Published in Soil Sci. Soc. Am. J. 69:423-426 (2005).
© 2005 Soil Science Society of America
677 S. Segoe Rd., Madison, WI 53711 USA
Division S-3—Notes
SOIL GRINDING INCREASES THE RELATIVE ABUNDANCE OF EUKARYOTIC PHOSPHOLIPID FATTY ACIDS
V. J. Allison* and
R. M. Miller
Building 203, E-133, Environmental Research Division, Argonne National Laboratory, 9700 South Cass Avenue, Argonne, IL 60439-4843
* Corresponding author (vallison{at}anl.gov)
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ABSTRACT
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Pretreatment of soil samples, such as grinding and sample size, can potentially affect phospholipid fatty acid (PLFA) extraction efficiencies. The objective of this study was to determine how soil pretreatment affects PLFA analysis and interpretation. Two grinding (ground and control) and five soil sample sizes (0.125–5 g) in factorial combinations were applied to restored or remnant prairie soils. In the restored soil, the smaller soil aliquots were more likely to result in outlying values of community composition, suggesting small-scale heterogeneity in microbial community distribution. Grinding reduced this variability in community composition, with no outliers, even with the lowest size in ground soils. However, grinding increased the abundance of eukaryotic relative to prokaryotic PLFAs, and we hypothesize that grinding exposed root cells and the interior of fungal hyphae to extraction. We suggest that in soils with high root densities, soils should not be ground as grinding may obscure changes in the microbial community by exaggerating the eukaryotic signal from roots. However, in soils with low rooting densities, grinding will reduce heterogeneity and ensure that eukaryotic biomarkers are not underestimated.
Abbreviations: GC, gas chromatography • PLFA, phospholipid fatty acid • RA, reciprocal averaging
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INTRODUCTION
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PHOSPHOLIPID FATTY ACIDS maintain membrane fluidity and are present in the cell membranes of all organisms (White et al., 1979). Because they turn over rapidly during metabolism and are quickly degraded following cell death, amounts of PLFAs are proportional to viable biomass (Vestal and White, 1989; Frostegård and Bååth, 1996; Zelles, 1999). The chemical composition of PLFAs differs among microbial functional groups in terms of C chain length, branch position, saturation, and substitution (Vestal and White, 1989; Zelles, 1999), and consequently PLFAs can be used to characterize the microbial community.
In a previous study, we found variability in PLFA composition among replicates, suggesting small-scale heterogeneity may be obscuring broader patterns. This problem could potentially be resolved by homogenizing soils before extraction. In this paper, we examined the effect of pretreatment effects on microbial community composition by assessing relative abundance of PLFAs. Two soils were used: a remnant prairie soil with high organic matter content and microbial biomass, and a restored prairie soil with much lower microbial biomass. We predicted that larger sample sizes and grinding would reduce variability in community composition, assessed as the relative abundance of signature PLFAs.
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Materials and Methods
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Site and Experimental Design
Soils were collected from tallgrass prairie in the National Environmental Research Park at Fermi National Accelerator Laboratory (Fermilab), Batavia, IL, USA. The site had been under cultivation since the mid 1800s. Since 1975, prairie vegetation has been restored, now totaling approximately 450 ha of prairie. The soil is a Drummer silty clay loam (fine-silty, mixed, mesic Typic Haplaquoll), a deep, poorly drained soil that forms under prairie vegetation in level to depressed areas. Two sites were chosen that differ in organic matter content: an area restored 17 yr before this study (restored site), and a remnant prairie site adjacent to a railway line (remnant site). These sites are approximately 1000 m apart.
Three soil cores (4.8 cm in diameter by 10 cm deep) were taken at each site, approximately 10 m apart, in July 2002 and frozen within 5 h at –20°C pending further processing. Cores were defrosted overnight in a refrigerator, weighed, and then passed through an 8-mm sieve and then a 2-mm sieve to remove roots and break up large soil aggregates. Soil was freeze-dried (–50°C, 80 x 10–3 Mbar) in a Labconco Freezone 4.5 freeze drier (Labconco, Kansas City, MO). Any visible fine roots were removed from dry soil by hand picking. The study was a 2 x 5 factorial completely randomized design, with two grinding treatments (ground and control, unground), and five soil sample size treatments (0.125, 0.25, 0.5, 1.0, and 5.0 g), with three replicates. Grinding was for 30 s at room temperature in a Spex mill (Spex-Certiprep, Metucher, NJ). These treatments were applied to soil from both sites.
Soil Analyses
Soil C concentrations were analyzed using a Carlo Erba NC2000 elemental analyzer (Fisons Instruments, Milan, Italy), and averaged 3.8 ± 0.5% (SD) at the restored site and 10.1 ± 2.1% (SD) by weight at the remnant site.
Lipids were extracted from freeze-dried soil in a single-phase mixture of chloroform, methanol, and phosphate buffer (pH 7.4) in a ratio of 1:2:0.8, by an adaptation of the method described by Bligh and Dyer (1959). After 2 h, water and chloroform were added to separate the mixture into polar and nonpolar fractions, and total lipids were extracted from the nonpolar chloroform phase. The PLFAs were separated from other lipid classes by using silicic acid column chromatography, and methylated by using an adaptation of the mild-alkaline analysis described by White et al. (1979).
Before analysis, PLFAs were thawed and dissolved in 1 mL of hexane. Phospholipid fatty acid separation was by high-resolution fused-silica capillary gas chromatography (GC), using a HP5890 GC, with an HP7673 autosampler (Agilent Technologies, Palo Alto, CA). A 25-m HP-Ultra 2 column was used, with hydrogen as the carrier gas at a constant flow rate of 0.8 mL min–1. A 1-µL splitless injection was made for each sample, with the inlet temperature set at 290°C. The oven temperature was held at 60°C for 2 min, increased at 10°C min–1 to 150°C, and then increased at 3°C min–1 to 250°C and held for 5 min. Detection of PLFAs was by flame ionization at 320°C.
Phospholipid fatty acids were identified by retention time in comparison with known standards, and quantified using a bacterial quantitative standard (Catalog no. 1114; Matreya, State College, PA). Phospholipid fatty acids for which no quantitative standard peak was available were quantified by using the response factor for fatty acid 15:0. We chose to use an external rather than internal standard both to determine the degree to which PLFA response factors vary, and whether 19:0 (a commonly used internal standard) was present in samples. Although the absence of an internal standard can potentially lead to variation due to differential injection and evaporation, samples were run in randomized order. In addition, we find very consistent peak areas: over 20 injections, we found that the standard deviation of peak area was less than 3.5% of the mean (results not shown).
Phospholipid fatty acid nomenclature follows Tunlid and White (1992). Briefly, PLFAs are identified by C chain length, the number of double bonds, the position of the double bond from the methyl end of the atom, and the orientation (cis or trans) about the double bond. For example, the PLFA 16:1
5c (an indicator of arbuscular mycorrhizal fungi in some systems) is 16 carbons long, with one double bond positioned five carbons from the methyl end of the PLFA, and cis orientation about that double bond.
Data Analysis
Soil microbial community composition was assessed by using a reciprocal averaging (RA) ordination (McCune and Mefford, 1999) on the relative abundances of the 10 dominant fatty acids. Data from all soil sample sizes were used in this and subsequent analyses. Outliers were identified (more than 2.0 SD from the mean), but data did not require transformation to meet assumptions of normality and homogeneity of variance. The effect of grinding on abundance of individual fatty acids was also assessed, using a pooled variance t test for each fatty acid at each site (Systat 10; SPSS, 2000). Because of the large number of comparisons made, differences in this analysis were considered significant at p 
0.005.
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Results
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Axis 1 of the RA ordination plot explains 86% of the variation in PLFA composition, and corresponds to a division between sites. The restored site (squares) falls to the left on Axis 1, while the remnant prairie site (triangles) falls to the right (Fig. 1A). Axis 2 explains 8% of the variation in PLFA composition, and is determined by the grinding treatment; ground soils (open symbols) appear lower than unground soils (filled symbols) on Axis 2 (Fig. 1A). Grinding most markedly increases the relative abundance of PLFAs 18:26,9, 18:19c, and 18:17c (Fig. 1B).
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Fig. 1. (A) Reciprocal averaging (RA) analysis on relative molar abundance of 10 common fatty acids, in restored or remnant prairie soil, either unground or ground. Outliers (samples more than two standard deviations from the mean) are indicated by asterisks, and the soil aliquot size from which they were extracted. (B) Average position of each fatty acid within the ordination space.
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Grinding prairie remnant soils did not affect variability, with data points clustered no closer in ground than unground soil (Fig. 1A). Although this is also true of many of the samples from the restored soil, there are three outliers, all of which were extracted from unground soil of low sample size (Fig. 1A). Removal of outliers does not qualitatively affect results, with Axis 1 (90% of variation explained) still determined by site, and Axis 2 (5% of variation explained) dependent on grinding (results not shown).
When assessed as actual rather than relative abundance, the restored soil (Fig. 2A) was dominated by the fatty acids 16:0, 16:15c, and 18:19c while the remnant prairie soil was dominated by 16:0 and 18:19c (Fig. 2B). In the restored soil, grinding significantly increased abundance of 18:19c (t = –4.3564, p 0.0002), 18:26,9 (t = –3.8721, p 0.0006), and 18:17c (t = –3.1928, p 0.0035). In the remnant soil, grinding increased the abundance of 18:19c (t = –5.3219, p 0.0001), 18:26,9 (t = –8.3961, p 0.0001), and 18:17c (t = –5.7660, p 0.0001), and also significantly increased the abundance of 16:0 (t = –3.5782, p 0.0013) and had a marginally significant positive effect on abundance of 16:15c (t = –3.0237, p 0.0053).
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Fig. 2. Abundance (±SD) of the 10 dominant fatty acids in soil of the (A) restored and (B) remnant prairie, when unground and ground. Asterisks indicate that means of ground and unground soils were significantly different, using a pooled variance t test (p 0.005).
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Discussion
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Soil heterogeneity and aggregation can potentially influence measures of community composition (Schutter and Dick, 2002), and we proposed that consistency of PLFA analysis might be improved by grinding soils before extraction. We tested this prediction at two sites: a remnant prairie with high microbial biomass, and a restored prairie with considerably lower microbial biomass. We found that grinding improved reliability of community composition when assessed as the relative abundance of PLFAs, at the restored but not remnant site. Small-sized samples from the restored site were more likely to be outliers (Fig. 1A), when soils were unground but not when ground. However, grinding unexpectedly altered the relative abundances of signature PLFAs in both soil types (Fig. 1A), increasing the relative abundances of the eukaryotic PLFAs 18:26,9c and 18:19c (Zelles, 1997), and also of 18:17c (Fig. 1B). Although PLFA 18:17c is found in high concentrations in prokaryotes (Zelles, 1997), it is also found in eukaryotes including mycorrhizal fungi (Olsson et al., 1995).
We have two hypotheses to explain an increase in eukaryotic PLFAs: (i) grinding releases the fatty acids from fine roots in soil, and (ii) grinding more effectively exposes the interior surfaces of fungi to extraction. There is some evidence that this may occur. Olsson and Johansen (2000) found that ball milling before extraction increased the amount of PLFA 16:15c extracted from fungal hyphae by more than 200% relative to extraction from unmilled freeze-dried tissue. Similarly, we found that PLFA 16:15c was significantly increased by grinding at the remnant site (Fig. 2B). In addition, while 18:17c is generally regarded as a bacterial PLFA (Frostegård et al., 1993; Zelles, 1997), it is also present in arbuscular mycorrhizal fungi (Olsson et al., 1995), and thus the increase in this PLFA may be due to increased extraction from fungi.
Although arbuscular mycorrhizal fungi (AMF) comprise a substantial portion of the microbial biomass at this site (Miller et al., 1995), grinding increased other eukaryotic PLFAs to a greater degree than it increased PLFA 16:15c (Fig. 2). This suggests that the increase in eukaryotic signatures is at least partly due to extraction from fine roots, in spite of careful hand removal of all visible roots before grinding. Although Schutter and Dick (2001) found that addition of plant residues had little effect on fatty acid methyl ester (FAME) profiles, they incorporated oven-dried plant tissue into soil. We have found that oven-drying greatly reduces the amount of PLFA extracted (unpublished data, 2004), and suggest that viable plant tissue does have the potential to influence the PLFA profile. In a previous study, Petersen and Klug (1994) found that passing soil through a 2-mm sieve decreased the fungal signature 18:26,9, and suggest this was the result of damage to fungal hyphae. An alternative possibility is that sieving reduced the eukaryotic signature by removing roots from the soil.
Although the effect of grinding is substantially lower than that of site (Axis 1 explains 86% of the variation, while Axis 2 explains 8%), we nonetheless demonstrate that soil pretreatment significantly affects PLFA profiles of soils. We suggest that whether or not soils are ground before PLFA extraction depends on the rooting densities at a given site. In soils with high root densities, grinding may obscure changes in the microbial community by exaggerating the eukaryotic signal. In soils with low rooting densities, grinding will ensure that the eukaryotic signal is not underestimated.
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ACKNOWLEDGMENTS
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We thank Krissa Skogen for field assistance, Sarah O'Brien for laboratory assistance, and Julie Jastrow and Roser Matamala for helpful comments on the manuscript. This work was supported by the U.S. Department of Energy, Office of Science, Office of Biological and Environmental Research, Environmental Sciences Division, Global Change Research Program, under Contract no. W-31-109-ENG-39 to Argonne National Laboratory.
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NOTES
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The submitted manuscript has been created by the University of Chicago as operator of Argonne National Laboratory under Contract no. W-31-109-ENG-38 with the U.S. Department of Energy. The U.S. government retains for itself, and others acting on its behalf, a paid-up, nonexclusive, irrevocable worldwide license in said article to reproduce, prepare derivative works, distribute copies to the public, and perform publicly and display publicly, by or on behalf of the government.
Received for publication March 18, 2004.
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ABSTRACT
INTRODUCTION
