Rock magnetic, chemical and bacterial community analysis of a modern soil from Nebraska

Earth and Planetary Science Letters 251 (2006) 168 – 178 www.elsevier.com/locate/epsl

Rock magnetic, chemical and bacterial community analysis of a modern soil from Nebraska Yohan Guyodo a,⁎, Timothy M. LaPara b , Amy J. Anschutz c , R. Lee Penn c , Subir K. Banerjee d , Christoph E. Geiss e , William Zanner f a

f

Laboratoire des Sciences du Climat et de l'Environnement, LSCE/IPSL (laboratoire CEA-CNRS-UVSQ), Campus du CNRS, 12 Avenue de la Terrasse, 91198 Gif-sur-Yvette Cedex, France b Department of Civil Engineering, University of Minnesota, 500 Pillsbury Drive SE, Minneapolis, MN 55455, USA c Department of Chemistry, University of Minnesota, 207 Pleasant Street S.E., Minneapolis, MN 55455, USA d Institute for Rock Magnetism, Department of Geology and Geophysics, University of Minnesota, 310 Pillsbury Drive SE, Minneapolis, MN 55455, USA e Trinity College, Department of Physics, 105 McCook, 300 Summit St., Hartford, CT 06106, USA Department of Soil, Water and Climate, University of Minnesota, 570 Borlaug Hall, 1991 Upper Buford Circle, St. Paul, MN 55108, USA Received 3 March 2006; received in revised form 25 July 2006; accepted 5 September 2006 Available online 9 October 2006 Editor: C.P. Jaupart

Abstract Detailed rock-magnetic and bacterial community analyses have been conducted on a 2.2 m-long soil profile developed over loess in Nebraska. Results show magnetic enhancement in the A-horizon of the soil profile, in part due to increased concentration of pedogenic superparamagnetic (SP) ferrimagnetic minerals (i.e., magnetite, maghemite). The magnetic enhancement correlates well with higher microbial biomass and lower iron hydroxide concentrations (here determined using a goethite-specific magnetic method), therefore supporting the idea of bacterially-mediated topsoil magnetite formation through iron-reduction of iron hydroxides. Cultivation-independent bacterial community analysis biased towards the Geobacteraceae (common dissimilatory iron-reducing bacteria) failed to show the involvement of this phylogenetic group in SP magnetite formation at this site. Although preliminary in nature, our failure to implicate dissimilatory Geobacteraceae with magnetic enhancement in modern Nebraskan soil and Dearing et al. (Dearing, Hannam, Anderson, Wellington, Geophysical Journal International 144 (2001) 183–196) earlier failure to detect magnetotactic bacteria in enhanced British soils points to the need to focus further multidisciplinary investigations on the possible roles of other dissimilatory iron reducing bacteria in contributing to magnetic enhancement. Many authors have produced strong circumstantial evidence of a microbial role in causing magnetic enhancement in topsoils of temperate climate zones, yet a precise role of specific cultured microbes in enhancing soil magnetism is still lacking. © 2006 Elsevier B.V. All rights reserved. Keywords: rock magnetism; mineral magnetism; iron reducing bacteria

⁎ Corresponding author. Tel.: +33 1 69 82 35 62; fax: +33 1 69 82 35 68. E-mail address: [email protected] (Y. Guyodo). 0012-821X/$ - see front matter © 2006 Elsevier B.V. All rights reserved. doi:10.1016/j.epsl.2006.09.005

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1. Introduction One of the major successes in the young field of environmental magnetism [1,2] has been the discovery that the central Chinese loess deposits can serve as continental environment change recorders. Magnetic susceptibility fluctuations in these deposits show a qualitative but unexpectedly detailed correlation with the marine oxygen isotope stages (SPECMAP [3]), which represent global climate signals as recorded by marine microfaunal shells. Precisely which of the three primary climatic parameters (temperature, rainfall, wind intensity), or a permutation or combination thereof, represents the sediment magnetic fluctuation, is still unknown. The simple geochemical model that has been proposed for enhanced topsoil magnetization derives from early work by LeBorgne [4]. His first model predicted that increased maghemite (γ-Fe2O3) production was due to natural fires converting weakly magnetic iron-bearing precursors (clay, amphibole). Kletetschka and Banerjee [5] proposed this as a possible mechanism for converting less magnetic parent material of Chinese loess to more magnetic topsoil. But the ubiquity of the phenomenon, irrespective of the amount of easily burnable grassland savanna in the Chinese loess plateau region, makes this mechanism a less likely overall explanation. LeBorgne's second mechanism, which he called ‘fermentation’, has been embraced much more strongly [4]. LeBorgne suggested that organic plant litter creates a complex reducing environment that would convert iron-bearing silicates to magnetite (Fe3O4). In the absence of such a reducing environment, the conversion of iron would proceed further to weakly magnetic hematite (α-Fe2O3). Given residence times on the order of a thousand years, LeBorgne [4] predicted that pedogenic magnetite would convert to only slightly less magnetic maghemite (γ-Fe2O3), which he identified as the putative magnetic mineral responsible for magnetism of topsoils. Research in the late 1980s and early 1990s supported the hypothesis of rainfall-driven over wind-driven magnetic topsoil formation [6,7]. However, statistical transfer functions constructed from a correlation of modern rainfall with modern topsoil magnetism [8] have been questioned by Han et al. [9], who point to important (but again only qualitatively so) correlations of magnetization not only with local rainfall but also with temperature. An alternative model for magnetic enhancement has been proposed by Dearing et al. [10] and Maher and Thompson [8], which hypothesizes that dissimilatory ironreducing bacteria (DIRB) are responsible for production of highly magnetic nanophase minerals (1–100 nm in size) in

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loess deposits. DIRB use poorly crystalline and weakly magnetic iron oxides (e.g., hematite) and iron oxyhydroxides (e.g., ferrihydrite, goethite) as terminal electron acceptors for metabolism [11–13]. This metabolism produces Fe(II) ions that can combine with pre-existing Fe(III) ions to produce nanophase magnetite [14], which has very high magnetic susceptibility due to its superparamagnetism. Such magnetite may even be the sole source of topsoil magnetization, without the requirement to produce maghemite, as LeBorgne had proposed. To return to the ultimate question of which climatic parameter, or combinations of parameters, ultimately controls the topsoil magnetism, we appear to be at a juncture where it has become critical to evaluate quantitatively the role of biotic (DIRB) vs. abiotic (presence of plant litter without microbes) controls on the production of nanophase magnetite. We allow the possibility that with the passage of time (a few hundred years to a thousand years) part or all of the nanophase magnetite may undergo ambient temperature oxidation in air to form metastable maghemite, which has a saturation magnetization that is only 10%–15% less than that of magnetite. The nature of the biotic versus abiotic pathway to magnetite production most likely encompasses several aspects. Nevertheless, we believe that a critical question is whether DIRB (or, in general, other microbes such as iron-oxidizing bacteria as well) are actively helping to produce nanophase magnetite, as opposed to the passive role of plant litter to create a less oxic environment in the topmost A-horizon of soils. Although the latter model may indicate that the length of the summer season may be a contributory factor, the former model may indicate a dominant role for the DIRB, which are substantially more active during warm/humid summer seasons than during the winter. Depending on the climatic zone of a site (semitropical vs. temperate vs. cold/arid), topsoil magnetic enhancement may thus be strongly influenced by the length of summer vs. winter (“seasonality” of climate) as well as ambient temperature, rainfall and wind intensity (responsible for delivering the parent material, i.e., the less magnetic iron oxides and silicates). In this contribution we report our first attempt to define and evaluate the role of microbes in general, and DIRB in particular, in influencing the magnetic variations in a 2.2-m deep soil profile in Nebraska. We have combined magnetic and chemical characterization with bacterial community analysis in order to develop an understanding of this complex material. Using our results, we are attempting to establish a transfer function for modern rainfall in the central U.S. where the parent

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material of the soil is predominantly wind-blown loess deposits, as in China. This is not the first attempt to quantify the role of microbes in producing magnetic topsoil in a loess-dominated site at temperate climates. In U.K. soils Dearing et al. [10] attempted (and failed) to detect magnetotactic bacteria, which produce internal chains of magnetic single-domain magnetite crystals. Hanesch and Petersen [15] carried out a topsoil microcosm experiment in the loess-derived brown soil (“parabraunerde”) of Germany. They enriched natural soil with an electron donor (lactate) and detected an increase in nanophase magnetite. They suspected DIRB to be the magnetic enhancer, but had no evidence beyond increased nanophase magnetite. 2. Methods 2.1. Site description and sample collection Our study site is located at Prairie Pines, approximately 5 km east of Lincoln (NE), a 145-acre property donated to the University of Nebraska in 2001. In 2002 we obtained a 2.2-m long core from the top of a nearly flat loess ridge using a hydraulic soil probe. According to its previous owners, Walter and Virginia Bagley, the 10-acre parcel around site PRA 02-A (N40.84242, W096.55912) has been grazed but never ploughed. The parent material consists of deep, massive Peoria loess, a late Pleistocene eolian deposit widespread in the midwestern United States [16]. The Lancaster County soil survey [17] shows the site as Sharpsburg silty clay loam, a fine smectitic mesic Typic Argiudoll, which is common on stable upland areas from southeastern Nebraska to northwestern Missouri as well as parts of Iowa and Kansas. Part of a collection of cores used by Geiss et al. [18], our core was immediately subsampled into sterile plastic

tubes, which were kept on ice in the field and stored in a refrigerator within hours of sampling. After subsampling, the core was described in the field using standard Natural Resources Conservation Service terminology [19]. Our description of site PRA 02-A is summarized in Table 1. Parts of the refrigerated samples were first used for the bacterial community analysis, and the remaining material was used separately for the magnetic and chemical study. 2.2. Bacterial community analysis Samples (0.5 g each) were suspended in 1 ml of lysis buffer (120 mM sodium phosphate, 5% sodium dodecyl sulfate, pH 8.0). Cells were lysed by performing three consecutive freeze–thaw cycles, followed by 90 min incubation at 70 °C and bead beating. Genomic DNA was extracted using a Fast DNA Spin Kit for Soil (Qbiogene; Vista, Calif.) per manufacturer's instructions. The quantity of DNA in these extractions was determined with a TD-700 fluorometer (Turner Designs, Sunnyvale, Calif.), using Hoechst dye 33258 and calf thymus as a DNA standard. Two different protocols were used to study the bacterial community structure within the soil sequence by denaturing gradient gel electrophoresis (DGGE) of polymerase chain reaction (PCR) amplified 16S rRNA gene fragments. The first protocol was used to study the general bacterial community while the second targeted the Geobacteraceae. The structure of the general bacterial community was analyzed by direct amplification of the variable V3 region of the 16S rRNA gene using the purified genomic DNA extracted from the soil as template. The primers for this PCR were PRBA338F (5′-ACT CCT ACG GGA GGC AGC AG-3′) [20] and PRUN518R (5′-ATT ACC GCG GCT GCT GG-3′) [21] primers, with a GCclamp [21] attached to the forward primer. The PCR

Table 1 Soil description of PRA 02-A Depth (cm)

Horizon Color

0–28 28–42 42–70

A AB Bt1

70–86

Bt2

86–100

BC

100–170 C1 N170 C2

Very dark gray (10YR 3/1) 10YR 3/2 Brown (10YR 4/3)

Texture

Structure

Comments

Silt loam Granular Heavy silt loam Fine subangular blocky Silty clay loam Medium subangular blocky Brown (10YR 4/3) Silty clay loam Coarse subangular blocky Yellowish brown (10YR 5/4) Silt loam Prismatic Common FE depletions (10YR 5/2) and concentrations (10YR 5/8) Yellowish brown (10YR 5/4) Silt loam Prismatic Common Fe depletions and concentrations Yellowish brown (10YR 5/4) Silt loam Massive Many Fe concentrations and depletions, many Mn concentrations, carbonates present below 217 cm

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protocol included a 5 min initial denaturation at 94 °C, 30 cycles of 92 °C for 45 s, 55 °C for 45 s, and 72 °C for 45 s, followed by a final extension at 72 °C for 10 min. Fingerprints of the bacterial community were also generated using a nested PCR-DGGE approach biased towards the Geobacteraceae, which include known dissimilatory iron reducing bacteria that belong to the Geobacter genus. The initial PCR amplified a fragment of the 16S rRNA gene using primers 8F (5′-AGA GTT TGA TCC TGG CTC AG-3′) [22] and Geo825R (5′TAC CCG CRA CAC CTA GT-3′) [23]. The PCR protocol included a 5-min initial denaturation at 94 °C, 30 cycles of 94 °C for 1 min, 55 °C for 1 min, and 72 °C for 2 min, and a final extension for 10 min at 72 °C. PCR products were then diluted 103-fold and used as template for PCR amplification using primers PRBA338F and PRUN518R. PCR mixtures (volume = 50 μL) contained 1 × PCR buffer with MgCl2, 4 nmol deoxynucleoside triphosphates, 25 pmol of forward and reverse primers, and 1.25 units of Taq DNA polymerase (Promega). PCR was performed using a PTC 100 thermal cycler (MJ Research; Watertown, Mass.). Denaturing gradient gel electrophoresis (DGGE) was performed using a D-Code apparatus (BioRad; Hercules, Calif.). Approximately equal amounts of PCR products were loaded onto 8% w/v polyacrylamide gels (37.5:1, acrylamide: bisacrylamide) in 0.5 × TAE buffer [24] using a denaturing gradient ranging from 25 to 50% (100% denaturant contains 7 M urea, 40% v/v formamide in 0.5 × TAE buffer). Electrophoresis was performed at 60 °C, initially at 20 V (15 min) and then at 200 V (180 min). The gel was stained with SYBR Green I (Molecular Probes; Eugene, Oreg.; diluted 1:5000 in 0.5 × TAE buffer), viewed on a UV transilluminator, and photographed with a CCD camera (BioChemi System; UVP; Upland, Calif.). Specific PCR-DGGE bands were manually excised from the gel, suspended in 20 μl of sterile water, and incubated overnight at room temperature. PCR-DGGE was repeated using these samples as template until a single band remained in each lane. A final PCR step was performed without the GC clamp attached to the forward primer. PCR products were then purified using the Geneclean II Kit (QBiogene) and nucleotide sequences were determined fully in both directions for each PCRDGGE band using PRBA338f and PRUN518r as sequencing primers. Sequencing was performed at the Advanced Genetic Analysis Center at the University of Minnesota using an ABI 3100 Genetic Analyzer (Applied Biosystems; Foster City, Calif.). Reported nucleotide sequences do not include the original PCR primer sequence. Reference nucleotide sequences were ob-

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tained from the GenBank database [25]. The nucleotide sequences obtained in this study have been deposited in the GenBank database under accession no. AY428951– AY428955. 2.3. Magnetic measurements Several types of magnetic measurements were performed on the samples that were collected for the comparative study (samples were dried, powdered and homogenized before analyses). Low field magnetic susceptibility and anhysteretic remanent magnetization (ARM) measurements were performed on standard cubic samples using a Kappa Bridge susceptometer (KLY2, Geofyzika) and a cryogenic magnetometer (2G), respectively. Hysteresis loops were acquired on a vibrating sample magnetometer (VSM, by Princeton Measurements) and low-temperature magnetic measurements on a magnetic properties measurement system (MPMS, by Quantum Designs), both using powders contained in gelatine capsules. Hysteresis loops in magnetic inductions up to 1.8 T were performed at room temperature, allowing extraction of several magnetic parameters such as coercivity (Hc), saturation magnetization (Ms), and magnetization of remanence after saturation (Mr). A first set of low temperature measurements consisted of the thermal demagnetization of an isothermal remanent magnetization (IRM) acquired at 10 K with an applied magnetic induction of 2.5 T, after having cooled the sample either in zero field (zero field cooling, ZFC) or in a 2.5 T induction (field cooling, FC). This experiment was followed by zero-field low-temperature cycling of an IRM acquired at 300K using a 2.5-T induction. A second set of low temperature experiments consisted of measuring AC magnetic susceptibility with an induction of 0.35 mT and AC frequencies of 1, 10, and 100 Hz. Details of these magnetic techniques can be found in Evans and Heller [2]. 2.4. Oxidation-reduction experiments The third component of our multi-disciplinary project examines the relative reactivity of the soils towards chemical reducing and oxidizing agents. Specifically, we wanted to determine the depth-dependent variation in oxidative and reductive reactivity of accessible iron to compare with, and confirm, the magnetic determination of iron oxide and hydroxide concentrations in the whole soil profile. The potential oxidizing and reducing behavior of the soils was characterized by quantifying the rate and extent of hydroquinone (QH2) oxidation and benzoquinone (Q) reduction [26,27]. All preparations

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Fig. 1. Examples of magnetic measurements obtained for samples located at various depths in the section. (a) Hysteresis loops in magnetic inductions up to 1.8 T. (b) Low-temperature field cooled and zero filed-cooled remanent magnetization acquired at 10 K in 2.5 T and low-temperature cycling of an IRM acquired at 300 K in 2.5 T. (c) Low-temperature variations of in-phase (χ′) and quadrature (χ″) magnetic susceptibility for field frequencies of 1, 10, and 100 Hz.

and reactions were performed in a catalytically maintained anaerobic environment (∼ 3% H2 in N2, vinyl anaerobic chamber, Coy Laboratory Products Inc., O2 b 100 ppm, as measured by an oxygen/hydrogen gas analyzer). Suspensions were stirred using Teflon®coated stir bars, and all reaction bottles were covered with aluminum foil to prevent exposure to light. Soil samples were ground using a glass mortar and pestle, and 150 mg of each was placed in a clean, 30 mL Nalgene bottle containing 5.0 mL of 40 mM, pH 3.75v acetate buffer (prepared using glacial acetic acid (Mal-

linckrodt, ACS grade) and NaOH) that had been purged with N2 gas for at least 20 min. Suspensions were tightly capped, removed from the anaerobic chamber, and sonicated for 10 min. After returning the bottles to the anaerobic chamber and stirring overnight, the appropriate volumes of either 10 mM hydroquinone (QH2, Sigma, 99%, reduction reactions) or 10 mM p-benzoquinone (Q, Acros, 99+%, oxidation reactions) stock solution and acetate buffer (to bring the final reaction volume to 25.0 mL) were added. The samples were stirred continuously throughout the experiment. At desired time

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intervals, 1.0 -mL aliquots were removed and filtered using a 0.2-μm Pall nylon filter membrane. The concentrations of QH2 at time t, [QH2]t, and Q at time t, [Q]t, were immediately quantified (b1 min) via High Performance Liquid Chromatography (HPLC). The stop time was recorded as the time of filtering. Blank samples containing only QH2 or Q in acetate buffer were used to account for the spontaneous oxidation of QH2 to Q and reduction of Q to QH2, respectively. The concentrations of Q and QH2 in each filtered sample were quantified using an Agilent Technologies 1100 Series HPLC equipped with a Zorbax® C18 Stable Bond column. The flow rate was 0.75 mL/min, and the mobile phase consisted of 65 vol.% 40 mM, pH 3.75 acetate buffer and 35 vol.% acetonitrile (Pharmco, HPLC grade). The detecting wavelength was 235 nm. The injection volume was 10 μL. Using this method, the retention time of QH2 was 2.4 min, and the retention time of Q was 3.4 min. An 8-point calibration curve from 0 M to 1 × 10− 2 M QH2 and an 8-point calibration curve from 0 M to 1 × 10− 3 M Q were used. 3. Results Examples of magnetic measurements performed on the samples are reported on Fig. 1. Room-temperature hysteresis measurements yielded average values of 8 mT for Hc, 0.12 for Mr / Ms, and 3.60 for Hcr / Hc. These values would place the magnetite mixture in the magnetic pseudo single domain range of the Day plot [28], although it would probably reflect the mixing of different grain sizes and/or mineralogy [29,30]. Hyster-

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esis data were also used to extract the para/dia-magnetic component of the magnetic susceptibility, which allowed us to calculate the ferrimagnetic susceptibility (Fig. 2) and hence, the concentration of only the magnetically-ordered minerals. Low-temperature magnetic measurements were performed to further improve our reading of the magnetic assemblage present in the samples (Fig. 1b,c). Results from FC and ZFC measurements indicate the presence of magnetite throughout the entire section, as shown by the existence of the Verwey transition in the cooling of the room temperature IRM (Fig. 1b). The separation between FC and ZFC curves, up to room temperature, and the large increase in magnetization upon cooling of the room temperature IRM, suggest the presence of high coercivity, magnetically unsaturated material such as goethite in the samples [31–33]. Results from AC susceptibility measurements show a decrease in the amplitude of the in-phase magnetic susceptibility (χ′) when the temperature increases, in part due to the presence of paramagnetic minerals in the sample (Fig. 1c). More importantly, it can be observed in the figures that the amplitude of χ′ is frequency dependent, which is also expressed by the non-zero values and the frequency dependency of the quadrature susceptibility (χʺ). These results indicate the presence of superparamagnetic magnetite (30 nm and below) in the samples. It is much more pronounced at the top of the core, suggesting that these grains are not directly inherited from the parent material, but rather incorporated or produced in the top layer during soil formation.

Fig. 2. Room-temperature magnetic parameters as a function of depth in the section. (a) Low-field magnetic susceptibility. (b) Ferrimagnetic susceptibility. (c) Ratio of ferrimagnetic susceptibility to saturation magnetization. (d) Ratio of anhysteretic remanent magnetization to remanent magnetization at saturation. Soil horizons are shown on the right side of the figure.

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Several magnetic parameters are reported as a function of depth in Fig. 2. The ferrimagnetic susceptibility (Fig. 2b) was obtained by subtracting the high-field slope of the hysteresis curves from the low-field magnetic susceptibility (Fig. 2a). Both data sets show higher values in the A horizon, indicating higher concentrations of ferrimagnetic minerals (e.g., magnetite, maghemite) at the top of the core. This is also reflected in the ratios χferri/Ms and ARM/Ms, which indicate higher concentrations of superparamagnetic and pseudo single domain grains in the A horizon, respectively. Such higher concentrations, particularly of superparamagnetic magnetite particles that are nearly absent in the parent material, could be due to in situ formation of ultrafine grains through iron reduction of Fe(III) minerals such as hematite, goethite or ferrihydrite. In Fig. 1b, the large increase in magnetization upon cooling of the room temperature IRM is interpreted as resulting from the presence of goethite in the samples. Given that goethite is a possible source for iron reduction, we attempted to follow the relative changes in goethite concentration in the samples. One major obstacle is that goethite is a very weak magnetic mineral that can easily escape magnetic detection when strongly magnetic magnetite is also present in a sample. We therefore designed a new experiment enabling us to target more efficiently this mineral, which is illustrated in Fig. 3. In the first step of the experiment, the sample is heated in the MPMS to a temperature of 400 K, which corresponds to the Néel temperature of goethite when the mineral becomes paramagnetic. A high intensity magnetic induction is then applied to the sample (usually 2.5 T) and the temperature is decreased down to 300 K, leaving the field on. This procedure tends to maximize the amount of magnetization carried by goethite [34] when cooling through its Néel temperature. The remanent magnetization of the sample is then continuously measured during a cooling/warming cycle between 300 K and 10 K. As it can be seen in Fig. 3a for a synthetic sample containing a mixture of goethite and magnetite standards the dominant signal is carried by magnetite, which displays a large drop of intensity at the Verwey transition characteristic of magnetite. At 300 K, the sample is removed from the MPMS and demagnetized using an alternating field (AF) demagnetizer with a peak field of 200 mT (the maximum field achievable with the instrument). The purpose of this AF demagnetization is to remove the signal carried by ferrimagnetic minerals with coercivities lower than 200 mT (e.g., magnetite), and therefore to enhance, by contrast, that carried by goethite. The sample is then placed back in

Fig. 3. Illustration of a new experiment targeting goethite, using a synthetic sample made of goethite and magnetite (a), a synthetic goethite sample (b), and a sample from a depth of 175 cm in our section (c). The sample is first heated to 400 K. At that temperature, a magnetic induction of 2.5 T is applied, and maintained while the sample is cooled back to room temperature. The remanent magnetization acquired by the sample is then measured during a cooling/ warming cycle between 10 and 300 K (steps 1 and 2). Next, the sample is demagnetized using an alternating field with a peak induction of 200 mT. The remaining remanent magnetization is consequently measured between 300 and 10 K (step 3), then from 10 K to 400 K (step 4), and back to 10 K (step 5). The difference in magnetization at 300 K between steps 3 and 5 is assumed to be proportional to the amount of goethite in the sample.

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Fig. 4. Comparison of magnetic data with microbial DNA as a function of depth in the section. Values at 290 K of the quadrature magnetic susceptibility (a) and of the frequency dependency of magnetic susceptibility (b) are taken as proxies for the amount of superparamagnetic magnetite particles. Mgeothite (c) is taken as a proxy for the concentration of goethite in the section. The total microbial genomic DNA (d) is used as a measure of microbial activity as a function of depth. Data set (e) represents the initial rate (black) and extent (light grey) of hydroquinone oxidation to benzoquinone as a function of depth.

the MPMS and the remanent magnetization is measured from 300 K to 10 K. On Fig. 3a, the Verwey transition of magnetite is suppressed after the AF treatment, and intensities are much lower than when measuring the sample without AF demagnetization, showing that the magnetite signal has been efficiently removed. In the subsequent step, the sample is warmed back up to 400 K, before being cooled again to 10 K. Since the Néel temperature of goethite is about 400 K, all the remanence carried by this mineral should be lost at 400 K, and the difference at room temperature between the magnetizations before and after thermal demagnetization should be proportional only to the amount of goethite present in the sample. In Fig. 3b, the same steps as in Fig. 3a are presented for a synthetic sample exclusively composed of goethite, which confirms that the magnetic and thermal behavior observed earlier for the mixture (Fig. 3a) after AF demagnetization is truly that of goethite. The procedure was applied to all the natural samples and is illustrated in Fig. 3c for a loess sample located at 175 cm below the soil surface. The goethite signal is clearly observed in this natural sample, although it can be noticed that some magnetization remains after thermal demagnetization of the goethite. This could be due to the presence of a very small quantity of minerals with coercivities higher than 200 mT and with Curie or Néel temperatures higher than 400 K. A small thermal hysteresis is present in the temperature range of 200 to 250 K near the Morin transition of hematite, which suggests the presence of

hematite in the sample, which could be responsible for the undemagnetized remanence. All the values obtained for Mgoethite are reported in Fig. 4c and compared to the quadrature magnetic susceptibility (Fig. 4a), the frequency dependency of the susceptibility (Fig. 4b), the total microbial genomic DNA (Fig. 4d), and the extent to which the soil reduces hydroquinone in batch experiments (Fig. 4e). It can be seen on this figure that lower concentrations of goethite are found in the top layer of the soil, which also corresponds to the layer where pedogenic superparamagnetic particles of magnetite are present, and the highest concentration of microbial biomass can be found. In contrast, the ability of the soil to oxidize hydroquinone (i.e., the susceptibility of ferric iron in the soil to be reduced by the reducing agent hydroquinone) is strongest at depths exceeding 50 cm, which corresponds to higher concentrations of goethite as found from Mgoethite measurements. Fingerprints of the total bacterial community were complex but virtually identical to each other down to a depth of 25 cm (Fig. 5). Below this depth, there was a gradual but substantial shift in community composition as well as a substantial decrease in the number of populations that were detectable by PCR-DGGE. The community profiles resulting from the nested PCR biased towards the Geobacteraceae were also complex and suggested substantial differences within the community as a function of depth in the section. Numerous bands were specific to the soil sections near the

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cesses. Finally, in experiments using benzoquinone in order to characterize the reductive character of the soils, no measurable reduction was observed. This could be the result of inaccessible reducing agents (i.e., the magnetite particles) or could be the result of the exceedingly low concentration of the magnetite nanoparticles in the soils (ca. 0.5 mg/g or less). Calculating the maximum expected hydroquinone production in the experiments yields a concentration that is similar to or less than our limit of detection. The lack of sensitivity for the reductive character of the soils highlights the superior strength of the magnetic methods for the characterization of exceedingly low concentrations of magnetic iron oxides. 4. Discussion

Fig. 5. PCR-DGGE fingerprint of the bacterial community, for various depths in the section.

surface (b 50 cm) or specific to the deeper section; no populations, however, were detected throughout the core. Ten prominent bands were excised and their phylogenetic identity was ascertained by nucleotide sequence analysis. Five bands were phylogenetically placed within the Deltaproteobacteria, which includes the Geobacteraceae, however they were most closely related (92–98% identity of ∼ 160 nucleotides) to other bacterial populations that had been detected without cultivation. The remaining bands grouped with the Acidobacteria, candidate division OP8, or failed to generate useful sequence data. Chemical characterization examining the relative reactivity of the soils towards hydroquinone, a chemical reducing agent, and benzoquinone, a chemical oxidizing agent, yielded interesting trends. First, the reduction experiments showed that a substantial amount of ferric iron is accessible for reduction. Fig. 4e shows the initial rate and extent of hydroquinone oxidation to benzoquinone as a function of depth. The maximum in both the initial rate and extent occurs with the 75 and 150 cm soil samples. Interestingly, the lowest reductive reactivity is observed in the soils with the greatest mass percent of DNA. This may imply reduced accessibility to ferric iron or depletion of reactive ferric iron by other pro-

Several mechanisms for pedogenic magnetite formation in loess soils have been proposed by different authors [4,8,10,35,36]. One major objective of this study, therefore, was to correlate the composition of the soil bacterial community to the properties of a Nebraska soil profile. Our results show that the total quantity of microbial biomass (i.e., bacterial plus eukaryotic microbes; here related to the quantity of DNA extracted from soils) changed as a function of depth in a pattern similar to that of the magnetic susceptibility. In parallel, the concentration of goethite, an important component of the soil parent material, is lower in the interval where superparamagnetic magnetite is formed. Furthermore, the location where superparamagnetic magnetite is formed corresponds to the location where the soil is least susceptible to ferric iron reduction. At the same time, the chemical assay (oxidation-reduction) experiments show a major loss of reactive ferric ion (from goethite, for example) in the superparamagnetic magnetite-rich levels. Thus, the synthesis of three sets of results provides circumstantial evidence that supports our hypothesis of magnetite formation via the reduction of goethite mediated by soil bacteria. In contrast, there was no obvious correlation between the composition of the overall bacterial community and the magnetic susceptibility profile. Because the resolution of PCR-DGGE is such that only bacterial populations comprising at least 1% of the total bacterial community can be detected [21], we performed nested PCR-DGGE that was specifically biased towards Geobacter spp., which are anaerobes capable of dissimilatory iron reduction [12]. But our bacterial community analysis data failed to support the hypothesis of bacterially-driven (or at least Geobacter-driven) magnetite formation in this loess deposit. This could, of

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course, mean that non-enzymatic iron reduction [37] is responsible for topsoil magnetic enhancement at this site. However, there are two possible scenarios in which DIRB could be responsible for the increased magnetic susceptibility but yet have gone undetected in our study. First, DIRB are broadly distributed among bacteria [38]; if non-Geobacter DIRB are responsible for the observed magnetic enhancement, then these DIRB would have been gone undetected in our Geobacter-biased study. Second, the DIRB responsible for magnetite formation, regardless of their species, could have been at extremely low population densities such that they would not be detectable. Maher and Thompson [8] proposed a mechanism in which magnetite formation occurs during intermittent periods of wetting that causes anoxic microzones to form in the soil. Under these conditions, only a small number of DIRB would be present in loess soils except during wet periods. Our contrasted multidisciplinary results point to the complex nature of the phenomenon and emphasize the need for additional basic research in this field [39]. 5. Conclusion In this study, a correlation between DNA content, bacterial community structure and magnetic properties of the soil was attempted in a Nebraska soil profile. Dearing et al. [10] showed that magnetic enhancement in top soil was probably not due to the presence of magnetotactic bacteria and suggested DIRB as a better candidate. In an attempt to identify the DIRB possibly responsible for the production of ferrimagnetic mineral in the soil, we performed a Geobacter-biased nested PCR-DGGE. This procedure was not conclusive in the sense that none of the populations were obviously Geobacter. This could mean that non-Geobacter DIRB are responsible for magnetic susceptibility enhancement through the formation of superparamagnetic magnetite. Alternatively, it could be argued that the magnetic enhancement is non-bacterial in origin. However, our multi-disciplinary data in this paper and the results of previous studies [e.g., [10,15]], all point to a close link between bacterial activity, iron reduction and magnetite formation. Therefore, we are currently conducting lab-scale experiments to stimulate DIRB in loess parent material to correlate increased magnetic susceptibility to identifiable bacterial community dynamics. Acknowledgements This research was supported by the U.S. National Science Foundation (CAREER program and Bio-

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