at aflow rate of 1mlmin�1, with 6M guanidine hydrochlorideand 20mM phosphate buffer, pH 7.5, respectively. Beforeinjection, all samples were filtered with Acrodisck LC 13PVDF 0.45 mm filters. Retention time for catechol, PrP andtPrP were 3.5, 6.5, and 9.8min, respectively.

2.7. Atomic absorption spectroscopic and elemental analyses

Atomic absorption spectroscopic measurements of so-luble Mn were performed at 279.5 nm with a Perkin-ElmerAnalyst 700 by using flam adsorption spectroscopy. Beforeeach investigation standard solutions (0.5, 1.0 and2.0mg l�1) were freshly prepared by diluting theMnCl2 � 4H2O standard solution in HCl. The detectionlimit for Mn2+ determination was 0.0082mg l�1.Carbon and nitrogen contents of lyophilized insoluble

products (see above) were measured by the ash combustionprocedure with a Fisons 1108 elemental analyzer. Calibra-tion of the Fisons instrument with appropriate standard(acetanilide) was carried out. Accuracy (o0.05%) andrecovery of C and N (for both instrument detection limit10mg kg�1) were checked analyzing a sample of thestandard material after each set of eight sample analyses.

2.8. UV–Visible and FT-IR analyses

The UV–Visible spectra of products remaining in thesupernatants of the catechol–birnessite and catechol–birnessite–PrP or –tPrP mixtures were obtained withscanning from 900 to 200 nm using a Perkin Elmerspectrophotometer Lambda 25 instrument.The Fourier transform infrared spectra of insoluble

products were recorded by the Universal Attenuated TotalReflectance (UATR) method using a Perkin Elmer FT-IRspectrometer. Each spectrum represents a collection of 16scans recorded at a 4 cm�1 resolution. Before FT-IRspectrum analyses, the insoluble samples were washedtwice with NaCl 0.02M to avoid acetate buffer signals.

2.9. Electrophoresis analysis

Denaturing SDS-PAGE electrophoresis with 15% or agradient of 10–20% (w/v) acrylamide was performed using aBio-Rad apparatus and the Laemmli discontinuous system.Gels were stained with Coomassie brilliant blue R250.In all cases, at least three independent experiments were

carried out and the standard deviation was lower than 5%.

3. Results

3.1. Catechol–birnessite–protein interactions

3.1.1. Preliminary studies by CD spectroscopy

Preliminary experiments were carried out to establishproper experimental conditions, which did not affect

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protein stability and/or did not produce aggregationeffects. A particular care was dedicated to the analysis ofPrP behavior in the presence of Mn2+, which is producedin detectable amounts in the polymerization reaction.Indeed, PrP is known to have a strong tendency to formaggregates in the presence of metal ions such as Cu2+ andMn2+ (Giese et al., 2004). Therefore, this preparative taskwas aimed at avoiding PrP aggregation under the experi-mental conditions used.

No aggregation was induced by Mn2+ at any of theMn2+:PrP ratios, as described in ‘‘Materials and methods’’section. Indeed, CD spectra of Mn2+:PrP solutions after18 h incubation were completely superposable to those ofPrP obtained in the absence of Mn2+ (data not shown).Furthermore, the presence of Mn2+ did not affect PrPstability, since its melting temperature, as measured bythermal denaturation, remained unaltered (Tm ¼ 79 at pH5.5). It is worth noting that thermal denaturation leads,under these experimental conditions, to a random coil stateand not a b structure. Indeed the CD spectrum recorded at80 1C (Fig. 1) shows a clear minimum close to 200 nm, butnot at 214 nm (Rath et al., 2005). Notably, also in thepresence of Mn2+ the denaturation process was fullyreversible (Fig. 1).

Further experiments were performed to evaluate possibleeffects by the chemicals used in the extraction/desorptionstudies reported below. While no effects on PrP stabilitywere observed with phosphate or propanol, a clearinfluence by pyrophosphate on PrP aggregation was noted.Indeed, CD spectra performed on PrP before and afterincubation for 18 and 24 h with pyrophosphate showedthat it induced protein aggregation, as the minimum at208 nm (typical of a-helix) became less deep after 18 and24 h incubation time (data not shown). At this time a slightopalescence was also observed and, consistently, the CDsignal decreased. This roughly corresponded to a proteinloss of 20% (w/v). Incidentally, interference on CDanalysis was observed when catechol polymers alone(supernatants of Cat–Bir mixtures) were mixed with

190 200 210 220 230 240 250

-10

-5

0

5

10

15

[Θ]

(deg

cm

2 d

mo

l-1)

10-3

Wavelength (nm)

20°C

40°C

60°C

70°C

80°C

20°C after refolding

aa

bb

cc

d

d e

e

f

f

Fig. 1. CD spectra of PrP in the presence of Mn2+ at different

temperatures. Experiments were performed in acetate buffer, pH 5.5,

and with a Mn2+:PrP ratio of 100. Spectra were recorded in the range 20–

80 1C and at 20 1C after refolding.

pyrophosphate, probably because of the release of poly-meric materials by pyrophosphate (data not shown). Nointerference occurred with pyrophosphate or Cat–Biralone.

3.1.2. Formation of catechol–birnessite–protein complexes

Preliminary experiments were performed with onlycatechol and birnessite to select the most appropriateconditions to utilize in the formation of catechol–birnes-site–protein complexes. These conditions should simulatetwo possible situations encountered by a protein in soil:protein molecules interacting with soluble organic matterin the first phases of its formation or with insoluble organicmatter which already underwent partial or strong con-densation.Investigations performed by increasing catechol concen-

trations from 1 to 5mM and birnessite amounts from 0.2to 5mgml�1 under buffered conditions (0.1M Na acetate,pH 5.5) showed that after 2 h incubation catechol removalincreased from 20% (at 1mM catechol and 0.2mgml�1

birnessite) up to 100% at the highest catechol andbirnessite amounts. A corresponding release of solubleMn2+ occurred; after 2 h incubation of 5mM catechol and5mgml�1 birnessite, the concentration of Mn2+ raised upto 0.131mgml�1. Neither catechol removal nor Mn2+

release occurred in the controls lacking birnessite orcatechol, respectively. Moreover, under buffered condi-tions at pH 5.5, the pH of the mixtures was stable (amaximum increase of 0.3 pH units was detected) and theprotein was not subjected to unfolding phenomena at roomtemperature.Soluble polymers were obtained at 3mM catechol,

whereas at 5mM catechol insoluble polymers prevailed,as demonstrated by several experimental evidences. Indeed,the absorbance in the visible region 900–350 nm of thesupernatant of 3mM catechol–birnessite mixtures wasmuch higher than supernatants of 5mM catechol mixtures.For instance, at 400 nm the absorbance value of 3mMcatechol supernatants was 1.15 against 0.65 measured atthe same wavelength for 5mM catechol samples, whereinsoluble materials simultaneously precipitated at thebottom of the incubation tubes, with a corresponding,visible at naked eye clarification of the supernatants.The formation of insoluble catechol polymers at 5mM

catechol was supported by both CHNS and FT-IRanalysis. Although the oxidative coupling reaction withcatechol caused the dissolution of 0.2mg birnessite(corresponding to 0.116mg of Mn2+ determined insolution by AAS), the weight of the precipitate did notchange when compared to the initial MnO2 amount, sincenew insoluble organic materials enriched it. Detectableamounts of carbon (C, deriving exclusively from catechol,because no C can be released by birnessite, and the solidsamples were acetate-free) were measured by elementalanalysis, thus confirming the formation of insolublecatechol polymers adsorbed on the surfaces of birnessite.For instance, 0.131mg of C corresponding to about 36%

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of the initial catechol quantity was found in the solid phaseof Cat5–Bir samples.

FT-IR spectra of precipitates also supported the aboveresults (data not shown). As reported by Russo et al.(2005), the higher the catechol/birnessite ratio (reducent/oxidant ratio), the lower the intensity of the peak at512 cm�1, characteristic of birnessite, and the higher theintensities in the range 1620–1000 cm�1, thus confirmingthe corresponding consumption of birnessite and theappearance of more complex products.

On the basis of these results, the interaction of PrP ortPrP with birnessite and catechol was studied in 0.1Macetate buffer at pH 5.5, using 5mgml�1 Bir, 0.5mgml�1

PrP or tPrP and 3 and 5mM catechol in four independentexperiments as described in detail in ‘‘Materials andmethods’’ section.

Full-length PrP or tPrP molecules neither affected theextent of catechol removal (100% in all samples whetherpresent or not the two proteins) nor the correspondingrelease of soluble Mn2+ (in larger amounts with Cat5)(Table 1). A simultaneous, complete removal of theproteins from the mixture at the two catechol concentra-tions was measured both when they were incubated at thebeginning with catechol and when added after catecholpolymerization (Table 1). Neither removal of catechol andPrP or tPrP, nor release of soluble Mn2+ occurred in therelative controls Cat, PrP, tPrP or Bir. Moreover, whenPrP or tPrP interacted with only birnessite in the absence ofcatechol (Bir–PrP and Bir–tPrP samples), 65% and 55% ofproteins were removed from the solution and adsorbed onbirnessite surfaces, respectively (Table 1). All of theseresults were supported by electrophoresis and HPLCanalyses performed on the relative supernatants of allsamples. Neither protein bands nor protein amounts weredetected in any of the supernatants, except for those of Bir–PrP and Bir–tPrP samples.

Notably, catechol polymerization still occurred withbirnessite loading PrP molecules previously adsorbed on its

Table 1

Percentage of protein removal and concentration of Mn2+ released during

the complete catechol polymerization with SD values

Samples Protein removal (%) Mn2+ (mgml�1)

PrP tPrP PrP tPrP

Cat3–Bir–protein 100a 100a 0.06470.003 0.06370.002

Cat5–Bir–protein 100a 100a 0.13270.005 0.13070.004

Cat3–Bir+protein 100a 100a 0.06970.003 0.06770.003

Cat5–Bir+protein 100a 100a 0.12670.006 0.12870.005

Cat3–Bir ND ND 0.06170.003 0.06170.003

Cat5–Bir ND ND 0.11670.005 0.11670.005

Bir–protein 6572 5571 0.00370.001 0.00370.001

Protein 070 070 ND ND

Bir–protein+Cat5 6572 ND 0.13270.005 ND

Cat3 ¼ 3mM catechol; Cat5 ¼ 5mM catechol; Bir ¼ birnessite.

ND ¼ not determined.aSD are below the sensitivity of the adopted method.

surfaces. When 5mM catechol was added to birnessitecoated by PrP (65% of adsorption), the complete catecholremoval was observed (Table 1), as when protein-freebirnessite was used.

3.1.3. Features of catechol–birnessite–protein complexes

The presence of PrP strongly affected the patterns andthe absorbance values of the supernatants of both 3 and5mM catechol mixtures and the formation of insolublecatechol–birnessite–PrP aggregates. The absorbance ofsupernatants with 3mM catechol in the visible region wasstill higher than that of 5mM catechol samples, thusconfirming the presence of a greater amount of solublepolymers at lower catechol concentration. When PrP wasadded after catechol polymerization (Cat3–Bir+PrP, andCat5–Bir+PrP samples), the absorbance of the super-natants markedly diminished in the whole UV–Visibleinterval, with a decrease much more conspicuous in thevisible region (Fig. 2A), and an evident and rapidformation of brown insoluble material, visible at nakedeye, accumulated spontaneously at the bottom of theincubation flasks. Similar UV–Vis spectra were obtainedwith tPrP (data not shown).The presence of PrP in the precipitates with birnessite

was also confirmed by the elemental analyses. Indeed, anincrease of C content and the appearance of detectableamounts of N were measured, compared to the corre-sponding samples without protein. For instance, for thesamples obtained with 5mM catechol, the C contentincreased from 0.131 up to 0.398 and 0.060mg of N weredetected. While the measured values of C did not allow toreveal the presence of the protein in the solid phase, thedetection of N (deriving exclusively from proteic material)demonstrated that PrP was present in it.FT-IR measurements of precipitates were performed.

They supported the hypothesis that the majority of thereaction products of catechol polymerization, with andwithout PrP, were apparently adsorbed on birnessitesurfaces, and thus could not be detected in the super-natants. Fig. 3 shows the FT-IR spectra of Cat5–Bir, Cat5–Bir–PrP and Cat5–Bir+PrP solid aggregates. The twosignals at 1622 cm�1 (aromatic CQC, H-bonded CQO oralkenes in conjugation with CQO) and 1254 cm�1 (C–Ostretching and aromatic CQC) in the Cat5–Bir spectrumare characteristic spectroscopic features of humic-likearomatic compounds. As respect to the Cat5–Bir sample,any modification observed in both Cat5–Bir–PrP andCat5–Bir+PrP spectra must account for the sharp amideI (CQO stretching at 1644 cm�1)and amide II (N–H bending at 1520 cm�1) bands of theprotein. Moreover, the reinforcement of the band at1254 cm�1 (weak in the Cat5–Bir sample) could beattributed to the presence in the same region of thebending (amide III) signal of the protein. Similar indica-tions were provided by the spectra of the catechol–birnessite–PrP complexes obtained at 3mM catechol (datanot shown).

ARTICLE IN PRESS

300 400 500 600 700 800 9000.0

0.2

0.4

0.6

(B)

(A)

Abs

orba

nce

300 400 500 600 700 800 9000.0

2.0

3.0

4.0

5.0

1.0

300 400 500 600 700 800 9000.0

0.1

0.2

0.3

0.4

(C)

Abs

orba

nce

Wavelength (nm)

Cat3-Bir-PrPCat3-Bir+PrPCat5-Bir-PrPCat5-Bir+PrPCat3-BirCat5-Bir

Cat3-BirCat3-Bir+PrPCat3-Bir+tPrP

Abs

orba

nce

Cat5-BirCat5-Bir+PrPCat5-Bir+tPrP

aa b

b

cc d

d

e

e

ff

aa b

b

c

c

aa b

b

c

c

Fig. 2. UV–Visible spectra of (A) supernatants of catechol–birnessite–PrP

mixtures after 4 h incubation. Spectra of Cat3–Bir and Cat5–Bir are also

shown (see insets); (B) supernatants of Cat3–Bir mixtures and (C)

supernatants of Cat5–Bir mixtures after 2 h incubation with 0.5mgml�1

PrP and tPrP. The spectra of Cat3–Bir and Cat5–Bir supernatants

obtained after a 2 h incubation of catechol and birnessite, and prior to

protein addition, are also reported as a reference.

Tra

nsm

ittan

ce

4000 3500 3000 2500 2000 1500 1000

Wavenumber (cm-1)

1644,19

1539,88

1254,21

1621,56

Cat5-Bir

Cat5-Bir+PrP

Cat5-Bir-PrP

Fig. 3. Fourier-transform infrared (FTIR) spectra of Cat5–Bir, Cat5–

Bir–PrP and Cat5–Bir+PrP solid aggregates.

M.A. Rao et al. / Soil Biology & Biochemistry 39 (2007) 493–504 499

3.2. Interaction of PrP and tPrP with catechol polymers in

the absence of birnessite

In order to distinguish the contribution of the solublecatechol polymers (i.e., in the absence of the mineral partloading or not organic material on its surfaces) to theinteraction with either PrP or tPrP, the proteins were addedto the supernatants of Cat3–Bir and Cat5–Bir mixturesafter 2 h incubation. Protein–catechol insoluble productswere formed. A rapid, immediate flocculation of insoluble,brown material possibly including the protein occurredwith a concurrent clarification of the solution. Thisdecoloration was confirmed by a progressive decrease ofthe UV–Visible signal in the wavelength range 900–200 nm(Fig. 2B and C). CD spectra, recorded immediately afterthe solution preparation and followed for a few hours, alsoshowed an almost immediate and progressive decrease ofthe CD signal in the whole wavelength range (Fig. 4). Inthe case of PrP, a removal of 92% and 74% was detectedafter 2 h contact time with Cat3–Bir and Cat5–Bir super-natants, respectively, by HPLC and confirmed by SDS-PAGE electrophoresis (Fig. 5). A significantly reducedremoval was observed for tPrP by HPLC (15% in Cat5–Birand 33% in Cat3–Bir), consistent with CD spectra (Fig. 4)and SDS-PAGE electrophoresis (data not shown). A lessintense flocculation of insoluble material was also ob-served. After 8 h incubation, electrophoresis and HPLCanalyses on aliquots of the sample supernatants showedthat both the protein forms completely disappeared fromthe reaction mixtures, thereby indicating an immediate andprogressive interaction of the proteins with soluble catecholpolymers to form insoluble aggregates which accumulatedin the precipitate.The greater protein removal in Cat3–Bir supernatants is

likely due to the higher amounts of soluble polymers inthese samples, which can interact with protein molecules(Fig. 2A).These results indicate that (i) in the presence of soluble

catechol polymers, the removal of both PrP and tPrP issignificantly slower than in the corresponding heteroge-

200 210 220 230 240 250

-15

-10

-5

0

[Θ] (

deg

cm2

dmol

-1)

10-3

Wavelength (nm)

Cat3-Bir+tPrP

Cat5-Bir+tPrP

tPrP

Fig. 4. CD spectra of supernatants of Cat3–Bir and Cat5–Bir mixtures

after 2 h incubation with 0.5mgml�1 tPrP. The spectrum of tPrP in 10mM

acetate buffer, pH 5.5, is also reported as a reference. Only samples

containing tPrP are shown, for clarity.

ARTICLE IN PRESS

Fig. 5. SDS-PAGE electrophoresis of supernatants of Cat3–Bir and

Cat5–Bir mixtures after the addition of 0.5mgml�1 PrP.

M.A. Rao et al. / Soil Biology & Biochemistry 39 (2007) 493–504500

neous systems including birnessite, and (ii) the lack of theN-terminal fragment (residues 23–93) in tPrP significantlyslows down the process of protein–catechol polymerinteraction. This is consistent with the presence in theprotein N-terminus of positively charged residues, whichare potential interacting moieties.

3.3. Extraction/desorption tests

Different experimental conditions were tested to desorband/or to extract the PrP or tPrP from the insolublecomplexes obtained.

The pellets obtained after centrifugation of each samplewere subjected to sequential extraction tests with 0.1Mphosphate at pH 7.0, 8.0 and 8.5. No detectable removal ofPrP or tPrP from all the samples was observed after 1 h ofcontact, as assessed by HPLC-protein determination andSDS-PAGE electrophoresis. A positive response to elec-trophoresis analysis was observed only with the Bir–PrPsample.

With 0.14M pyrophosphate at pH 7.0, the Protein AssayESL identified detectable amounts of PrP (30%, 18% and15% for Cat3–Bir–PrP, Cat3–Bir+PrP and Cat5–Bir-PrP,respectively). However, these results were not confirmed byelectrophoresis analyses (no detection of protein bands)indicating a false-positive response likely due to thecapability of pyrophosphate to release polymeric materialfrom the formed catechol–protein complexes. Indeed, aspreviously reported a detectable amount of protein (20%)along with polymeric material was actually released in thepresence of pyrophosphate. Pyrophosphate was proven tobe very effective in the extraction of proteins from soil and/or soil organic complexes (Tabatabai and Fu, 1992;Nannipieri et al., 1996).

PrP extraction tests were also performed using CuCl2 orpropanol solution by incubating for 1 h in a shaker or forlonger time (72 h), with a magnet stirring procedure. Cuions show a strong affinity for the prion protein

(Brimacombe et al., 1999; Brown et al., 2000; Brown,2001; Giese et al., 2004) and propanol is capable ofinterfering with the hydrophobic interactions possibleestablished between the protein and the organic complexes(Wershaw, 2004). In both experimental conditions, witheither the copper or propanol solutions, the protein wasnot detected in the extracts.Finally, sarkosyl, a strong anionic detergent, usually

utilized for the cleaning of glass and instruments inlaboratories, was tested. Release of PrP or tPrP wasrevealed neither by HPLC nor by electrophoresis analyses,even from Bir–PrP or Bir–tPrP samples.

4. Discussion

The results of the present work highlight the strong andirreversible interactions of recombinant PrP with organicand organo-mineral compounds, thereby emphasizing theimportance that organic or organo-mineral soil colloidsmay play in the fate of prion proteins in soil. For thesestudies, soil colloid models were derived by birnessite-mediated polymerization of catechol. Although the com-plexity of natural soil systems was not reproduced, thesemodel systems can simulate some natural processesoccurring in soil.The experiments were performed under buffered condi-

tions at pH 5.5 to favor the formation of insoluble oversoluble phenol polymers (Naidja et al., 1998) and topreserve the a-helix conformation of the recombinant PrP(Rezaei et al., 2000).A complete removal of both PrP and tPrP from the

solution was observed under all experimental conditionsinvestigated. Strong interactions of PrP occurred with theoxidative polymerization products whichever soluble poly-mers or insoluble aggregates (Table 1). These conclusionswere drawn from several experimental evidences such asthe disappearance of protein molecules from the solution,the decrease of UV–Visible absorbance of supernatants,the FT-IR spectra and elemental analyses of solid-phaseresidues, and the nil amounts of PrP or tPrP detected byHPLC and SDS-PAGE electrophoresis.Moreover, the interaction of PrP or tPrP with birnessite

and birnessite-mediated catechol polymers was so strongthat both protein forms could not be released from theformed solid aggregates neither by (a) phosphate andpyrophosphate buffers, (b) organic solvents, (c) solutionsof copper, nor by a strong anionic detergent like sarkosyl.The oxidation of catechol by Mn(IV) oxides is a rapid

multistep, redox-surface process (Matocha et al., 2001),leading to the formation of soluble and highly insolubleorganic products accumulating on birnessite surfaces.Given some constant experimental conditions (e.g., in-cubation temperature and time, pH, amount of birnessite),the transformation of catechol in either soluble or insolublepolymers depends on the initial catechol concentra-tion. Our results have clearly indicated that solublepolymers prevailed over insoluble ones using a catechol

ARTICLE IN PRESSM.A. Rao et al. / Soil Biology & Biochemistry 39 (2007) 493–504 501

concentration of 3mM whereas an opposite situationoccurred at 5mM (Fig. 2).

When the protein was also present in the reactionmixture along with catechol and birnessite, a rathercomplicated interplay of physically mediated processesand chemical reactions probably occurred, thereby leadingto a very complex sequence of pathways and productsbeing formed (Fig. 6). The investigated systems differedsubstantially from each other whether the protein wasmixed with catechol and birnessite at the same time (Cat–Bir–PrP) (Fig. 6A), after the preventive incubation betweencatechol and birnessite (Cat–Bir+PrP) (Fig. 6B), or withcatechol polymers in the absence of birnessite(Fig. 6C). To attempt an explanation of the obtainedresults, it is necessary to separately analyze the contribu-tion of the different phenomena to the whole process.Namely, the proteins interact with (i) birnessite alone, (ii)catechol polymers alone, (iii) birnessite partly covered bycatechol polymers, and (iv) birnessite and catechol.

The most complex situation holds when the proteins,catechol and birnessite were mixed at the beginning of theincubation (Fig. 6A). In this case, along with the formationof soluble catechol polymers, the involvement of NH andOH groups of the proteins in cross-coupling reactions withphenoxy radicals produced by birnessite action is likely tooccur (Fig. 6A). Several findings have demonstrated thatamino acids, humic substances and phenols may give riseto cross-coupling products during the oxidative polymer-ization of phenols promoted by birnessite (Bollag et al.,1995; Huang, 1995). In this process, the higher number ofNH groups in PrP may have favored the entrapment of theprotein in catechol–birnessite insoluble precipitates. Thisphenomenon is less probable when the protein was addedafter 2 h incubation where free catechol was completelytransformed (Cat–Bir+PrP and soluble catechol poly-mer+PrP samples; Fig. 6B and C).

+

+

Cross-coupling of Cat wi

Formation of soluble Cat

Adsorption of PrP on Bir

Adsorption of Cat polyme

Cat

PrP

Bir

(A)

Adsorption of Cat polymers on Bir

Formation of soluble Cat polymers

(B)

(C)

Cat

Bir

Fig. 6. Scheme of possible physical and chemical pr

In this reactive context, we also evidenced the adsorptionof either PrP or tPrP on birnessite surfaces (Fig. 6A), sincehigh percentages of the two proteins were removed fromthe solution in the presence of the sole birnessite. Thisfinding was consistent with the presence of amide I andamide II bands in the FT-IR spectra of Bir–PrP insolublecomplex (data not shown). At the adopted experimentalpH (5.5), birnessite surfaces [point of zero charge (PCZ)1.81, according to Matocha et al. (2001)] and PrP ortPrP are negatively and positively charged, respectively.Therefore, strong electrostatic interactions may be estab-lished, as already observed in the literature (e.g., tyrosinasewas adsorbed on birnessite surfaces up to 89% of itsinitial amounts, Naidja et al. (2002)). In addition, aninteraction of PrP with birnessite sites coated with catecholpolymers is also likely to occur (Fig. 6B), in particular forCat3–Bir+PrP and Cat5–Bir+PrP samples (where PrP isadded after a preventive incubation of catechol andbirnessite). In this case (Fig. 6A and B) it is possible tohypothesize that the two phenomena, i.e. adsorption of thePrP on birnessite surfaces and its interaction with phenolpolymers adsorbed on these latter, may simultaneouslyoccur.Besides interacting with birnessite–catechol complexes

and with birnessite alone, proteins may also interact withsoluble catechol polymers, a situation which we simulatedby incubating either PrP or tPrP with the supernatants ofCat5–Bir and Cat3–Bir samples (Fig. 6C). Protein interac-tions with soluble phenolic polymers during or after theirformation involve several phenomena like (i) multidentatebinding of phenolic polymers to the protein, a processwhich can become irreversible with time and form insolublephases (Ladd and Butler, 1975); (ii) protein–catecholpolymers association through coulombic and/or hydro-phobic interactions; (iii) protein–phenolic polymeric com-plexes may transform from flexible soluble structures into

PrPFormation of Cat-PrP soluble andinsoluble aggregates

PrP

Adsorption on birnessite sites eithercoated or not by Cat polymers

th PrP

polymers

rs on Bir

Formation of Cat-Bir-PrPinsoluble complexes

Formation of Cat-PrP insolubleaggregates

ocesses occurring in the model systems studied.

ARTICLE IN PRESSM.A. Rao et al. / Soil Biology & Biochemistry 39 (2007) 493–504502

insoluble, reticulated structures and microporous micellesin the presence of positive charges (Gianfreda et al., 2002);(iv) quinoid compounds may covalently interact with theprotein (Bittner, 2006; Suderman et al., 2006).

Fig. 7. (A) Ribbon representation of ovine PrP structure. PrP C-terminal dom

strands (b1, b2). The flexible PrP N-terminal arm (gray) was modeled as con

sequence of the PrP form under investigation. (B) Electrostatic potential surfa

Multidentate binding is strictly influenced by thestructural features of both phenolic polymers and proteinand is assisted by conformational flexibility. In this respect,the long flexible N-terminal end in PrP is likely to play a

ain (PDB 1UW3, green) contains three a helices (H1, H2, H3) and two b—taining mainly polyproline II (Blanch et al., 2004). The inset shows the

ce of PrP as calculated using GRASP (Nicholls et al., 1992).

ARTICLE IN PRESSM.A. Rao et al. / Soil Biology & Biochemistry 39 (2007) 493–504 503

strong role in PrP interaction with catechol polymers alongwith charge interactions favored by the polycation-likebehavior of the PrP N-terminal region. This region con-tains as many as six positively charged residues (Fig. 7) thatare probable candidates for establishing favorable cou-lombic interactions with catechol polymers. Also, PrP N-terminus (residues 23–93) adopts mainly polyproline II andb conformations (Blanch et al., 2004). Polyproline II is asecondary structural motif with the peculiar feature oflacking any intra-helix backbone hydrogen bonds (Berisioet al., 2006), and as such involved in the molecularrecognition processes and/or in molecular interactions(Rath et al., 2005). All of its backbone hydrogen bonddonors/acceptors are non-saturated and are prone to formhydrogen bonds with neighboring molecules. Therefore,the abundance of positively charged residues in PrP as wellas the capability of PrP N-terminal arm to form hydrogenbonds concurs to account for the different behaviorobserved for PrP and tPrP in the interaction with solublecatechol polymers.

A role of the N-terminal arm of PrP was also suggestedby Revault et al. (2005) and Rigou et al. (2006) in theirstudies with montmorillonite. Electrostatic interactionsbetween the positive charges of the Arg and Lys residuesat the N-terminal of PrP and the negative surfaces ofmontmorillonite were hypothesized (Revault et al., 2005).A cleavage at N-terminal site due to a strong interaction inthis region was observed by Johnson et al. (2006) when theprotein was desorbed from montmorillonite with denatur-ing extracting agents.

The investigations performed on soils with differentphysico-chemical characteristics demonstrated that PrPentirely bound on soils, and desorption occurred only inconditions which may potentially denature protein mole-cules, and therefore very far from natural phenomena(Rigou et al., 2006; Johnson et al., 2006). Besides the well-known contribution of clay minerals, our findings indicatethat in the complex soil system humic materials may likelyplay an important role also in the interaction with PrPmolecules, thus exerting a stabilizing effect (Wershaw,2004).

In conclusion, overall the results reported here havecontributed to shedding light on the interaction of prionprotein with synthetic organic- and organo-mineral com-plexes simulating soil organic and organo-mineral colloids.Both in the protein only theory (Prusiner, 2004) and in themore general conception of ‘prion’, PrP is recognized as aninfectious protein particle. The irreversible interaction ofPrP demonstrated both with catechol polymers andbirnessite–catechol polymers mixtures, which simulatehumic substances and humic–mineral complexes, suggeststhat prion proteins should be strongly retained in soils richin organic matter, with very low risks of release andsubsequent dissemination in soil. Further investigationsare, however, mandatory to better clarify the strength andthe type of interactions between prion protein and soilorganic matter and to evaluate PrP behavior in real soil

samples representative of different natural situations.Additional studies should be also devoted to validate therole of the long and flexible N-terminal arm in the PrP-catechol polymer interactions, and mainly to evaluatewhether the immobilized and/or the released prion proteinstill retains its infectivity potential. Both kinds of investiga-tions are in progress.

Acknowledgments

This work was carried out with the support of Quality ofLife Program and partially of COFIN 03, Italy. Theauthors are grateful to Dr. J. Grosclaude and Dr. H.Rezaei, from the Virologie et Immunologie Moleculaires,INRA (Jouy-en-Josas, France) for a stimulating discussionon the achieved results and the supply of PrP and tPrPsamples.This paper represents Journal Series No. 115 from

DiSSPA.

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