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School of Environment and Natural Resources, Ohio State Univ., 2021 Coffey Rd., Columbus, OH 43210
* Corresponding author (Richard.Dick{at}snr.osu.edu).
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ABSTRACT |
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 TOPNOTES ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONSUMMARYREFERENCES |
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Abbreviations: D/R, drying and rewetting treatment • FAME, fatty acid methyl ester • G+, Gram positive bacteria, G–, Gram negative bacteria • PCA, principal component analysis • tDNA, total DNA • tFAME, total FAME
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INTRODUCTION |
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TOPNOTESABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONSUMMARYREFERENCES |
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A few studies have produced mixed results when comparing 4 or –20°C storage effects on microbial biomass C (by chloroform fumigation [Ross et al., 1980; Stenberg et al., 1998]) or fatty acid methyl ester (FAME) profiles (Schutter and Dick, 2000). Pesaro et al. (2003), using a PicoGreen assay and restriction fragment length polymorphism (RFLP) profiles, found that –20°C freezing and thawing caused a 24% reduction in soil total DNA along with a dramatic reduction in archaeal RFLP fingerprints. Enzyme activities after air drying are generally reduced (Pancholy and Rice, 1972; Bandick and Dick, 1999), but a few reports have shown specific enzyme activities to increase (urease, McGarity and Myers, 1967; arylsulfatase, Tabatabai and Bremner, 1970). Bandick and Dick (1999) found, for one soil type, that air drying either increased or decreased urease activity depending on soil management, but had no effect on arylsulfatase activity regardless of soil management. There are limited data showing that recovery of biological indicators varies with the assay method (Pesaro et al., 2004) and soil type (Fierer et al., 2003) after re-equilibration by drying, rewetting, and incubation under optimal conditions.
The studies to date by and large have determined the effects of soil storage on a single analytical method for one soil type. We found no lab incubation studies of storage effects on measurement of basal respiration and little known about the effects of ultrafreezing (–80°C) on soil microbial properties. In short, there have been no systematic studies that determine the effect of common storage methods or drying and rewetting of soil samples on a range of microbial properties and soil types. Therefore, the objective of this study was to determine the effects of sample handling and storage conditions on FAME profiles and total FAME (tFAME) concentrations, total genomic DNA (tDNA), seven enzyme activities, soil microbial biomass C, and respiration for three different soil types.
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MATERIALS AND METHODS |
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TOPNOTESABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONSUMMARYREFERENCES |
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Laboratory Analyses
Soil pH was determined in a 1:1 soil/water (v/v) ratio. Total C and N were measured by dry combustion (950°C) with a Vario Max CN analyzer (Elementar, Hanau, Germany). The particle size distribution was determined by the pipette method (Kilmer and Alexander, 1949).
Microbial biomass C was determined by the chloroform fumigation–incubation method (Anderson and Domsch, 1978). Cellulase activity (EC 3.2.1.4 ß-glucan 4-glucanohydrolase) was measured as described by (Gander et al., 1994) with the following modifications: samples were incubated for 24 h with 64 mM acetate buffer (pH 5.5), and the soil extracts were then diluted threefold with the same buffer after filtering through Whatman no. 2 filters.
N-acetyl-ß-D-glucosaminidase (EC 3.2.1.30 NAGase) activity was determined using the protocol of Ekenler and Tabatabai (2002). Analyses of arylsulfatase (EC 3.1.6.1 arylsulfate sulfohyrolase), ß-glucosidase (EC 3.2.1.21 ß-D-glucoside glucohydrolase), urease (EC 3.5.1.5 urea amidohydrolase), acid phosphatase (EC 3.1.3.2 orthophosphoric-monoester phosphohydrolase), and alkaline phosphatase (EC 3.1.3.1 orthophosphoric-monoester phosphohydrolase) activities were performed as described by Tabatabai (1994) with the following adaptations: for the analysis of arylsulfatase, ß-glucosidase, and acid and alkaline phosphatase, toluene was not used due to the short incubation times; and for urease, the NH4+–N product was measured using the colorimetric method described by Keeney and Nelson (1982).
Genomic DNA was isolated from 300 mg of soil, with two analytical replicates for each of the three soil replicates. The Bio101 Fast Spin Kit for Soil (Qbiogene, Inc., Carlsbad, CA) was used with slight modification. Two additional washes were done using the SEWS M buffer to minimize humic acids and other contaminants. A PicoGreen assay (Molecular Probes, Inc., Eugene, OR) was used for quantification of DNA, with relative fluorescence based on a standard curve, and determined using an ND-3300 Fluorospectrometer (NanoDrop Technologies, Wilmington, DE). The DNA extracts were diluted 20-fold to accommodate the calibration range of the assay. The total assay volume was 50 µL, and 2 µL were used for quantification on the NanoDrop.
Fatty acid methyl ester analysis was performed as described by Schutter and Dick (2000). Methyl nonadecanoate served as an internal standard, which allowed calculation of FAME concentrations (Zelles, 1996). The FAME detection and quantification were performed with a Hewlett-Packard 5890 Series II gas chromatograph (GC) equipped with a HP Ultra 2 capillary column and a flame ionization detector. The measurement was done with the MISystem, Version 4.5 (MIDI Inc., Newark, DE), using the TSBA 40 method. The GC temperature program ramped from 170 to 270°C at 5°C min–1. The reports generated by the MISystem software provided peak areas (response) and peak names (according to the peak match with the TSBA 40 method library). Standard nomenclature for the FAMEs includes the number of C atoms counted from the omega (
) end (i.e., opposite the carboxyl end), followed by the number of double bonds after the colon; cis conformations are designated with the suffix c, and the prefixes i and a are given for iso- and anteiso-branched FAMEs, respectively. The suffix 10 methyl indicates a methyl group at the 10th C atom, while OH stands for hydroxy and cyc for cyclopropane groups. The FAME compounds 18:26,9c and 18:19c served as fungal biomarkers (Kaur et al., 2005). The proportion of fungal FAMEs was calculated as a percentage of the total FAMEs, where fungal FAMEs were summed with bacterial FAMES (15:0, a15:0, i15:0, i16:0, 16:17c, 16:19c, 17:0, a17:0, i17:0, 17:0cyc, 17:18c, 18:15c, 18:17c, 19:0cyc). Markers for Gram positive (G+) bacteria were a15:0, i15:0, i16:0, a17:0 and i17:0. Selected monounsaturated and cyclopropane FAMEs served as indicators for Gram negative (G–) bacteria: 16:17c, 18:17c, 17:0cyc and 19:0cyc (Zelles, 1999).
Statistical Analysis
All microbial data were expressed on the basis of soil dry weight, and were tested for normal distribution (Kolmogorov–Smirnov test) before statistical analysis. Enzyme, tFAME, and tDNA data were normally distributed, but some parameters, including individual FAMEs, were log-transformed. Data comparisons between fresh soil and all other storage pretreatments were performed with the Dunnett's test. Significant differences between properties in the three soils within one pretreatment were tested with Duncan's new multiple range test. Correlations between tDNA, tFAME, and microbial biomass C data were calculated with SPSS for Windows (Version 14.01) using Pearson's correlation coefficient. Principal component analysis (PCA) was performed using PC-ORD for Windows (Version 4.01, MjM Software, Gleneden Beach, OR).
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RESULTS |
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TOPNOTESABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONSUMMARYREFERENCES |
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20 0ncrease in the Gilpin compared with fresh soil. Acid phosphatase was also unaltered at 4 and –20°C in two of the three soils (Hoytville and Gilpin). Cellulase activity clearly increased after –80°C storage (by 20 0n Granby and Hoytville and by 50 0n the Gilpin). Urease was unaffected after cold or frozen storage except in the Hoytville soil, where it was reduced by nearly half following all pretreatments. Arylsulfatase and alkaline phosphatase generally had lower activity for all storage methods in all soils.
Total DNA
The average tDNA extracted from the three fresh soils and the recovery percentage under various storage conditions is shown in Table 3. Recovery of tDNA from field-fresh soil was much higher in the Gilpin (forest) soil than the other two soils (agricultural). For the Granby, average tDNA recovered from field-fresh soil was significantly greater than for any of the other pretreatments except the soil stored at –80°C (P < 0.05). Storage at –20 and –80°C did not alter the tDNA concentration in the Hoytville soil. All storage pretreatments resulted in significant loss (32–51%) when compared with fresh soil for the Gilpin soil. Drying and rewetting resulted in significantly less tDNA compared with fresh soil in all three soils.
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In all soils, the G+/G– bacterial ratio increased due to soil D/R (Table 3) but the other storage treatments varied with soil type for this ratio. Storage at 4°C did not significantly affect the G+/G– ratio in the Granby soil, and it was similar for fresh soil and –80°C in the Hoytville. It was not affected by storage at 4, –20, or –80°C in the Gilpin.
The relative proportions of the fungal biomarkers in the Hoytville soil were not significantly different after the various storage methods. The fungal biomarkers in the Gilpin soil were reduced by D/R, whereas they increased in the Granby soil after storage at –20 and –80°C.
Correlation between Measures of Microbial Biomass
Total DNA, biomass C, and tFAME data were compared using simple linear correlation. Although these represent measurements of very different properties, they are each frequently used to estimate change in microbial biomass. Across all three soils and all storage treatments, tDNA and biomass C, tFAME and biomass C, and tDNA and tFAME were positively and highly correlated (r = 0.90, 0.90, and 0.92, respectively; P < 0.0001). When comparing these measurements in each of the three soils, however, correlation was only found between tDNA and tFAME in all soils (Granby: r = 0.80, P < 0.0001; Hoytville: r = 0.62, P < 0.015; Gilpin: r = 0.83, P < 0.0001). In addition, tDNA correlated with biomass C in the Granby (r = 0.53; P < 0.044) and Hoytville (r = 0.63; P < 0.011) soils.
Principal Component Analysis
Six PCA plots (two for each soil) (Fig. 1A and B) were obtained by analyzing two data sets per soil that contained measurements of microbial biomass and respiration, tFAME, tDNA, and enzyme activities (Fig. 1A), and unique FAME concentrations (Fig. 1B). Figure 1A plots from the Granby and Gilpin soils revealed that the fresh soil clustered close to the 4 and –20°C treatments. The Fig. 1A PCA plot for the Hoytville soil indicated that the sample pretreatments at 4, –20, and –80°C clustered together and were located closer to the fresh soil than the D/R pretreatment.
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DISCUSSION |
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TOPNOTESABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONSUMMARYREFERENCES |
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Recovery of tDNA was reduced by the D/R treatment. Pesaro et al. (2004) found that air drying reduced cell counts but not DNA concentration, and that after rewetting, there was a recovery of cell count but DNA concentration decreased by 50%. The probable explanation is that DNA from cells lysed by drying is not degraded until the soil is rewetted and populations recover and resume nuclease activity, causing rapid degradation of DNA from the lysed cells.
Total fatty acid concentration decreased with the D/R cycles compared with fresh soil. In our study, it appeared that the G+/G– ratio could serve as an indicator for microbial osmotic stress. In our case, G– bacteria were reduced after D/R, whereas G+ bacteria were less affected. This might be related to a strong cell wall of G+ bacteria (907fb3e69ceptidoglycan cross-linked by amino acids), whereas G– bacteria have 107fb3e69ceptidoglycan (Madigan et al., 1997). This explanation is consistent with our results, in which the drying–wetting cycle caused an increase in the G+/G– ratio (Table 3). Overall, the FAME data showed that D/R treatment caused a large decrease in total biomass and a shift of microbial community structure.
Storage at 4°C
Soils stored at 4°C were similar to fresh soils for acid phosphatase activity, ß-glucosaminidase activity, and the fungal markers across all three soil types, and for respiration in two of the three soils (Tables 2 and 4). Statistically significant differences in storage effects tended to be small for most of the enzymes. No other storage treatment had as many microbial measurements that were similar to the fresh soil across soil types. This is probably due to the fact that many enzyme activities are associated with extracellular enzymes stabilized in the soil matrix. These enzymes are not associated with viable cells, and during 4°C storage they remain protected from denaturation, unlike enzymes under treatments such as freezing or D/R, in which soil aggregates may be disturbed.
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In the present study, biomass C after 4°C storage was unchanged in the Granby and Hoytville soils, and significantly increased in the Gilpin. This may be related to the amount of bioavailable C in each of the soils. For example, the Gilpin soil had the highest organic C and probably could sustain microbial growth, as evidenced by an increase in biomass C when stored at 4°C over that of fresh soil. This same treatment increased ß-glucosidase activity, which hydrolyzes cellobiose and releases glucose, an important energy source for microorganisms. This would suggest a microbial induction of this enzyme in response to availability of low-molecular-weight C substrates. This follows Coxson and Parkinson (1987), who found a slow depletion of bioavailable substrates in refrigerated soil due to ongoing microbial activity. Even though microbial biomass C might stay the same when stored at 4°C, the community structure is probably changed. Evidence for this is seen in the Hoytville soil, where tDNA and tFAME decreased (57 and 50%, respectively), and PCA showed FAME profiles clustered separately between 4°C storage and fresh soil (Fig. 1B), but microbial biomass C remained unchanged by 4°C storage.
Frozen Storage
Freezing at –20 or –80°C resulted in little alteration of most microbial properties for the Hoytville soil when compared with fresh soil. For the other two soils, many microbial properties were altered by both of these storage temperatures. This is in agreement with other studies that have shown that freezing moderately affects soil microbial properties (Schutter and Dick, 2000; Petersen and Klug, 1994).
Respiration was unaffected by freezing except in the high-organic-matter Gilpin soil, where it increased over fresh soil by as much as 73%. The response of enzyme activities to storage pretreatments varied with soil type, but there was a trend toward –20°C having less effect on these assays than –80°C. This is shown in Tables 2 and 4, where from three to five enzyme assays were unaffected by –20°C storage across all soils, compared with a range of one to three assays unaffected by storage at –80°C. Activities of ß-glucosidase and cellulase increased, particularly at –80°C storage. This again, as mentioned above for the D/R cycle, is probably due to physical disruption and exposure of stabilized enzymes during the freezing and thawing cycle.
In two of the three soils, biomass C was unaltered by storage at –20 and –80°C, and tDNA was unaltered by –80°C storage. Sample storage at –80°C is frequently recommended before DNA extraction, and our results suggest that this is appropriate for the two agricultural soils. Forest soils with high organic matter content such as the Gilpin may be more susceptible to the effects of storage. Pesaro et al. (2003) found that freezing at –20°C reduced soil DNA concentration by 24%, and our results were in agreement, with a range of 18 to 35% reduction after –20°C storage. Total FAME was the only microbial property reduced (16–47%) after either freezing treatment across all soils. While reductions in tDNA and biomass C can only confirm overall losses in recovery of the microbial community, FAME profiles provide information about specific groups. After –20 and –80°C storage, all three soils clustered separately from the fresh soil in the PCA plots obtained from FAME profiles (Fig. 1B). This indicates that freezing changes the microbial FAME composition in soils substantially.
Effects of Soil Type
Storage treatment effects on microbial analyses varied with soil type with a few exceptions. Two of the enzyme assays (ß-glucosaminidase and acid phosphatase) and the FAME fungal markers were unaffected by 4°C storage across all soil types. Total FAME was reduced by all storage treatments in all soils, and the D/R pretreatment altered most microbial measurements in all soils. While microbial biomass and enzyme activities were mostly altered in the sandy Granby soil, however, respiration was either reduced (D/R and 4°C) or increased (–20 and –80°C) most prominently in the high-organic Gilpin soil. Total DNA also showed more significant reductions in the Gilpin soil.
Ranking of Soils
Clearly, maintaining microbial properties that represent field conditions is of critical importance for most soil microbiology studies. When microbial assays are to be used as soil quality indicators for commercial applications, however, pretreatment or storage methods are required that facilitate high throughput. For these applications, the exact value may not be as important as a consistent result across soil types or soil management systems that are being characterized. In other words, as long as a pretreatment results in a consistent ranking of soils or land management practices in the same order on a relative basis as fresh soils, the procedure could be calibrated for assessing soil quality.
To evaluate the microbial properties for consistency of ranking, soils were arranged from highest to lowest by microbial property and pretreatment in Table 4. The Gilpin soil generally showed the highest value measured in all sample pretreatments; for the other two soils, ranking varied with microbial property. The ranking among soil types was the same for microbial biomass and arylsulfatase activity, with the highest values in the Gilpin and the lowest value in the Granby soil. Total DNA revealed similar ranking (and significant differences) in fresh soil and soils stored at 4 and –80°C. Total FAME concentrations and fungal FAMEs indicated that storage had a profound effect on the microbial community composition since the ranking of each property changed by comparing the three soils. The ß-glucosidase and alkaline phosphatase activities showed similar ranking in the fresh soil compared with the D/R soil. The results for ß-glucosidase activity after D/R are consistent with Bandick and Dick (1999), who found that this assay maintains the ranking of soil management effects between fresh and air dried soil. Acid phosphatase and the G+/G– ratio were similar in fresh soil and after soil storage at 4°C.
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SUMMARY |
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TOPNOTESABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONSUMMARYREFERENCES |
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NOTES |
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TOPNOTESABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONSUMMARYREFERENCES |
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Received for publication June 28, 2006.
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REFERENCES |
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TOPNOTESABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONSUMMARYREFERENCES |
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0.05, Duncan's test) of microbial properties in three soils (Gi = Gilpin, Gr = Granby, Hy = Hoytville) within a sample pretreatment. The soils are ranked from left (highest value) to right (lowest value) for each microbial property (> indicates significant difference, = indicates no significant difference among soils within a given pretreatment). Italic ranking of the soils means the order of ranking was the same as fresh soil.