Appl Environ Microbiol 69(11): 6793-6800

Microbial Community Dynamics Associated with Rhizosphere Carbon Flow

Departments of Crop and Soil Science,^{Microbiology, Oregon State University, Corvallis, Oregon 97331}²

^{Corresponding author. Mailing address: Department of Crop and Soil Science, Oregon State University, 3017 Agricultural and Life Science Building, Corvallis, OR 97331-7306. Phone: (541) 737-5737. Fax: (541) 737-5725. E-mail:}ude.etatsnogero@dloryM.divaD.

^{Present address: Harvard Forest, Petersham, MA 01366.}

^{Present address: University of Georgia, Athens, GA 30606.}

Received 2003 Apr 7; Accepted 2003 Sep 1.

Abstract

Root-deposited photosynthate (rhizodeposition) is an important source of readily available carbon (C) for microbes in the vicinity of growing roots. Plant nutrient availability is controlled, to a large extent, by the cycling of this and other organic materials through the soil microbial community. Currently, our understanding of microbial community dynamics associated with rhizodeposition is limited. We used a ^{C pulse-chase labeling procedure to examine the incorporation of rhizodeposition into individual phospholipid fatty acids (PLFAs) in the bulk and rhizosphere soils of greenhouse-grown annual ryegrass (}Lolium multiflorum Lam. var. Gulf). Labeling took place during a growth stage in transition between active root growth and rapid shoot growth on one set of plants (labeling period 1) and 9 days later during the rapid shoot growth stage on another set of plants (labeling period 2). Temporal differences in microbial community composition were more apparent than spatial differences, with a greater relative abundance of PLFAs from gram-positive organisms (i15:0 and a15:0) in the second labeling period. Although more abundant, gram-positive organisms appeared to be less actively utilizing rhizodeposited C in labeling period 2 than in labeling period 1. Gram-negative bacteria associated with the 16:1ω5 PLFA were more active in utilizing ^{C-labeled rhizodeposits in the second labeling period than in the first labeling period. In both labeling periods, however, the fungal PLFA 18:2ω6,9 was the most highly labeled. These results demonstrate the effectiveness of using}^{C labeling and PLFA analysis to examine the microbial dynamics associated with rhizosphere C cycling by focusing on the members actively involved.}

Abstract

It is widely recognized that root-deposited photosynthate serves as an important carbon (C) source for microorganisms in the vicinity of growing roots. In turn, plants rely on the microbially mediated decomposition of organic materials for their supply of available nutrients. Several studies have examined the partitioning of photosynthate throughout the plant-soil system (e.g., see reference 23); a few of these studies have monitored the incorporation of this photosynthate (rhizodeposition) into the soil microbial biomass (e.g., see reference 21). Although these studies have revealed a great deal of information on C partitioning and soil organic matter dynamics, the microbial biomass is generally considered a single entity, which reveals nothing of the structure of the microbial community actively involved in nutrient cycling.

Several molecular and biochemical methods have been developed over the past decade that are useful for studying the great diversity of microorganisms in soil, most of which are unknown and unculturable (32). For example, analyses of microbial DNA and phospholipid fatty acids (PLFAs) have proven extremely useful for describing the general structure of soil and aquatic microbial communities. Analysis of PLFAs has been used to monitor changes in microbial community structure in response to several factors, such as agricultural management activities (6, 27, 28, 36) and heavy metal contamination (12). Analysis of PLFAs has also been utilized to examine the structure of rhizosphere microbial communities (14, 17, 18, 29, 30, 31). Although these studies have provided considerable information regarding the dynamics of microbial community structure in rhizospheres, they did not reveal any information regarding the function of microbial communities associated with rhizosphere C cycling.

Only within the last few years has the use of C isotope techniques coupled with PLFA analysis been exploited (4). Because ^{C-PLFA analysis reveals information on the active portion of the microbial community (}33), the isotopic labeling of individual PLFAs has the ability to directly link microbial processes with the groups of organisms involved. Boschker et al. (4) first used this approach with ^{C-labeled acetate and methane to examine which members of aquatic microbial communities were involved in two processes: sulfate reduction coupled to acetate oxidation and methane oxidation. Subsequently, this approach has been used to trace}^{C-labeled C substrates into several cultured strains of bacteria and fungi (}1), to examine the activity of soil bacterial and fungal biomarkers through incorporation of ^{C-labeled acetate (}2), and to link toluene degradation with specific PLFA biomarkers (16).

The primary objective of this experiment was to trace photosynthetically fixed C (^{C) through the microbial community associated with the rhizosphere and bulk soils of annual ryegrass (}Lolium multiflorum Lam. var. Gulf) during two different stages of plant growth. We hypothesized that spatial (i.e., rhizosphere or bulk soil) or temporal (i.e., growth stage) changes in the quality and/or quantity of rhizodeposition would influence which members of the microbial community would most actively utilize rhizodeposition. To test these hypotheses, we used a ^{C pulse-chase labeling procedure coupled with PLFA analysis. Another paper (}8a) reports on the distribution of photosynthetically fixed ^{C throughout the plant-soil system along with the turnover of}^{C through the microbial biomass pool. The present paper describes the dynamics of microbial communities associated with rhizosphere C cycling.}

Acknowledgments

This work was supported by a grant from the National Science Foundation (DEB00-75777) to Peter Bottomley and David Myrold.

We thank Stacie Kageyama and Jennifer Parke for assistance with evaluation of mycorrhizae, Anthony D’Amato and Lisa Ganio for providing insights about proper use of MANOVA statistics, and two anonymous reviewers, who provided thoughtful and helpful comments which improved the presentation and interpretation of our results.

Acknowledgments

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