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This is to certify that the

thesis entitled

Enhancement of Soil Aggregation by the
Combined Influences of Soil Wetting and
Drying and Root-Microbial Associations

presented by
Fagaye Sissoko

has been accepted towards fulﬁllment
of the requirements for

M.S Soil Biophysics

 

degree in

 

Dr. Alvin ..M Smucker

 

Major professor

Date May 9, 1997

 

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ENHANCEMENT OF SOIL AGGREGATION BY THE
COMBINED INFLUENCES OF SOIL WETTING AND
DRYING AND ROOT-MICROBIAL ASSOCIATIONS

By

Fagaye Sissoko

A THESIS

Submitted to
Michigan State University
in partial fulﬁllment of the requirements
for the degree of

MASTER OF SCIENCE

Crop and Soil Science

1997

ABSTRACT

Plant root modifications of soil aggregates in the rhizosphere have been
reported frequently. However, there are conﬂicting reports of soil
aggregate stabilization by multiple cycles of root exudation and soil water
removal. Stimulation of soil microorganisms by root exudates was studied
in soils collected to depths of 20 cm in conventionally tilled (CT), no-tilled
(NT), and native grassland (NG) at the Kellogg Biological Station in the
southwestern Michigan. This study presents a method for measuring the
combined affects of root infusions of a mixture of 7 carbohydrates, 4
organic acids, and 12 amino acids compounds and root extractions of soil
water with multiple soil wetting and drying cycles. Polyvinyl chloride (PVC)
cylinders, 10 cm diameter x 12 cm length, prepared by drilling a
longitudinal row of 11 holes at 1 cm intervals, were used as containers for
soil samples. In this study, Rhizos soil solution samplers (SSS) have been
used to simulate plant roots. Soil microbial biomass and mean weight
diameter were estimated at the end of the wetting and drying cycles. Root
exudate compounds increased soil aggregate stability in macroaggregate
fractions by 5 times compared to the control after nine wetting and drying
cycles. Additions of root exudate compounds increased microbial activities
of soils adjacent to artiﬁcial roots for all management practices. Aggregate
stabilities of soils exposed to long-term tillage management, responded
more to additions of C and N compounds than did soil aggregates from

native grasslands.

To my wife Magnine Sidibe and daughters Ouriba and Sedinte

iii

ACKNOWLEDGMENTS

I wish to express my gratitude to Dr. Smucker for his guidance and helpful
suggestions as major advisor. Appreciation is also extended to Dr. Paul and Dr.
Perry for their helpful suggestions an for being members of my guidance
committee.

I want to thank Daniel, Santos, Yasemin, Ferguson for their supportive
assistance. I wish to express my sincere gratitude to Ravenscroft, Parker, Harris,
and Yabba for their assistance.

The ﬁnancial support provided by US—AID and MALI/SPARC Project is

fully acknowledged.

iv

Table of Contents

Page
1 .0 INTRODUCTION ..................................................................................... 1
2.0 LITERATURE REVIEW ........................................................................... 8
Soil structure ......................................................................................... 8
Wetting and drying forces on soil aggregation. ................................... 13
Soil water content ................................................................................ 14
Root exudates. .................................................................................... 15
Simulated root studies ......................................................................... 19
Ions and water uptake by plant roots. ................................................. 20
Soil organic matter. ............................................................................. 23
Microbial biomass. .............................................................................. 25
Summary. ............................................................................................ 32
3.0 SIMULATED ROOT EXUDATES ENHANCE SOIL AGGREGATION
DURING WETTING AND DRYING CYCLES
Abstract ............................................................................................... 32
Introduction. ........................................................................................ 33
Materials and methods. ....................................................................... 34
Results and discussion. ...................................................................... 50
Summary ............................................................................................. 58
References. ......................................................................................... 59

4.0 ROOT MODIFICATIONS OF SOIL MICROBIAL AND AGGREGATION
PROCESSES IN THE RHIZOSPHERE

Abstract .................................................................................................. 61
Introduction ............................................................................................ 62
Materials and methods ........................................................................... 64
Results and discussion .......................................................................... 71
Agregate stability ............................................................................. 71

Soil microbial activities. ................................................................... 92
Summary. ............................................................................................... 99
References. .......................................................................................... 100
5.0 CONCLUSIONS. .................................................................................. 103
6.0 REFERENCES. .................................................................................... 105

LIST OF FIGURES

Page

Fig. 1. Interactions among soil structural components and multiple

Fig.

Fig.

Fig.

Fig.

Fig.

Fig.

Fig.

Fig.

compounds of the soil. Adapted from Tisdall, 1991; Kemper
and Rosenau, 1986 ............................................................................. 2

. Wetting/drying, root exudates, microbes and ions modifications

of soil aggregate stability. ................................................................... 6

. Split PVC cylinder with holes on a longitudinal row. ........................ 37

. Laboratory conﬁguration for introducing carbon and nitrogen

compounds to “rhizosphere” regions of soil: 1) Split PVC

cylinder containing soil, 2) Simulated root exudate solution,

3) Peristaltic pump, 4) Manifold of 5 Rhizos SSS tubes, and

5) TDR probe. ................................................................................. 38

. Schematic drawing of laboratory Rhizos SSS system for

introducing carbon and nitrogen compounds to “rhizosphere”

regions of soil: 1) Porous teﬂon hydrophilic tube (average

porosity of 0.1 pm), 2) Stainless steel wire support,

3) Adhesives, and 4) Polyethylene tube. .......................................... 39

. Diagrammatic representation of sample collection from

rhizosphere and bulk soil when cyclinders were opened after
multiple wetting and drying cycles. (1) Rhizos SSS
microtube. ....................................................................................... 44

. Nebulization system used to pre-saturate soil aggreagtes

before wet sieving. 1) Nebulizer system, 2) Mist conduent,
3) Manifold of PVC tubes supporting 4 mm sieve, and 4) Screen
supporting soil aggregates ............................................................... .45

. An example of the change in soil water content of the

rhizosphere region of a Kalamazoo loam soil contained

in the PVC column and ﬁtted with 6 TDR probes and

5 Rhizos SSS microtubes during each wetting and drying

cycles ................................................................................................ 51

. Volumetric soil water content for samples collected in CT of

Kalamazoo loam soil subjected to nine wetting and drying
cycles during a 94-day period. .......................................................... 52

vi

Fig.

Fig.

Fig.

Fig.

Fig.

Fig.

Fig.

Fig.

10. Mean weight diameter (MWD) for soil aggregates
4.75 - 6.30 mm across, following simulated root C and N
compounds and nine wetting and drying cycles of a Kalamazoo
loam soil subjected to 10 years of CT and NT and 40 years

of NG, n= 3 ........................................................................................ 56

11. Mean weight diameter (MWD) for soil aggregates 2 - 4.75 mm
across, following simulated root C and N compounds and three
wetting and drying cycles of a Kalamazoo loam soil subjected to

10 years of CT and NT and 40 years of NG, n= 3 .............................. 74

12. Mean weight diameter (MWD) for soil aggregates 1 - 2 mm
across, following simulated root C and N compounds and three
wetting and drying cycles of a Kalamazoo loam soil subjected to

10 years of CT and NT and 40 years of NG, n= 3 ............................. .77

13. Percentage water stable aggregates 2 - 4.75 mm of a

Kalamazoo loam soil after three wetting and drying cycles .............. .78

14. Percentage water stable aggregates 1 - 2 mm of a

Kalamazoo loam soil after three wetting and drying cycles ............... 80

15. Mean weight diameter (MWD) for soil aggregates 4.75 6.30 mm
across, following simulated root C and N compounds and nine
wetting and drying cycles of a Kalamazoo loam soil subjected to

10 years of CT and NT and 40 years of NG, n= 3 ............................. 85

16. Mean weight diameter (MWD) for soil aggregates 2 - 4.75 mm
across, following additions of simulated root C and N compounds
and nine wetting and drying cycles of a Kalamazoo loam soil

subjected to 10 years of CT and NT and 40 years of NG, n= 3 ........ 87

17. Mean weight diameter (MWD) for soil aggregates 1 - 2 mm
across, following additions of simulated root C and N compounds
and nine wetting and drying cycles of a Kalamazoo loam soil

subjected to 10 years of CT and NT and 40 years of NG, n= 3 ........ 88

vii

LIST OF TABLES

Page
Table 1. Selected characteristics of the Agroecosystem site soil
at Kellogg Biological Station, 1994 and 1997 ........................................... 36

Table 2. Carbon and nitrogen compounds and quantities of
simulated corn root exudates used in laboratory injections
into soil columns of a Kalamazoo soil ....................................................... 42

Table 3. Inﬂuence of rhizosphere distances from root of 0 - 5
and 0 - 10 mm on microbial biomass estimates following
simulated C and N compounds and nine wetting anddrying
cycles of a Kalamazoo loam soil subjected to conventional
tillage (CT), n=3 ........................................................................................ 56

Table 4. Mean weight diameter for Kalamazoo loam soil collected from
three management practices which received 10 years of conventional
tillage (CT) and no tillage (NT), and from native grassland (NG).
Samples were air-dried then sieved to aggregate sizes from
2 - 4 mm across ........................................................................................ 74

Table 5. Inﬂuences of simulated root C and N compounds added
during three wetting and drying cycles of a Kalamazoo loam
soil, sampled from CT, NT, NG ﬁelds on the delta MWD
estimations of aggregate stability for sieved aggregates
ranging from 2 - 4.75 mm across, n=3 ...................................................... 75

Table 6 Inﬂuences of simulated root C and N compounds added
during three wetting and drying cycles of a Kalamazoo loam
soil, sampled from CT, NT, NG ﬁelds on the delta MWD
estimations of aggregate stability for sieved aggregates
ranging from 1 - 2 mm across, n=3 ........................................................... 78

Table 7. Inﬂuences of simulated root C and N compounds added
during nine wetting and drying cycles of a Kalamazoo loam
soil, sampled from CT, NT, NG ﬁelds on the delta MWD
estimations of aggregate stability for sieved aggregates
ranging from 4.75 - 6.30 mm across,n=3 .................................................. 84

viii

Table 8. Inﬂuences of simulated root C and N compounds added
during nine wetting and drying cycles of a Kalamazoo loam
soil, sampled from CT, NT, NG ﬁelds on the delta MWD
estimations of aggregate stability for sieved aggregates
ranging from 2 - 4.75 mm across,n=3 ....................................................... 85

Table 9. Inﬂuences of simulated root C and N compounds added
during nine wetting and drying cycles of a Kalamazoo loam
soil, sampled from CT, NT, NG ﬁelds on the delta MWD
estimations of aggregate stability for sieved aggregates
ranging from 1 - 2 mm across, n=3 ........................................................... 86

Table 10. Microbial biomass from samples collected in the three
management practices in Kalamozoo loam soil subjected to
10 years CT, NT and at least 40years of NG .......................................... 93

Table 11. Microbial biomass estimates of rhizosphere and bulk soils
following simulated C and N compounds and three wetting
and drying cycles of a Kalamazoo loam soil subjected to
Conventional tillage (CT), No tillage (NT), and Native Grassland
(NG), n= 3 ................................................................................................. 94

Table 12. Delta values of soil microbial biomass in rhizosphere and
bulk soils of aggregates from a Kalamazoo loam, <2 mm after three
wetting and drying cycles of the Ap horizon, n=3 .................................... 95.

Table 13. Microbial biomass estimates of rhizosphere and bulk soils
following simulated C and N compounds and nine wetting and
drying cycles of a Kalamazoo loam soil subjected to Conventional
tillage (CT), No tillage (NT), and Native Grassland (NG), n=3 .................. 96

Table 14. Delta values of soil microbial biomass in rhizosphere and
bulk soils of aggregates from a Kalamazoo loam, <2 mm after
nine wetting and drying cycles of the Ap horizon,n=3 .............................. 98

Table 15. Microbial biomass estimates of rhizosphere and bulk soils
following simulated C and N compounds and nine wetting
and drying cycles of a Kalamazoo loam soil subjected to
Conventional tillage (CT), No tillage (NT), and Native Grassland
(NG), n=3 .................................................................................................. 99

ix

INTRODUCTION

Soil structure, deﬁned as the arrangement of primary soil particles into
secondary particles or peds (Soil Sci. Soc. Am., 1984), controls and indirectly
describes the distribution of soil pore space. The development of a stable soil
structure is important for controlling several soil characteristics such as water
retention, gaseous diffusion, hydraulic conductivity, mechanical resistance, and
erodibility.

Processes controlling the formation of stable soil aggregates are complex
and include many biological, chemical, and physical components of the soil
system. Combinations of textural units, ions, water, organic matter, microbial and
fauna activities determine the various degrees of aggregate stability (T isdall,
1991). The development and stability of soil aggregates provide multiple
foundations for soil productivity and sustainable crop production. Therefore, it is
important to understand the roles of as many soil components involved in soil
aggregation as possible.

Individual factors, controlling the formation of soil aggregates, Fig 1 , have
not been completely separated to determine their relative importance. Soil
structure is greatly affected by clay content and mineralogy because of cohesive
and adhesive properties associated with the clay and organic matter (Kemper

and Rosenau, 1986).

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Climate is highly variable and an independent factor. Temperature has an

important affect on the rate of chemical reactions including those bonds of
biological metabolism, e.g. microbial biomass (Pawluk, 1988; Formanek et al.,
1984). Soil drying improves contact of particles by bringing them in closer
proximity. The strength of a single aggregate depends on the number of mineral
contact points (Horn, 1990). Utomo and Dexter (1982) investigated the effects of
wetting and drying on sterile and nonsterile soils which have been tilled, non-
tilled, and remolded. Their work demonstrated that variation in soil structural

stability was also affected by microbial activity.

Improving soil structure, particularly in agricultural production systems, is
an essential component to consider, when developing the best management
practice. Best management practices which augment or maintain soil organic
matter (SOM) content appear to have the most dramatic effect on soil structure
(T isdall and Oades, 1982). Agricultural activities, such as spreading manure,
adding imported plant residue, and rotating cultivation seasons with periods of
legume-based green manure crops, may counteract losses of SOM. However,
the level of SOM is seldom maintained by conventional agricultural production
practices. The inﬂuence of a cropping system on soil aggregation is a function of
the varied and combined effects of diverse physical, biological, and chemical

agents on the formation and degradation of soil aggregates (Harris, et al., 1966).

Microbial populations in the soil are responsible for the degradation of

plant tissue to produce humidiﬁed SOM. Microorganisms and plant roots exude

4
organic compounds into the soil which directly bind soil particles together. Root

exudates also serve as a rich source of carbon (C) and nitrogen (N) for many
microorganisms. Microorganisms prefer metabolizing fresh exudes to dead

tissue (Reid and Goss, 1982).

Fine roots have been also reported to physically bind soil particles
together (T isdall and Oades, 1982). They constitute a major source of organic
material for the soil and affect its structure, aeration, and biological activity.
Difﬁculties associated with past root measurements should not detract one from
the tremendous need to quantify numerous morphological and physiological
components of roots. This lesser known area of the plant-soil-atmosphere
continuum must be quantiﬁed before we can predict the ﬁxation and utilization of
carbon at the plant, landscape, or at the global level by either functional or
mechanistic models (Smucker, 1990). Rovira (1973) found that the major
exudation in plant roots occurs in the elongation zone. Root cap cells and
polysaccharide exuded at the apex are the major components of root
contributions to the rhizosphere. Baldock et al. (1987) studied bromegrass and
corn rotation effect on soil structural stability. No signiﬁcant differences in total
carbohydrates were found. In their study, eight sugars were found in all soils.
Corn roots had more galactose and glucose. Only very small aggregates (0.5
mm diameter) were found to have signiﬁcant differences in total carbohydrates.
Angers and Muhuys (1989) found that continuous cropping of corn for 2 years

decreased carbohydrate content by 30% and organic matter by 9 %. They found

5
a strong correlation between carbohydrate contents and mean weight diameter

of soil aggregates. When wetting and drying cycles are associated with root
releases of organic compounds (exudates), soil structure is usually improved
(Fig. 2). During the wetting and drying cycles, parallel clay crystals (about 5pm
diameter) are grouped together closely enough to behave in water as a unit
(Emerson, 1959, 1977).

The objectives and hypotheses of this study were:

1) To evaluate artiﬁcial corn root exudate contributions to soil aggregation in the
rhizosphere.

Hypothesis: Root exudates increase soil aggregate stability more in the
rhizosphere than in bulk soils.

2) To correlate soil microbial biomass with soil aggregate stability in the
rhizosphere.

Hypothesis: Higher microbial populations in the rhizosphere promote soil
aggregate stability.

3) To correlate soil aggregate stability with the frequency and duration of wetting
and drying cycles.

Hypothesis: Wetting and drying cycles improve soil aggregate stability.

4) To determine the relationship between soil management history and the effect
of C and N additions on aggregate stability.

Hypothesis: Soil aggregates are more stable in grassland than in conventional

and no tillage management systems.

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7
2.0 LITERATURE REVIEW

Soil structure

Soil aggregates, composed of primary particles and binding agents, are
the basic units of soil structure. Good structure for crop growth depends on the
presence of aggregates of soil particles 1 to 10 mm across which remain stable
when wetted (T isdall and Oades, 1982). Roots and hyphae stabilize larger soil
aggregates (macroaggregates) deﬁned as greater than 250 pm across.
Consequently, macroaggregation is controlled by crop and soil management
systems, as management generally inﬂuences the growth of plant roots (T isdall
and Oades, 1982). The importance of soil aggregation in crop production is
related to water and air relationships within the soil. A productive agricultural soil
must have a wide range of pore sizes (Lynch and Bragg, 1985), which allow
rapid inﬁltration and drainage (Tisdall and Oades, 1982). The deterioration of soil
structural stability is a major factor contributing to increased rates of soil
degradation by processes such as erosion, and compaction (Coote et al., 1988;
Wall et al., 1988), with consequent effects on productivity and the environment.

Aggregates having diameters of 20-250 pm are very stable, partly
because they are small, but also because they contain several types of binding
agents whose effects are additive. Water-stable aggregates 2-20 pm across
consists of particles bonded together so strongly by persistent organic bonds
that they are not disrupted by agricultural practices (T isdall and Oades, 1982).

Clay particles in these aggregate particles are oriented tangentially to the

8
bacterial surface. However in soils, perfect alignment of clay plates occurs rarely.

Small aggregates produced by slaking, remove clay particles and block some
pores (Arnold, 1978). In the ﬁeld, dispersed clay blocks pores which transmit or
store water. Slaking and dispersion together produce undesirable structures
such as surface crusts (T isdall and Oades, 1979). The stability of surface
aggregates is most important because aggregates below the surface are
protected from rapid wetting (Lynch and Bragg, 1985). When surface aggregates
are unstable, crusts can be formed and inhibit the movement of water and air
into the soil. Organic binding agents involved in stabilizing soil aggregates can

be considered in three main groups.

Transient binding agents are organic materials (polysaccharides) which
decompose rapidly and are associated with large aggregates. Readily available
substrates (glucose) increase water-stable aggregation which is transient
(several weeks) because the glues are decomposed readily (T isdall and Oades,
1982). The signiﬁcance of polysaccharides as glues in soil aggregates was

reviewed by several authors (Martin, 1971)

Temporau binding agents include roots and fungal hyphae, particularly
vesicular-arbuscular (VA) mycorrhizal hyphae (T isdall and Oades, 1979). These
binding agents build up in the soil within a few weeks or months as root systems
and associated hyphae grow, and are probably associated with young
macroaggregates (Tisdall and Oades, 1982). Stabilization of aggregates by fungi

in the ﬁeld is limited to periods when readily decomposable material are

9
available. Most of the microbial ﬁlaments which have been reported to stabilize

aggregates in the ﬁeld in the presence of plants may have been VA mycorrhizal

fungi (T isdall and Oades, 1978).

Persistent binding agents consist of partially degraded, aromatic humic
material associated with amorphous iron, aluminum and iron- aluminosilicates.
These form large organo—mineral fractions of temperate and tropical soil
(Turchenek and Oades, 1979) and are thought to be at the centers of highly

stable aggregates.

Inorganic binding compounds may be regarded as cementing agents. If
they are dominant, the presence of organic glues may be of little extra beneﬁt.
Regular cultivation may reduce the content of organic matter and the chemical
fertility, with little inﬂuence on the physical properties of such soils. However, in
the surface layers of many agricultural soils, it appears that organic matter plays
a major role on binding aggregates because of the rapid wetting and raindrop

impacts on aggregates at the soil surface.

Two forces holding particles together in aggregates in moist soils are the
surface tension of the air to water interface and cohesive tensions associated
with the liquid phase (Kemper and Rosenau, 1986). As soil dries, the water
phase recedes into capillary wedges surrounding particle-to-particle contacts and
ﬁlms between closely adjacent platelets. The interfacial tension and internal
cohesive tension pull adjacent particles together with great force as soil dries. As

the highly adhesive liquid phase of thin moisture ﬁlms pulls adjacent mineral

10
particles into closer proximity, there are also more opportunities for hydrogen

and other cationic bonding between adjacent oxygen and hydroxyl groups and

other mineral surfaces.

Much of the work on the stability of hydrated aggregates utilized elutriation
through various sized tubes to separate the aggregates into several sizes. Yoder
(1936) pointed out the deﬁciencies of this method. It is difﬁcult to avoid some
turbulence in water ﬂowing in wide columns, and some aggregates stay in the
tube if only laminar ﬂow occurred. Therefore, he developed a wet-sieving
procedure that used a nest of sieves which oscillate vertically and rhythmically,
so that water is made to ﬂow up and down through the sieves. Kemper and Koch
(1966) concluded that aggregate stability can be best determined by using a
single sieve. This involves much less investment and it can be well correlated
with important ﬁeld phenomena. It is generally considered that weight of large
aggregates is more indicative of good structure for most agricultural purposes
than is a weight of small aggregates (Tisdall and Oades, 1982). Van Bavel’s
(1949) concept of the mean weight diameter (MWD), based on weighing the

masses of aggregates of the various size classes, has been used widely.

The MWD is an index of the stabilities of soil aggregates of different size-

fractions. It can be calculated by equation (1).

MWD=£x,wi (1)

11
where x, is the size fraction of each sieve and w. is the proportion of the total

sample weight. The equation generally overestimates the original MWD where
only ﬁve separate size-fractions are used. The use of Van Bavel’s concept is
time consuming. However, the correlation using ﬁve size-fractions is excellent.
The geometric mean diameter (GMD) is an index of aggregate-size distribution.

It is calculated by the equation (2).

n n
GMD = exp [2 w, log x12: w. ] (2)
i=1 i=1
where wi is the weight of aggregates in a size class with an average diameter xi
and 2: wi is the total weight of the sample. Gardner (1956) found that the
aggregate size distribution in most soils is approximately log-normal rather than
normal. This approach describes aggregate-size distributions of most soils, using
two parameters, the GMD and the log standard deviation. The main
disadvantage of expressing data in terms of GMD and log standard deviation is
the extensive work involved in obtaining them. The log standard deviation must
be obtained by either graphical or differential interpolation from the data. The
best methods for presenting aggregate size distribution data seemed to be MWD
or GMD and the log standard deviation. The correlation coefﬁcient between
MWD and GMD was about 0.9. The GMD and the log standard deviation give a
more complete description of the size distribution than the MWD. However, the

MWD used for this study is easy to calculate and easy for most individuals to

12
visualize. Both can be used to represent aggregate size distribution for statistical

analysis.

Wetting and drying forces on soil aggregation

Wetting processes can be highly disruptive to soil aggregates. Ion
hydration and osmotic swelling forces pull water between clay platelets, pushing
the clay platelets apart and causing swelling of the aggregates (Kemper and
Rosenau, 1986). If soil aggregates are wetted slowly at atmospheric pressure,
the binding is still sufﬁciently strong to hold most of the primary particles together
in aggregates. Drying of soil allows particles to come in contact or closer
proximity, and the strength of a single aggregate depends on the number of
contact points or the forces that can be transmitted at each contact point (Horn,

1 990).

Wet-dry cycles can increase the mineralization of organic C and N in soil
(Birch and Friend, 1961; Seneviratne and WM, 1985). A proportion of the
increase in mineralization can be attributed to the death of signiﬁcant numbers of
microorganisms during soil drying (Van Gestel, et al., 1991), but most of the
increase in mineralization is due to increased availability of soil organic C. Wet-
dry cycles increase the amounts of organic C available to microorganisms
because of the additional exposure of organic C sequestered within the interiors
of soil aggregates. Disruption of micro-aggregates exposes C if located at the

interiors of broken soil aggregates, during drying and re-wetting (Powlson and

13
Jenkinson, 1976; Van Gestel et al., 1991). Sorensen (1974) showed that the

wet-dry cycles have a larger effect on the mineralization of more recently
accumulated, and possible more labile, pools of organic matter. Utomo and
Dexter (1982) found that 6 wetting and drying episodes of a ﬁne sandy loam soil
increased the stability of remolded aggregates by up to 4 times. Jager and
Bruins (1975) reported that soils can become completely structureless after 60
cycles of wetting and drying. Air-drying of clay and loam soils before wet-sieving
increased the stability of aggregates because of increased particle-to-particle
contact which favors the formation and adsorption of inorganic and organic

compounds (Haynes and Swift, 1990; Reid and Goss, 1982).

Soil water content

The variable amount of water contained in a unit mass or volume of soil,
and the energy state of water in the soil are important factors affecting the
growth of plants (Hillel, 1982). Many soil properties depend on soil water content
(swelling and shrinkage, bulk density, air content, and gas exchange of the soil).
Soil water content affects the growth and respiration of roots, the activity of
microorganisms, and the chemical state of the soil (oxidation-reduction

potential).

Volumetric water content: The volume wetness (often termed the
volumetric water content or volume fraction of soil water) is generally computed

as a percentage of the total volume of the soil rather than on the basis of the

14
volume of particles alone (Hillel, 1982). This author has shown that the

volumetric water contents of sandy soils ranges from 40 to 50 %; for medium-
textured soils it is approximately 50 %; and for clayey soils it can approach 60 %.
The Time Domain Reﬂectometry (T DR) technique has been used to measure

volumetric water contents (Dalton, 1992).

Root exudates

Plant root systems anchor plants, absorb ions and water for them. They
also produce quantities of C and N compounds by exudation and root
decomposition. For many years the quantity of organic compounds released
from roots were underestimated primarily because the results were based upon
measurements of plant roots grown in nutrient solutions and/or axenic conditions
(Smucker, 1984). This author showed also that root turnover or the death and
decomposition of ﬁne roots, may account for a large loss of organic compounds
into soil. Coleman (1976) demonstrated that root production is often the largest C
input to the ecosystem as 54 % of grass root tips appear to survive for less than
1 month. Healthy roots also release many organic compounds (Rovira et al.,
1978). Roots of many species have been shown to exude organic compounds of
varying composition and complexity. Organic materials lost from plant roots have

been classiﬁed by Rovira et al. (1978).

Root pxudates: Root exudates are substances (sugars, amino acids,

organic acids, hormones and vitamins) leaking (passively) from intact cells along

15
a concentration gradient. There are low-molecular-weight compounds, including

water-soluble and volatile compounds (Rovira, 1969, 1973; Pearson and
Parkinson, 1961). Root exudates diffuse through soil and are used by
microorganisms. The loss of root exudates appears to be proportional to the
concentration of these organic compounds at the root surface since removal of
the compounds increases exudation (Smucker, 1984). Secretions: Compounds
of low or high molecular weight (polymeric carbohydrates and enzymes) which
are actively released by root tissues. _l=y§a_te§_: Compounds released by autolysis

of epidermal cell walls and increase from increasing distance from the root apex.

Mucilageszare secreted by Golgi vesicles primarily in the root cap area. They
also originate from hydrolysates of the polysaccharide of the primary cell wall,
between the epidermal cells and sloughed root cap cells. Root-cap mucilage,
which accumulates in the corn rhizosphere as the tip extends through the soil, is
composed of polysaccharide molecules with many complex oligosaccharide

branches which have neutral sugars at their terminals (Watt et al. 1993).

Rovira (1969) listed several environmental conditions which affecte root
exudation. These include plant species, plant age, temperature, light, plant
nutrition, soil moisture, root damage, and foliar applications. Physiological age of
plants also affect root exudation composition and quantity (Martinez-Toledo et
al., 1988). Tyrosine is only released by tomato roots during the fruit formation
and is absent during the purely vegetative phases of growth. Rovira (1969)

showed that substantially smaller quantities of sugars, organic acids, and some

16
amino acids are released by plant roots during fruiting than in the earlier

vegetative phases of growth. He also suggested that soil microbes may affect
root exudation either by altering root metabolism or the permeability of cells, or
by the microbial assimilation of certain constituents of the root exudate. Rovira
(1969) showed that a wide range of organic materials, including sugars, amino
acids, and organic acids, are lost to the soil in the exudate, secretion, lysate
fractions. Many of these compounds are readily used by a wide range of soil
microorganisms, and they are largely responsible for the types and numbers of
microorganisms in the rhizosphere. Cheshire and Mundie (1990) showed that the
release of large amounts of water-soluble carbohydrates from com roots was
due to cell lysis. Root exudate solutions contained mainly glucose but also
galactose and mannose. Darbyshire and Greaves (1973) released the chemical
composition of wheat root exudate (12 organic acids, 12 sugars, and 25 amino
acids). Barber and Martin (1976) reported that gaseous and soluble C released
by root systems growing in soils or solution culture amounted to 20 to 39 % of
the C translocated to the roots. Carbon transfer from the root to the rhizosphere

results in a net loss of photosynthates from plant root systems (Smucker, 1984).

The rhizosphere is deﬁned as the volume of soil that is adjacent to and
inﬂuenced by the plant root (Hiltner, 1904). In another words, the rhizosphere is
the physical location in soil where plants and microorganisms interact. The work
of Lochhead et al. (1947) helped to understand the qualitative as well as the

quantitative differences between rhizosphere soil and bulk soil. Clark (1949)

17
demonstrated that the ratio of organisms in rhizosphere is different from the

organisms in the bulk soil. He demonstrated that not only the plant species and
soil water contents inﬂuence rhizosphere microorganisms but also the quantity of
soil adhering to roots from which suspensions were prepared for counting. He
proposed the term “rhizoplane” for the microbiology of the root surface. The

rhizoplane is deﬁned as the contact at the root-soil interface.

Polysaccharides deﬁned as high molecular weight carbohydrates
containing many (hundreds or thousands) of monomeric units connected to one
another by a type of covalent bond, referred to as a glycosidic bond (Brock and
Madigan, 1988). Cellulose is the simplest and most frequently occurring
structural polysaccharide of the cell wall. Tillage disrupts a variety of bonding
mechanisms in soil and reduces soil stability. It exposes SOM to microbial attack
and decreases the fungal biomass (Rovira and Greacen;1957; Gupta and
Gerrnida, 1988). Most of the temporary increases in porosity, following
cultivation, is the result of a ﬂush of biological activity of soil microorganisms,

resulting in a temporary improvement in aggregate stability (Molope et al., 1987).

Root exudates of corn consist of 61 % of carbohydrates, 8 % of proteins,
2 % of amino acids and 29 % of organic acids (Guckert et al., 1991; Buyanovsky
and Wagner, 1997). Glucose, arabinose, galactose, and mannose were the
predominant sugars. Brock and Madigan (1988) showed that amino acids are
monomeric units of proteins. Rovira and Greacen (1957) showed that the low

solubility of some amino acids (glutamic, glutamine, cystine, lysine, histidine,

18
arginine) inﬂuenced the relative abundance of these elements in the extracted

materials, giving an incorrect impression of the balance of amino acids in the root
exudate. The largest quantity of amino acids is synthesized by the root hair
zone, older parts of the root contain smaller amounts. Wallace and Lochhead
(1950) showed that the growth requirements of 80 % of the amino acid
dependent rhizosphere bacteria could be satisﬁed by supplying methionine as
the sole amino acid. They suggested that the methionine may be one of the

major amino acids excreted into the rhizosphere.

Simulated root studies

Information on the water and nutrient dynamics in the root is essential for
appropriate soil management and fertilizer application, since the root zone is the
area where soil and plant interact. Richards (1941) grouped the methods of soil
sampling into ﬁve categories: suction, displacement, compaction, centrifugation,
molecular adsorption. The only practical method to collect the soil solution
nondestructively is the suction method. A porous ceramic cup has been used as
a suction material (Debyle et al., 1989). This device has some limitation due to
the heterogeneous medium as root zone. Yanai et al. (1993) used a hollow ﬁber,
instead of a porous cup, as a soil solution sampler. Hollow ﬁber is a
semitransparent ﬁber used for hemo dialysis and water puriﬁcation, with internal
diameter of 475 pm and external diameter of 900 pm. It operates as a micro-

sieve, which blocks 90 % of particles 40 nm in diameter but allows solutes to

19
pass in spite of the elimination of suspension and colloid materials as well as

bacteria. To evaluate the performance of the sampler, Yanai et al. (1993)
examined ion adsorption-desorption. They found that the effect of the sampler
was too small to affect the composition and concentration of the soil solution.
Bouldin (1989) suggested that soil solution composition changes and ion
concentrations are interrelated. The looped hollow ﬁber sampler has outstanding

properties such as ﬂexibility, small size, and negligible ion exchange.

A Rhizos soil solution sampler (SSS), another device, was used to simulate
roots. It is produced from a hydrophilic porous polymer. This material is ideal for
greenhouses, laboratories, waste water sampling, and forestry studies. Sampling
by rhizos SSS is appropriate when successive samples form one position in soil

are needed.

Ions and water uptake by plant roots

Fresh plant matter is about 80 to 95 % water. Initial entry of water into the
root is mainly through that region which extends for a few centimeters behind the
tip (Epstein, 1972). The vascular tissue is the pathway of water in the root,
through the stem, and into the petioles and the veins of leaves. When water
moves through the soil to the root surface, it brings with it ions and other solutes.
If these arrive at the root surface faster than they are taken up, they accumulate
in the rhizosphere. Esptein (1972) showed that water uptake depends on root

length distribution in soil, phenomenological stage of the plant, air content, water

20
content and water conductivity of the soil, soil temperature and potential

transpiration. Osmotic component of soil water potential has little effect on water.
Water migrates throughout the mesophyll much as it does through the cortex of

the root.

Extensive root proliferation within the rooted zone result in rapid soil water
depletion. Meyer et al., (1990) showed that during a drying period, wheat roots
removed from an undisturbed soil 154 mm of water in 49 days (3.1 mm day"),
while in repacked soil they removed 174 mm in 57 days (3.1 mm day"). The rate
of root extension remained reasonably constant in undisturbed soil, but generally
increased with time in the repacked soil. The production of new roots enables the

exploration of new soil volume.

Soils contain considerable quantities of inorganic ions which are bound to
the charged surfaces of clay and organic soil particles. The gradual release of
these reservoirs of bound ions provides a continuous supply of nutrients for plant
absorption. Soil particles, particularly the silicates have an overall negative
charge and are capable of binding or absorbing cations added to solution.
Aluminosilicates are negatively charged because some of the O valencies

remain free.

The inorganic ions released by the root are in exchange for the cations
and ions absorbed into the root. Signiﬁcant release occurs when cations and
anions absorption are unequal. The degree of disparity between cation and

anion uptake varies with nitrogen, plant species, and stage of plant growth

21
(Esptein, 1972). Plant roots release organic materials as well as inorganic ions.

The effect of organic ions released by roots on diffusion or mass-ﬂow is related
to their effect on ion solubility. The increased concentration inﬂuence the

availability of other nutrients that reach the root mainly by diffusion.

At least 16 chemical elements (C, H, O, N, S, P, K, Ca, Mg, Fe, B, Mn,
Cu, Zn, Mo, and Cl) are required for plant growth. Eight of these elements are
absorbed in the form of oxides (C, H, O, N, S, P, Mo, B) while the other eight are
taken up as metallic ions. Traditionally, the elements have been divided into two
groups, the macronutrients and micronutrients. Macronutrients (C, H, O, and N)
make up more than 96 % of plant dry weight at ear leaf sample of com, the
nitrogen content is 2.76 - 3.5 %, phosphorus 0.25 - 0.50 %, potassium 1.71 - 2.5
%, calcium 0.21 - 1.00 %, magnesium 0.16 - 0.60 % and sulfur 0.16 - 0.50 %.
Micronutrient contents are generally reported as pg 9". Quantities of these,
compounds range from: manganese 20 -150, iron 21 - 250, boron 4 -20, copper
6 -20, zinc 20 -70 and molybdenum 0.1 -2.0 (Vitosh et al., 1994). Nutrients in soil
solution arrive at the surface of the root as a result of two kinds of movement:
bulk ﬂow and diffusion. High ion concentrations of the outer solution seem to
favor passive uptake processes (inﬂux). At low concentrations, net efﬂux may
occur through the pores. One third of the C ﬁxed by higher plants is converted to
cellulose, and massive amounts of lignin and aromatic derivatives are also
formed. Both NH: and NO; are commonly present in soil solution, and both are

readily taken up by roots. Potassium is absorbed in great amounts even when its

22
concentration in soil solution is low (Kolek and Kozinka, 1992). Potassium uptake

by root tissues is an active process (taken up against the electrochemical
potential gradient) (Ussing, 1949). Phosphate occurs in plants in its most highly
oxidized form. Phosphate absorption by roots takes place in epidermal cells, but

the main site of its uptake is the cortical tissue.

Soil organic matter

Organic matter consists of partially decayed plant residues that are no
longer recognizable as plant material, the microorganisms, the small fauna
involved in decomposition, and the by products of decomposition that undergo a
process called humiﬁcation to form the material known as humus (Paul and
Clark, 1996). The process of formation of soil organic matter or humus is
primarily a biological one, in which nearly all of the ﬂora and the fauna living in or
on the soil play a direct or indirect role. The microorganisms that are so active in
the decomposition of plant and animal residues are using a portion of these for
the building of their own bodies which become a considerable portion of soil
organic matter (Allison, 1973). Paul and Clark (1996) showed that an amino acid
(glucine) reacting with a phenol (catechol) derived either from the partial
degradation of lignin or from microbial pigments such as those produced by
fungus, form aminoquinone and can condense to form brown high molecular-
weight nitrogenous humates. The polyphenol reaction is considered important in

forming SOM from lignin or melanin-degradation products. The browning

23
reaction involves sugars (glucose) reacting with an amino compound such as

glucine to form three intermediates compounds (3-C aldehydes and ketones,
reductones, and furfurals). All of those react with amino compounds to form dark
colored end products. Nitrosation, a mechanism in which oxidized N reacts with
phenols, is often associated with N loss from soils. It is also believed to
participate in the formation of SOM (Paul and Clark, 1996). They listed the
factors affecting the decomposition of soil organic matter (composition and
particle size, microorganisms, available nutrients, water, temperature, pH, and

aeration).

The organic fraction of the soil consists of a complex system of
substances (Kononova et al., 1961). Components of decompositing plant and
animal residues, consist of various nitrogenous and non-nitrogenous organic
compounds (carbohydrates, organic acids, fats, waxes, resins etc.). These
compounds form 10 -15 % of the total amount of SOM. In developed soils, humic
substances form up to 85 - 95 % of the humus (Kononova, 1961). Humic acids
contain 50 - 58 % C and 5.0 - 5.5 % N. Soil humic materials generally have more
C than carbohydrates and amino acids (45 - 48 %). Ninety-ﬁve percent or more

of the nitrogen in soil surface is usually present in organic form (Allison, 1973).

Continuous cultivation increases the mineralization rates of SOM and
tends to exert a detrimental effect on soil tilth. These responses are enhanced
when soil physical conditions are suboptimal for mechanical manipulations

(Lynch and Bragg, 1985). Tillage affects soil structure in signiﬁcant ways.

24
Comparisons of virgin to cultivated soils have shown that cultivation profoundly

reduces the stability of soil structure (Elliot, 1986), such deterioration can occur
after a few years. Haynes and Swift (1990) found that long-term pasture samples
had a greater aggregate stability than long-term arable samples. They also found
that the aggregate stability of a regrassed site (13 years of arable plus 2 years of
pasture) was marvedly higher than that of a corresponding site from 15 years of
arable cropping. Mytton et al. (1993) suggest that clover is more effective than

ryegrass in developing rapid improvements in soil structure.

Microbial biomass

Aggregation within the surface horizons of many soils is predominantly a
function of the microbiological production and decomposition of soil binding
agents. In the presence of a suitable energy source, diverse fungi,
streptomycetes, and bacteria bind soil particles into water-stable aggregates.
The temporary increase in aggregation was frequently observed following
incubation of soils amended with organic materials, and is related closely to
microbial activity. Microbial metabolism of soil organic amendments is
accompanied by the production of organic aggregating materials which are
metabolized by the soil microﬂora in the absence of a more readily available
energy source. Microbial polysaccharides and fungal mycelia play major roles as
soil-binding materials. Fungal biomass in soil is the main component of the

microbial biomass.

25
Microbial biomass is deﬁned as the dry weight, volume, or other

quantitative estimation of organisms; the total mass of living organisms in an
ecosystem. The amount of biomass in soil can be assessed by the quantiﬁcation
of a particular cell constituent of the microorganisms. Jenkinson and Ladd
(1981) mentioned the basic requirements for this approach: the substance used
to estimate microbial biomass must be present in all organisms in the same
known concentration at all times. The microbial biomass itself may represent a
labile pool of C and nutrient elements. In agricultural soils 200-1000 pg biomass
C 9'1 soil is often found. This cell mass ﬁxes 100-600 kg N and 50-300 kg P

ha'1 in the upper 30 cm of soil (Martens, 1993). These amounts often exceed the
annual application of nutrients supplied as fertilizer to soils in agricultural
practice.

Soil microbial biomass is an important component of the soil organic
matter, and therefore, has an important impact on soil structure. It is a labile
component of the soil organic fraction containing 1 to 3 % of the total soil C and
up to 5 % of the total soil N (Smith and Paul, 1990). These materials are
converted by microorganisms in order to generate energy and to produce new
cellular metabolites to support their maintenance and growth (Martens, 1993).
Microbial biomass has been shown to be a sensitive indicator of differences in
sustainable cropping systems (Anderson and Domsch, 1989). The size and
activity of the soil microbial biomass must be assessed to understand nutrient

ﬂuxes in managed and natural ecosystems.

26
Comparative studies have generally measured temporal ﬂuctuations in

microbial biomass of natural and perturbed agroecosystems. The change in
microbial biomass values due to single effects such as tillage, soil type, climate,
and crops has been the focus of much research (Smith and Paul, 1990). Soil
microbial biomass has been used in studies of degradation of added organic
chemicals, residue decomposition, and polluted soils. The toxicity of pollutants
and the degradation of organic compounds can be monitored by following
changes in the soil microbial biomass (Horwath and Paul, 1994). These biomass
measurements when combined with tracer techniques will help answer questions
concerning soil processes of importance to agricultural management and
understanding the ecosystem functioning (Smith and Paul, 1990). We now seem
to understand the signiﬁcance of microbial biomass values, but the question
remains, “how does microbial biomass inﬂuence soil structure?”.

Among the methods to measure soil microbial biomass carbon, the four
most commonly used are; (1) chloroform fumigation (COz-C ﬂush), (Jenkinson
and Powlson, 1976); (2) substrate-induced respiration (SIR), (Anderson and
Domsch, 1978); (3) soil adenosine triphosphate content (ATP), (Jenkinson and
Oades, 1979); and (4) microbial biovolume derived by microscopy, (Jenkinson et
al., 1976). It is important to underline that none of these methods directly

enumerates the C content of the microbial biomass needing to be converted.

27
Chlpgpform fumigation incubation methpd (CFI)

The most common methods to measure soil microbial biomass are based
on chlorofon'n-fumigation. F umigation of soil with chloroform (CHCI3) increases
the amount of C extractable with 0.5 M of K2804 (Jenkinson 1966).

F umigation of soil samples with chloroform vapor causes a ﬂush of
decomposition during a subsequent 10 days incubation at 25° C compared with
an unfumigated soil (Jenkinson 1966). Chloroforrn fumigation may dissolve waxy
ﬁlms protecting decomposable substrates. The application CFI method is
confounded by the difﬁculty in ascertaining the contribution of non-microbial C to

the fumigation ﬂush (Fe) (Horwath and Paul 1994).

The chloroform fumigation extraction (CFE) method

CFE has few problems. It is rapid, although requiring additional analytical
procedures (i.e., C digestion followed by titration, Kjeldahl procedures for organic
N, etc.) for the determination of C and N which can lead to inconsistent results as
compared to CFI.

To obtain the microbial biomass C, the proportion (P) of the control (UFc)
should be subtracted from the fumigated ﬂush (Fe) and would vary as a function
of the ratio FclUFc. A linear function was used to determine the fraction of the
control to subtract from the fumigated ﬂush as described in equation (3):

P = K1(Fc/ UFC) +K2 (3)
where P is the fraction of UFc to subtract from Fe, K1 and K2 were estimated by

minimizing the sum of squares of the difference between MBC and the

28
microscopic biovolume. Therefore, microbial biomass C (MBC) can be calculated

by equation (4):

MBC = (Fc- UFc) P [0.41 (4)
Combining the two equations gives the formula in equation (5).

MBC = [Fc- (UFC(K1 (Fc/ UFc) + K2 ))] I 0.41 (5)

where K1= 0.224, K2= 0.33, Fc= fumigated and UFC= unﬁmugated. The CFI is
simple to perform, the estimation of C02 is easy. A critical concern is that
chloroform kills all microorganisms in soil which makes difﬁcult to estimate the
original microbial biomass. Also the direct quantiﬁcation of soil microorganisms
by other techniques does not correspond to the results of the fumigation-
incubation method. The fumigation with CHCI3 makes some of the humic fraction
of soils more available for degradation. The control problem when doing
fumigation is one of the most controversial, some authors try to estimate
microbial biomass without control but the result they obtain is twice as big as
when they use a control. Voroney and Paul (1984) advised caution in omitting a

control in measuring biomass in fumigation incubation.

The CFE method is rapid. It presents a good index for respiration. During the
process no ammonium immobilization or denitriﬁcation activity is noted and also
there is a low interference from nonmicrobial labile C and N substrates that can
be used during the incubation. CF E requires additional instruments. It is also

difﬁcult to measure small amounts of soil C.

29
The substrate-induced respiration (SIR) method was introduced by

Anderson and Domsch (1978) to rapidly estimate the amount of C held in living,
non-resting microorganisms in soil samples. Biomass estimate in amended soils
are of special interest under agriculture practice. In order to test the reliability
and limitations of the CFI, CF E, and SIR methods, agricultural soils were
amended with 1 % dried sewage sludge (Martens, 1985). Biomass C estimates
were carried out 3, 7, 17, 21, and 28 days after the addition of the organic
material. Three days after the amendment the CF I and SIR methods gave
undoubtedly erratic results. The CF I method showed the typical underestimation
caused by the problem with a large value for the control. The results of the SIR
method conﬁrmed the expected overestimation, indicating a shift in the
physiological ages of the microbial cells (Martens, 1985). With the SIR method,
amendments affected the biomass C estimates even after 4 weeks. Their results
showed that the SIR technique requires long pre-incubation times before reliable
estimates can be expected after the addition of a C source. Compared with the
results obtained at later sampling dates, the chloroform fumigation extraction
method obviously gave overestimates in two soils 3 days after the amendment

(Martens, 1985).

West et al (1986) pointed out that conversion factors obtained with a
relatively narrow range of soils may not be universally applicable. In studies on
New Zealand grassland soils they found a wide range of soil dependent

conversion factors between some of these methods. They evaluated and

3o
recalculated equations and common conversion factors given in the literature by

an application of statistical methods. For the kc factor of the fumigation-
incubation method they reviewed the published data and calculated a high
degree of variability. The relation between this method and the SIR method was
found to be uncertain.

The ATP extraction method was devised for the extraction and
measurement of adenosine 5'-triphosphate (ATP) in soil. ATP is a universal cell
constituent. It is present in all living cells and can be estimated with great
sensitivity by the luciferin-luciferase system. ATP extraction is rapid. One
disadvantage of the ATP method is that extraction of ATP from the cells is
usually incomplete and it is decomposed by enzymatic or chemical hydrolysis
during the extraction process. After extraction, ATP is strongly adsorbed by soil
constituents (West and Sparling, 1986).

Microscopic observations provide direct estimates of biovolume. However,
biovolume to weight, C, N conversions have remained problematic (Paul and
Clark 1996; Bottomley, 1994). Microscopy measurements are applicable to direct
ﬁeld studies. Microscopes are expensive and require care. Discrete organisms
with volumes greater than these categories were not included in volume
calculations. When required, volume was converted to biomass C by assuming
buoyant densities of 1.09 g ml’1 and dry matter contents (wlw) of 21 and 30 °/o for

mycelial organisms and bacteria respectively (West and Sparling, 1986).

31
Summary

This literature review is an overview of the interactions among plant roots
and their exudates, soil water, soil ions, and microbial biomass with a myriad of
activities involving soil aggregate development, function, and stability.

Carbon and nitrogen compounds from plant roots stimulate rhizosphere

microorganisms which contribute to soil aggregation processes. Frequent soil
wetting and drying cycles contribute directly to the positioning of soil particles.
Interactions of root exudates, soil ion and water contents, and augmented soil
cohesion improve soil aggregate stability. The best methods of measuring soil

aggregate stability and microbial biomass were reviewed and compared.

3.0 SIMULATED ROOT EXUDATES ENHANCED SOIL AGGREGATION

DURING WETTING-DRYING CYCLES
ABSTRACT

Plant root modiﬁcations of soil aggregates in the rhizosphere have been reported
frequently. However, there are conﬂicting reports of soil aggregate stabilization
by multiple cycles of root exudation and soil water removal. This study presents
a method for measuring the combined affects of root infusions of carbon and
nitrogen compounds and root extractions of soil water with multiple soil wetting
and drying cycles. Polyvinyl chloride (PVC) cylinders, 10 cm diameter x 12 cm
length, prepared by drilling a longitudinal row of 11 holes at 1 cm intervals, were
used as containers for soil samples. Several methods have been used to collect
soil solution nondestructively. In this study, a Rhizos soil solution sampler (SSS)
has been used to simulate plant roots. These small tubes, 2.5 mm x 100 mm
with an inside diameter of 1.4 mm, are constructed of hydrophilic polymers,
having an average pore diameter of 0.1 um. These root-like tubes were used to
extract soil water and exude carbon (C) and nitrogen (N) compounds, during
multiple cycles at one position in the soil. This study was designed to examine
the effects of C and N incorporation during wetting/drying cycles on soil
aggregation. A mixture of 7 carbohydrates, 4 organic acids, and 12 amino acids
was injected into soil by ﬁve porous Rhizos Soil Solution Samplers (SSS), and
submitted to 3 days incubation during each cycle. Soil water contents were

estimates by time domain reﬂectometry (T DR) during the wettint and drying

32

33
cycles. Soil microbial biomass and mean weight diameter were estimated at the

end the wetting and drying cycles.

INTRODUCTION

Several methods have been used to collect soil solutions
nondestructively. A large porous ceramic cup has been used to extract soil
solutions by suction (Debyle et al., 1989). This device has some limitations due
to the heterogeneous medium of the root zone. Yanai et al. (1993) used a hollow

ﬁber instead of a porous cup, as a soil solution sampler.

In this study a Rhizos soil solution sampler (SSS) was used to simulate
roots. This hydrophilic porous polymer is small, root-like, and can be used to
sample water contents in numerous soils. Extraction of soil solution and injection
of C and N compounds by Rhizos SSS seems appropriate for rhizosphere
measurements, especiallly when successive samples are injected into one
position of soil a volume.

Several methods for measuring the stability of soil aggregates have been
reported. The aim for aggregate stability tests is to describe and rank soil
behavior under rainfall in a way that can be related to real environments (Le
Bissonnais, 1996). Because of the range in ﬁeld conditions, under which the
samples are collected, the physical condition of samples needs to be
standardized before testing so that different soils can be compared. Major factors

in this standardization of methods includes soil moisture and structure of the

34
original sample. Numerous studies have shown that initial moisture content has a

large inﬂuence on soil aggregate stability when soil aggregate stability was

quantiﬁed by sieving (Haynes et al., 1990).

The activity of soil microorganisms and their inﬂuence on plants is largely
dependent on the utilization of substrates which originate either as root exudates
or plant residues. In this study, corn root exudates were simulated in the
laboratory to better understand the stimulatory contributions exudates have on
microorganism activities in soil aggregates during several wetting and drying
cycles. The chloroform fumigation incubation method was used to estimate

microorganism activities of the microbial biomass.

Speciﬁc objectives of the study were: 1) to evaluate artiﬁcial corn root
exudate contributions to microbial biomass by using Rhizos SSS microtubes, 2)
to evaluate the “exudation “ and “absorption” properties of Rhizos SSS

microtubes upon rhizosphere distances.

MATERIALS AND METHODS

Soil samples for this study were collected at Kellogg Biological Station in three
different soil management groups. The conventional tillage (CT) which is plowed,
the no tilled (NT), and the 40 years native grassland (NG). The percentage of
sand, silt and clay for the Kalamazoo loam soil samples used in this study are

listed in Table 1. The carbon and nitrogen contents for NG are high compare to

35

Table 1. Selected characteristics of the Agroecosystem site soils at Kellogg

Biological Station (K38), 1994, 1997.

 

 

 

 

pH Sand Silt Clay Org C N
1997 1997
9’;
CT 6.3 42.1 43.1 14.7 1.24 0.11
(0.03) (0.01)
NT 5.5 42.2 44.3 13.5 1.13 0.10
(0.10) (0.01)
NG 6.7 39.1 46.8 14.1 1.82 0.16
(0.09) (0.01)

 

Values in parentheses are standard deviations.
CT conventional tillage, NT no tilled, NG native grassland.

36
CT and NT. The ratios cm for the three management groups are 11 for CT, 10

for NT, and 12 for NG.

Soil column construction

Polyvinyl chloride (PVC) cylinders, 10 cm diameter x 12 cm length, were
prepared by drilling a longitudinal row of 11 holes, at 1 cm intervals, along one
side of the cylinder Fig. 3. Cylinders were then sawed in half, lengthwise through
the row of holes. Cylinders were reassembled and secured using duct tape
(Nashua, Watervliet, NY 12189). Four layers of cheese cloth, secured with duct
tape, were used to close one end of the cylinder. Soil aggregates 2 - 4 mm in
diameter, obtained by sieving air-dried soil samples, from each ﬁeld plot, were
uniformly packed into the PVC cylinders. The soil aggregates were saturated
from the underside by placing cylinders inside a container partially ﬁlled with
distilled water. Enclosed soil columns were saturated to the surface for 16 hours.
Following complete soil saturation, the second end of the cylinder was also
closed with four layers of cheese cloth and secured to the PVC tube by plastic

duct tape.

Simulated plant roots
Five Rhizos Soil Solution Samplers (SSS) (Ben Meadows Company. PO
Box 80549, Atlanta (Chamblee), GA 30366) were inserted into the saturated

soils at spacings of 2 cm, Fig. 4. An internal stainless steel wire, Fig. 5, with 0.8

 

: \...........3 I

12 cm

Fig. 3. Split PVC cylinder with holes on a longitudinal row.

38

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mm diameter and 150 mm length connected to the closed end of this porous

polymer tube, gives the sampler enough rigidity to be inserted it into the
soil.Stainless rods (3 mm x 100mm), sharpened on one end, were inserted,
parallel to each other above and below each artiﬁcial root for measuring
volumetric water contents of the rhizosphere by Time Domain Reﬂectometry

(T DR). Porous Rhizos SSS tubes were used for injecting simulated artiﬁcial corn
root exudate solutions. Rhizos tubes (2.5 mm x 100 mm) with an inside diameter
of 1.4 mm, were constructed of hydrophilic polymers which had an average pore
diameter of 0.1 pm. Solutions were added to the soil via the Rhizos tubes which
were connected to a manifold with PVC tubing (Fig. 4). A multiple channel
peristaltic pump (Cole-Parmer Instrument Company. Chicago, IL 60648) was

used for adding and withdrawing simulated corn root exudates and soil solutions.

Artiﬁcial root exudate solutions

Analyses of organic materials found on, in, or near corn roots reveal a
wide assortment of aliphatic amino acids, aromatic acids, amides, and sugar
compounds (Guckert et al., 1991; Paul and Clark, 1996; Buyanovsky and
Wagner, 1997). We simulated these corn root exudates by dissolving 12 amino
acids, 7 carbohydrates, and 4 organic acids at concentrations reported in Table
2. Carbohydrates constituted 63.35 %, amino acids 9.36 %, and organic acids
27.29 %. The pH of the solution was 2.2. In the mixture solution C content within

the solution was 40.74%, N 1.36%, H 6.69%, O 51.17%, and S 0.04%. The C/N

41

Table 2. Carbon and nitrogen compounds and quantities of simulated corn root
exudates used in laboratory injections into soil columns of a Kalamazoo loam

 

 

 

soil.
Carbohydrates gL'1 Amino acids gL’1 Organic acids gL’1
Arabinose 0.50 Alanine 0.02 Acetic acid 0.76
Galactose 1.45 Asparagine 0.16 Butyric acid 0.76
Glucose 1.31 Glutamine 0.16 Malonic acid 0.76
Mannose 1.04 Glycine 0.06 Succinic acid 0.76
Ribose 0.73 lsoleucine 0.02
Sucrose 1 .48 Leucine 0.03
Xylose 0.58 Lysine 0.03

Methionine 0.15

Phenylalanine 0.02

Proline 0.33

Tyrosine 0.16

Valine 0.1 1
Totals 7.06 1.25 3.04

 

Adapted from Guckert et al. (1991); Buyanosvsky and Wagner (1997); Paul and Clark

(1996).

42
ratio was 30:1.

Many other materials have been added to soils, such as straw and
sawdust, which contain C:N ratios from 80 and 400, respectively (Foth and Ellis,
1988). During the cycle, there is a decomposition and a continual loss of C as
respiratory C02, with an accompanying increase in the N percentage and
adecrease in the C:N ratio (Foth and Ellis, 1988). These added C and N
compounds were metabolized by the soil microorganisms during three day

incubation periods.

Wetting and drying cycles

The soil aggregates packed inside a PVC container were saturated for
16 hours. After saturation water was ﬁrst removed from the soil columns in a
manner which simulated root withdrawal of soil water by vacuum extraction
through the ﬁve Rhizos tubes, at the rate of 8.8 mllmin for two hours until air
entry and then at a minimum rate of 0.9 mein for 22 hours. After 24 hours of
water removal by the pump, water contents approximated ﬁeld capacity (22 to 25
%). A pulse of 5 ml of root exudate solutions plus a chase of 25 ml of distilled
water were added to soils adjacent to the ﬁve Rhizos SSS tubes in each
cyclinder at the rate of 6.3 mllmin. When the injection was completed, the
samples were incubated at 22° C for three days. During the incubation, soil
microorganisms metabolized the C and N added to the soil. After three days of

incubation, the cyclinders were air-dried by placing them on their sides with both

43
ends exposed to a high, 3.40 m3/min, ﬂow rate of air under the ventillation hood

(Kewaunee Scientiﬁc Equipment Corp. Adrian Michigan 49221), for four days.
When soil moistures approached air—dry water contents ranging from 2 - 5 %
(v/v), the cyclinders were removed for another cycle of saturation, vacuum
extraction pulse-chase, injection of root exudates, incubation and air drying. At
the end of the designated wetting and drying cycles, soil air-dry samples were
collected from the rhizosphere and bulk soil. A knife and saw were used to
separate rhizosphere and bulk soil from each cylinder as diagramed in Fig. 6.
Soils collected 1 cm around the Rhizos tubes were designated as rhizosphere
soil. The next 1 cm was collected as border soil to avoid the mixture of
rhizosphere and bulk soil. The adjacent 2 cm was collected as buk soil, and the
outlayer 1 cm , the portion in contact with the PVC was discarded to avoid the

effect of PVC.

Air-dried soil samples were prepared for wet sieving by saturating with a
nebulizer (Sunbeam Ultrasonic Humidiﬁer, Oster Household Products). A
humidiﬁer nebulizer was used to avoid rapid wetting of dry aggregates, causing
them to slake into smaller aggregates (Kemper and Rosenau, 1986). Four
preweighed and presieved samples, uniformly distributed on the top 15.2 cm
diameter sieve with openings of 4 mm were saturated by the humidiﬁer nebulizer
connected to four PVC plastic containers, Fig. 7. Uniform saturation of soil
aggregates by the humidiﬁer system was completed after 12 hours (Santos et al,

unpublished).

44

 

 

 

 

l
l
I 5m? I
|
I

Bulk soil I Border I Rhizosphere soil I Border I Bulk soil

Fig.6. Diagrammatic representation of sample
collected from rhizosphere and bulk soil
when cylinders were opened aﬁer multiple
wetting and drying cycles.

(1) Rhizos SSS microtubes.

45

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46
Aggregate stablllty

Soil aggregate stability was determined by using the wet sieving method
here after referred to as the Yoder method which is fully described by Kemper
and Rosenau (1986). Twenty grams of air-dry aggregates, subsampled for water
content, were saturated by nebulization, as described above, before placing on
the uppermost sieves of four nests. The sieving machine had an oscillation
speed of 33 rpm. Four nests containing 6 sieves (4, 2, 1, 0.5, 0.25, and 0.106
mm) were oscillated vertically for 20 min, so that water ﬂowed through the
screens containing the aggregates as described by Hillel, (1982).

After wet sieving, the nest of sieves were removed and the content of
each sieve transfered to a 50 ml beaker and oven dried for 24 hours. The
content of each sieve was weighed and thoroughly dispersed by adding 50 ml
sodium hexamethaphosphate (5 %), and dispersing in a machine (Multimixer.
Sterling Multi-products, Inc) by mixing for 20 min. The primary particles were
oven dried for 24 h and weighed. The mean weight diameter equation (1) was

used to estimate the aggregate stability, after correction for the coarse sands.

The delta value of the MWD was used for the statistical analysis. The delta value

is given by equation (6):
Am: ((MWD, - MWDc)/MWDC)*1OO (6)

where MVln)t is the mean weight diameter of the treatment (root water and root C

and N added), and MWDC is the mean weight diameter for water control.

47
Microbial biomass

The chloroform (CHCI3) used in the chloroform fumigation incubation
estimations of microbial biomass was triple distilled at 55° C with 5 % H2804
according to the method described by Jenkinson and Powlson (1976) to remove
ethanol and 10 g of anhydrous K2CO3 were added to the dry chloroform to
remove water.

Six subsamples of 25 g each, were weighed for microbial biomass
estimation. Three replicates were fumigated and three were unfumigated.
Vacuum desiccators containing these samples were lined with freshly moistened
paper towels prior to fumigation. For the fumigation treatment, the glass sample
bottles containing the soil were placed into a desiccator along with a 100 ml
beaker containing 50 ml CHCla. The desiccator was sealed and connected to the
vacuum pump, and evacuated until the chloroform boiled (four times). The
desiccator was sealed under vacuum and placed in the dark at 25 °C for 24
hours. The evacuation was repeated after 24 h and the samples were prepared
for incubation. The fumigated and control soil samples were placed in jars with
an open 20 ml scintillation vial containing 2 ml 2 M NaOH to trap 002. Three
black jars with no soil samples containing scintillation vials were placed to trap
the atmospheric C02. All the jars were tightly closed and incubated for 10 days
at 25 °C. Direct analysis of CO; was achieved by the double endpoint titration
method. An automatic titrator (Schott Gerate titrator, model TR 156, Yonkers, NY

10701 ) was standardized using 0.1 N HCI. Two ml of 2 M SrCIz were added to

48
the solution to precipitate the trapped CO; to strontium carbonate and titrated to

a pH of 7.0.
Soil water contents

Volumetric water contents were estimated by the Time Domain
Reﬂectometer (T DR) method (Topp et al., 1982; Campbell, 1990). Topp et al.,
(1980) proposed a polynomial which relates an empirical relationship between
volumetric soil moisture content and the apparent dielectric constant of soil (8).
They assumed that the dielectric constant of soil, for all practical purposes, was
insensitive to variations in bulk density, temperature, salinity, and mineral
composition.

Soil volume water contents were calculated by using the Topp, et al.,

(1982) equation (7).

e, = [-5.3«40’2 +(2.92*10'2*K,)-(5.5*10"*K.2)+(4.3*10-6*K,3)] .100 (7)

where K, is the apparent dielectric constant and K,= (c-rt/LZ); t is the signal travel
time in nanoseconds and t = (B-A)/(Vp*c); c is the propagation velocity of an
electromagnetic wave in free space and c = 30 cm/nsec; Vp=0.99; L is the length
of the transmission line (9.5 cm) which was the length of the TDR rod used in
these studies, A is the minimum and A the maximum. For this application, this

equation could be reduced to equation (8)

49
a, = - 0.053 + 0.02928 - 0.0005532 + 00000043.:3 (8)
where e is the electromagnetic wave in a medium of dielectric constant,

s= (c:2:-t2)/4L2

A critical step in TDR application to soil moisture determination is the
identiﬁcation of the second inﬂection point. The ﬁrst inﬂection point, which is the
junction of the probe and the transmission line, is unaffected by the soil water
content or salinity (Dalton et al., 1984). The second inﬂection point, which occurs
at the end of the probe, may be very weakly deﬁned in problem waves due to

noises in the circuitry (Dalton, 1992).

50
RESULTS AND DISCUSSION

Root extraction and infusion

Five Rhizos SSS tubes, connected to the peristaltic pump Fig.4, were
used to vacuum extract up to 8.8 ml per minute through the 5 Rhizos root-like
resulting in a speciﬁc absorption rate of 2.24 pl per mm2 of root surface tubes.
Soil water contents were reduced enough to permit air to enter the root-like tubes
while being pumped at this high rate for periods of two hours. Then tubes were
pumped at the rate of 0.9 mllmin (eg., 0.23 pl per mm2 of root surface) removing
both air and soil water for an additional period of 22 hours. Following this 24
hours “root extraction” period volumetric soil water contents, were reduced from
saturation 38.42 i 1.30, to 25.59 :t 3.77%, Fig. 8. Therefore, approximately 134
cc of soil water were removed from each soil column by the artiﬁcial roots during

each of the nine wetting and drying cycles, Fig. 9.

Following root water extraction, simulated corn root exudates were
injected into rhizosphere regions of the soil columns. Five ml of solution
consisting of 12 amino acids, 7 carbohydrates, and 4 organic acids, Table 2,
were injected at concentrations similar to root exudates (Guckert et al., 1991;
Paul and Clark, 1996; and Buyanosky and Wagner, 1997) assuming uniform
distribution among all ﬁve Rhizos tubes and uniform leakage ﬂow along the
surfaces of each tubes, then each Rhizos SSS injection of 56.75 pg of C and N
resulted in “speciﬁc root exudation” rates of 1.45 pg of C and N exudates per cc

of root surface. These C and N exudates were washed through the connecting

51

45

 

Percent soil water content (v/v)

 

 

 

 

O l T l l l l | l

O 25 50 75 100 125 150 175 200

Time (hour)

Fig. 8. An example of the change in soil water content of the
rhizosphere region of a Kalamazoo loam soil contained in the
PVC column and ﬁtted with 6 TDR probes and 5 Rhizos SSS
microtubes during each wetting and drying cycle.

45

 

2
40— Q

35-

 

 

30—

25—

20—4

15—

Volumetric soil water content (v/v)

 

10—

 

 

 

 

 

 

 

 

. . C C

0 l T l l l
0 500 1 000 1 500 2000 2500

Time (h)

Fig.9. Volumetric soil water contents for samples collected
in CT of Kalamazoo loam soil subjected to nine wetting and
drying cycles during a 94-day period.

 

 

 

53
lines and Rhizos tubes by 25 ml of distilled water. Each root injection and

subsequent washing resulted in a 3 - 4 % increase in soil water content ,
primarily in the rhizosphere regions of the PVC soil columns, Fig. 8. Microbes in
these soil columns metabolized these simulated root exudates for 3 days then
columns were air dried to approximately 3 % volumetric soil water contents,
Fig. 8.

Soil aggregate stabilization in response to root exudates and multiple soil
wetting, to saturation, and air drying were evaluated by repeating the processes
outlined above for 9 cycles, Fig. 9. There more similar volumetric soil water
contents of 38.42 :l: 1.30 % for the 9 saturation events and 3.42 :t 1.06 % for the
9 air dry events. Although soil water contents following root extraction 25.59 i
3.77 %, and root injection 29.04 i 2.32 % varied somewhat, all soil water
contents returned to nearly similar values during the 94-day period which

encompassed the 9 complete wetting and drying cycles, Fig. 9.

Soil microbial biomass

The distances which root C and N compounds moved beyond the
surfaces of the Rhizos tubes were evaluated by measuring microbial activities at
0 - 5 and 0 - 10 mm distances from the root surfaces as estimated by the
microbial biomass assay after 9 cycles of wetting and drying. Similar microbial

biomass values in experiments 0 - 5 and 0 - 10 mm, Table 3, suggest the Rhizos

54

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55

system is a repeatable estimate of root activities. Values for each soil position
and treatment are similar even though soil in the 0 - 5 and 0 - 10 mm
experiments were prepacked to different bulk densities of 1.30 g/cc and 1.15
g/cc. However, rhizospheres less than 5 mm appear to be too small for these
Rhizos experiments as signiﬁcant increases in the microbial biomass values
occurred between the C and N exudate treatments and the water controls of the
bulk soils, suggesting root exudates migrated into soil regions > 5 mm beyond
the root , Table 3. When rhizosphere soil samples were extended to 10 mm
beyond the root, then signiﬁcant differences in the microbial biomass values
could be detected between rhizosphere soils (0 - 10 mm) and bulk soils (> 10
mm) from the root surface. However, there were no signiﬁcant differences in the
microbial biomass values between the bulk soils of the water control and the C
and N exudates for the 0 - 10 mm experiment, Table 3.
Aggregate stability

Soil aggregate responses to simulated root exudation and root extraction
of soil water can be veriﬁed by measuring the mean weight diameter (MWD) of
soil aggregates adjacent to the Rhizos tubes ( ie., rhizosphere) and in bulk soil at
distances > 10 mm from the artiﬁcial root microtubes. The water stability of soil
aggregates, 4.75 - 6.30 mm across, removed from the rhizosphere region (ie., 0
10 mm) after 9 cycles of wetting and drying was always greater than for
aggregates removed from the bulk soil region of the soil column, Fig. 10. In tilled

(CT) and non tilled (NT) soils, additions and extractions of water and C and N

 

56

  

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and NT and 40 of NG.

Fig.10 Mean weight diameter CMWD) for soil aggregates 4.75-6.30 mm across,
following simulated root

n=3.

57
exudates by the simulated roots increased MWD of aggregates in the

rhizosphere 2 to 5 fold more than the water control without simulated roots.
Although this trend was also measured for soils taken from native grasslands
(N6), the contrast between rhizosphere and bulk soils were smaller. Aggregates
in the bulk region of the soil (eg., > 10 mm from roots) were less stable, lower
MWD, for soils subjected to all three management practices, Fig. 10.

These data suggest that wetting and drying partially disperse clay and
other ﬁne soil fractions from the surfaces of soil aggregates within the soil
column as observed by soil losses during the saturation phase of each wetting
and drying cycles (unpublished observations). Apparently water removal by the
nine vacuum extractions through the Rhizos tubes accumulated some of the
dispersed clay on soil aggregates located in the rhizosphere regions adjacent to
the simulated root (Reid and Goss, 1980). Subsequent additions of C and N
compounds stimulated microbial growth in the rhizosphere regions during the
3 day incubation periods. The resultant polysaccharides and other microbial
products (Lynch and Bragg, 1985) combined with the dispersed clay developed
more water-stable aggregates during the drying phase of each wetting and

drying cycle.

53
Summary

This experiment demonstrated the inﬂuence of artiﬁcial roots, root
exudates, wetting and drying modiﬁcations on microbial biomass and soil
aggregate stability. The presence of artiﬁcial roots used to removed water from
the soil during the wetting and drying cycles has an impact on the reorganization
of clay particles and microbial activity inside the cyclinder. Application of root C
and N as exudates improved aggregate stability by increasing the number of
microorganisms in the rhizosphere soil, and the quantity of larger aggregates
and their mean weight diameters. Aggregate stability increases in the
rhizospheres were 4 to 5 times greater compared to the baseline. The number of
wetting and drying cycles affect the aggregate stability. One of the powerfull
component of soil aggregate formation is the orientation of clay particles. The
further experiments can look the mechanisms of clay movement around the
Rhizos SSS, and the concentration of speciﬁc bacteria in the rhizosphere soil

which act as glues in soil aggregate formation.

59
REFERENCES

Buyanovsky, GA, and Wagner, G.H. 1997. Crop residue input to soil organic
matter on Sanbom ﬁeld. In Soil Organic Matter in Temperate
Agroecosystems. Long-Term Experiments in North America. Eds by Paul,
E.A., Elliot, E.T., Paustian, K., and Cole, C.V.

Campbell, GS. 1990. Dielectric properties and inﬂuence of conductivity in soils
at one to ﬁfty megahertz. Soil Sci. Soc. Am. 54:332-341.

Dalton, F .N. 1992. Development of time-domain reﬂectometry for measuring soil
water contents and bulk soil electrical conductivity. In: Advances in
measurements of soil physical properties: Bringing theory into practice
pp 140-167. (SSSA Special Publication no. 30).

Dalton, F.N., Herkelrath, W.N., Rawlins, D.S., and Rhoades, JD. 1984. Time-
domain reﬂectometry: Simultaneous measurement of soil water content
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Debyle, N.V., Hennes, R.W., and Hart, GE. 1989. Evaluation of ceramic cups for
determining soil solution chemistry. Soil Sci. 146:30-36.

Foth, H.D., and Ellis, BC. 1988. Soil fertility. Eds byJohn Wiley and Sons. pp.72.

Guckert, A., Chavanon, M., Mench, M., Morel, J.L., and Villemin, G. 1991. Root
exudation in Beta vulgaris: a comparison with Zea mays. Plant roots and
their environment. McMichaeI, B.L., and Person, H.(ed) Elsevier. pp.
449-462.

Haynes, R.J., and Swift, RS. 1990. Stability of soil aggregates in relation to
organic constituents and soil water content. J. Soil Sci, 41:73-83.

Hillel, D. 1982. Introduction to soil physics. Academic Press. San Diego,
California

Jenkinson, D.S., and Powlson, D.S. 1976. The effects of biocidal treatments on
metabolism in soi|:V. A method for measuring soil biomass. Soil Biol.and
Biochem. 8:209-213.

Kemper, W.D., and Rosenau, RC. 1986. Aggregate stability and size
distribution. Methods of Soil Analysis, Part 1. Physical and Mineralogical
Methods-Agronomy Monograph No. 9 (2nd ed.) Am. Soc. Agron.
Madison WI.

Le Bissonnais, Y. 1996. Aggregate stability and assessment of soil crustability
and erodibility: l. Theory and methodology. European J. of Soil Sci.
47:425-437.

Lynch, J.M., and Bragg, E. 1985. Microorganisms and soil stability. Advances in
Soil Science. Volume2. pp. 133-169.

Paul E.A., and Clark F.E. 1996. Soil Microbiology and Biochemistry. Academic
Press, San Diego.

Reid, J.B., and Goss, M.J. 1980. Changes in the aggregate stability of a sand
loam soil effected by growing roots of a perennial ryegrass (Lolium
perenne). J. Sci. Food Agr. 31 :325 -328.

Tisdall, J.M., and Oades, J.M. 1982. Organic matter and water stable aggregate
in soil. J. Soil Sci. 33:141-163.

60

Topp, G.C., Davis, J.L., and Annan, AP. 1980. Electromagnetic determination
soil-water content: measurement in coaxial transmission lines. Water
Resour. Res. 162574 582.

Topp, G.C, Davis, J.L., and Annan, AP. 1982. Electromagnetic determination of
soil water content using TDR: l. Application to wetting fronts and steep
gradients. Soil Sci. Soc. Am. J. 46:672-678.

Yanai, J., Araki, S., Kyuma, K. 1993. Use of a looped hollow ﬁber sampler as a
device for nondestructive soil solution sampling from the
heterogeneous root zone. Soil Sci. Plant Nutr. 39:737-743.

4.0 Root Modifications of Soil Microbial and Aggregation
Processes in the Rhizosphere

ABSTRACT

The aim of this study was to examine the effects of C and N incorporation and
wetting/drying cycles on soil aggregation. A mixture of 7 carbohydrates, 4
organic acids, and 12 amino acids was injected into soil by ﬁve porous Rhizos
Soil Solution Samplers (SSS), 10 cm long, 3 mm in diameter, and submitted to 3
days incubation periods during each cycle. Soils were subjected to three and
nine wetting and drying cycles to simulate natural soil conditions.The three
wetting and drying cycles did not generate the formation of aggregates 4.75 -
6.30 mm when a 2 - 4 mm aggregates were packed and submitted to wetting
and drying cycles. Carbon and N compounds accompanied by nine wetting and
drying cycles increased the mean weight diameter (MWD) in the rhizosphere 4.9
times in aggregates 4.75 - 6.30 mm. MWD increases in NT aggregates 1 - 2 mm
across, was 53 % when sampled from the root exudate rhizospheres. The
volumetric water contents were constant at saturation 38.05 %, after incubation
and 7 days air drying the water content was 2.94 %. Microbial biomass of the
samples collected from the three different management practices varied
depending on the season of sampling. Additions of root exudate compounds
increased microbial biomass of soil adjacent to artiﬁcial roots for all management
practices by approximately 50 % for NT soils. The application of C and N

compounds along with 9 cycles of wetting and drying caused signiﬁcant

61

62
increases in the microbial biomass and stabilities of soil aggregates in

rhizosphere adjacent to artiﬁcial roots.

Introduction

Soil aggregate stability, as determined by the wet sieving method, is
inﬂuenced by the number of wetting and drying cycles and the content of
microbial biomass carbon. The number of repeated wetting and drying cycles
have a great impact on the mean weight diameter (MWD) and the percentage of
water stable aggregates (VVSA). When an unstable air-dried aggregate is wetted
rapidly, it slakes into smaller sub-units (Emerson, 1977). Rovira and Greacen
(1957) and Utomo and Dexter (1982) found that aggregates collected from ﬁeld
soils and submitted to wet-dry treatments reduced aggregation by 14 to 48%.
Wet—dry treatments had signiﬁcantly reduced total macroaggregation of soil.
Total macroaggregation declined rapidly from 35 to less than 25% after two
cycles (Degens and Sparling, 1995). The ﬁrst wet-dry cycle resulted in large
declines in the proportions of aggregates from 0.25 - 0.5 mm, 1 - 2 mm, and > 2
mm by 54-65, 22-30, and 27-48%. When microorganisms were present, soil
aggregation increased in a few weeks, due primarily to microbial decomposition
of added organic material with the production of aggregating materials (McCalla
et al., 1957; Lynch and Bragg, 1985).

Wetting and drying cycles have been suggested to increase the amounts

of organic C available to microrganisms due to the greater exposure of organic C

63
during the disruption of microaggregates or soil pores during drying and re-

wetting (Powlson and Jenkinson, 1976; Van Gestel et al., 1991). The effect of
wetting and drying cycles on subsequent decomposition of organic matter
decreases with longer incubation of the organic material (Sorensen,1974). This
suggests that wetting and drying cycles have a larger effect on the mineralization
of more recently accumulated pools of organic matter (Degens and Sparling,
1995). Changes in aggregate stability in the rhizosphere have often been
attributed to the presence of root exudates (Oades, 1984; Reid et al., 1982;
Turcheneck and Oades, 1979). Due to their polysaccharide nature, root
exudates play a signiﬁcant role in the “cementation” of soil particles (T urcheneck
and Oades, 1979). The use of intact root mucilage, collected from maize plants,
produced additional evidence of the adsorption of exudates on clay minerals
(Habib et al., 1990; Morel et al., 1987).

The global objective of this study was to evaluate the separate and
combined affects of both root exudates and soil wetting-drying on soil aggregate
stability in the rhizosphere. Speciﬁc objectives were: 1) to evaluate the
contributions of artiﬁcial corn root exudates on microbial biomass activities; 2) to
evaluate soil aggregate stability responses to multiple wetting and drying cycles;
and 3) to estimate soil aggregate stability responses to both root exudates and

soil wetting and drying cycles.

Materials and Methods
Study site description and experimental design

Soils used for this study, collected at Kellogg Biological Station (KBS), are
classiﬁed as a Kalamazoo loam (Fine loamy, mixed, mesic, Typic Hapludalf: Soil
Conservation Service, 1990) near Hickory Comers, Michigan. Soil samples were
collected from plots subjected to three management practices. Two were from
agricultural sites which received 10 years of conventional moldboard plowing and
secondary tillage (CT), or no-tillage (NT), and the third was from a nearby native

grassland (NC), for the past 40 years.

Soil sampling

Soil samples from the Ap horizons of the three management practices
were taken to depths of 20 cm on November 25, 1995; May 2, 1996; and
November, 22 1996. Composite samples were then transported to the laboratory
and split into thirds. One third of each composite sample was sieved as moist
samples through nested sieves with 2 and 4 mm mesh sieves and was used to
estimate moisture content and microbial biomass within 24 h of sampling. A
second fraction was stored at 4° C, for estimating soil aggregate stability. A third
fraction was spread evenly on a greenhouse bench to air dry for 10 days. The

composite sample was then sieved to obtain a fraction from 2 - 4 mm.

65
Soil column construction and soil preparation

Polyvinyl chloride (PVC) cylinders, 10 cm diameter x 12 cm length, were
prepared by drilling a longitudinal row of 11 holes, at 1 cm intervals, along one
side of the cylinder (Sissoko and Smucker, 1997). Soil aggregates 2 - 4 mm in
diameter, obtained by sieving air-dried soil samples, from each ﬁeld
management system, were uniformly packed into the PVC cylinders at a bulk
density of 1.30 glcc for the ﬁrst 9—cycle experiment and 1.15 g/cc for the second
9-cycle experiment. The soil columns of aggregates were saturated from the
underside by placing cylinders inside a container partially ﬁlled with distilled

water. Enclosed soil columns were saturated to their surfaces for 16 hours.

Simulated plant roots

Five Rhizos soil solution samplers (SSS) (Ben Meadows Company. PO
Box 80549, Atlanta, GA 30366) were inserted into the saturated soils at spacings
of 2 cm (Sissoko and Smucker, 1997). In this study, Rhizos SSS tubes were
used to simulate roots. Rhizos microtubes consist of a hydrophilic porous
polymer which is constructed into a small root-like tube which can be used to
sample soil solutions. Extraction of soil solution and the injection of C and N
compounds by Rhizos SSS are most appropriate when successive samples are
extracted or added at one position within a soil.

Analyses of organic materials found on, in, or near corn roots reveal a
wide assortment of aliphatic amino acids, aromatic acids, amides, and sugar

compounds (Guckert, et al., 1991; Paul and Clark, 1996; and Buyanovsky and

66
Wagner, 1997). Corn root exudates were simulated by dissolving 12 amino acids

(9%), 7 carbohydrates (63%), and 7 organic acids (28%) in distilled water at
concentrations which composed an artiﬁcial root exudate containing 56.8 mg of

C and N compounds per injection (Sissoko and Smucker, 1997).

Wetting and drying cycles

Soil aggregates packed inside the PVC container were saturated for
16 hours. Water was ﬁrst removed from the soil column in a manner which
simulated root withdrawal of soil water, by vacuum extraction through the ﬁve
Rhizos microtubes until mostly air was withdrawn. At the end of each designated
number of wetting and drying cycles, air-dry samples were collected from the
rhizosphere and bulk soils (Sissoko and Smucker, 1997).

Air-dried soil samples were prepared for wet sieving by saturating with a
nebulizer (Sunbeam Ultrasonic Humidiﬁer, Oster Household Products). The
nebulizer was successfully used to avoid rapid wetting of dry aggregates, which
would have caused them to slake into smaller aggregates as reported by Tisdall
and Oades (1982). Four preweighed and presieved samples, uniformly
distributed on the top 15.2 cm diameter sieve, with openings of 4 mm, were
saturated by a nebulizer connected to four PVC plastic containers (Sissoko and
Smucker, 1997). Uniform saturation of soil aggregates by the nebulizer system

was completed after 12 hours (Santos et al, unpublished).

67

Soil microbial biomass

The chloroform (CHCI3) used in the chloroform fumigation incubation
estimations of microbial biomass was triple distilled at 55 °C with 5 % H2804
according to the method described by Jenkinson and Powlson (1976) to remove
ethanol and ten 9 of anhydrous K2CO3 were added to the chloroform to remove
water.

Six subsamples of 25 g each were weighed for microbial biomass
estimation. Three replicates were fumigated and three were unfumigated.
Fumigated samples were lined with freshly moistened paper towels. For the
fumigation treatment, the glass sample bottles containing the soil were placed
into a desiccator along with a 100 ml beaker containing 50 ml CHCI3. The
desiccator was sealed and connected to the vacuum pump, and evacuated until
the chloroform boiled (four times). The desiccator was sealed under vacuum and
placed in the dark at 25 °C for 24 hours. The evacuation was repeated after 24 h
and the samples were prepared for incubation. The fumigated and control soil
samples were placed in jars with an open 20 ml scintillation vial containing 2 ml 2
M NaOH to trap C02. Three black jars with no soil samples containing
scintillation vials were also established as controls to trap the atmospheric 002.
All the jars were tightly closed and incubated for 10 days at 25 °C. Direct
analysis of 002 was achieved by the double endpoint titration method. An
automatic titrator (Schott Gerate titrator, model TR 156, Yonkers, NY 10701) was
standardized using 0.1 N HCI. Two ml of 2 M SrClz were added to thesolution to

precipitate the trapped COZ to strontium carbonate and titrated to a pH of 7.0.

68
Mean weight diameter (MWD)

Soil aggregate stability was determined by using the wet sieving method
here after referred to the Yoder method as described by Kemper and Rosenau
(1986). Twenty grams of air-dry aggregates, subsampled for water content, were
saturated by nebulization, as described above, before placing on the uppermost
sieves of four nests. The sieving machine had an oscillation speed of 33 rpm.
Each nest containing 6 sieves (4, 2, 1, 0.5, 0.25, 0.106 mm) was oscillated
vertically for 20 min, so that water ﬂowed through the screens containing the
aggregates as described by Hillel, (1982).

After wet sieving the nest sieves were removed and the content of each
sieve transfered to a 50 ml beaker and oven dried for 24 h. The contents of each
sieve were weighed and thoroughly dispersed by adding 50 ml sodium
hexamethaphosphate (5 %), and dispersing in a machine (Multimixer. Sterling
Multi-products, Inc) was run for 20 min. The primary particles were oven dried for
24 h and weighed. The mean weight diameter equation (1) was used to estimate
aggregate stability, after correction for the coarse sands. The delta value of the

MWD was used for the statistical analyses. The delta value is given by equation

(6)

69

Percentage of water stable aggregates (WSA)

The air dry aggregates were placed on the uppermost sieve of the nest of
sieves and immersed in water. The nest of sieves were then oscillated vertically
forcing water to ﬂow up and down through the screens. After 20 min the nest of
sieves was removed from the water and the material left on each sieve was
oven-dried and weighed. The contents of each sieve were dispersed by adding
50 ml sodium hexamethaphosphate (5 %), and completely dispersed by mixing
in a dispersing machine for 20 min. The weight of sand retained after the second
sieving was then subtracted from the total weight of undispersed material
retained after the ﬁrst sieving. The percentage of water stable aggregates was
calculated by equation (9)

(weight retained) - (weight of sand)

WAS= x100 .(9)
(total sample weight) - (weight of sand)

 

Soil water contents

Time domain reﬂectometry (T DR) is a viable method for determining soil
water contents in small uniform sections of soils ranging from near saturation to
air-dry. The relationship between the dielectric constant and the volumetric water
content was found to be essentially independent of soil type, density, salt content
, and temperature (T opp et al., 1982). In this experiment volumetric water
contents in the rhizosphere region of the soil cores were monitored by TDR to
determine soil drying rates and uniformity of rewetting. Soil volume water

contents were calculated by using the Topp, et al., (1982) equation (7).

70
A critical step in TDR application to soil moisture determination is the

identiﬁcation of the second inﬂection point. The ﬁrst inﬂection point, which is the
junction of the probe and the transmission line, is unaffected by the soil water
content or salinity (Dalton et al., 1984). The second inﬂection point, which occurs

at the end of the probe, may be very weakly deﬁned in problem waves due to

noises in the circuitry (Dalton,1992).

71
RESULTS AND DISCUSSION

Aggregate stability

The _rr_1_e__a3 weight diameter (MWD) values of ﬁeld samples were different
for samples collected at different times of the year, Table 4. Samples collected in
November had higher MWD values compared to samples collected in May.
Lower MWD values for soil aggregates in May, after the frost had disappeared
from the soil, may have been the result of dilution and/or leaching of ions from
soils by frequent spring rains or could have resulted from lower microbial activity,
due to colder soil temperatures. Less stable soil structural units, during the
Spring of each year, could be one reason why some soil types are more

vulnerable to compaction by tillage in the Spring.

Three wetting and drying (w/d) cycles

Three wld cycles had little affect upon the stability of soil aggregates > 2
mm. No soil aggregates larger than 4.75 mm, could be extracted from NG, after
3 wld cycles. The MWD of soil aggregates, 2 - 4.75 mm, were the largest for NG
soils and smallest for CT soils, Fig. 11. MWD for water controls without Rhizos
roots appeared to be similar to or greater than water and C plus N exudate
treatments from the Rhizos microtubes. C and N additions to the rhizospheres of
Rhizos microtubes signiﬁcantly increased the MWD of aggregates 2 - 4.75 mm
across, from CT soils, but not from NT nor from NG soils Fig. 11 and Table 5.

Three wld cycles signiﬁcantly increased (positive delta MWD values) the

72

Table 4. Mean weight diameter for Kalamazoo loam soil collected from three
management practices which received 10 years of conventional tillage (CT) and
no tillage (NT), and from native grassland (NG). Samples were air-dried then
sieved to aggregate sizes from 2 - 4 mm across.

 

 

 

 

 

Mean weight diameter (MWD)
Date CT NT NG
mm
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May 1996 1.17 1.20 1.53
(0.09) (0.05) (0.06)
November 1996 1.38 1.90 2.14

(0.26) (0.06) (0.15)

 

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and NT and 40 of NG.

75

stabilities of soil aggregates 1 - 2 mm, for both CT and NT soils, Fig 12 and
Table 6. Increased (positive) delta MWD in the NT and CT treatments but not the
NG treatment, where aggregates were more stable initially, suggest that C and N
compounds were great enough in the NG treatment as to not respond to three
cycles of wetting and drying. Greater increases in the delta MWD values for the
C and N treatments of NT soils, Table 6, also veriﬁes that C and N compounds
may be lower in the tilled (ie., CT and NT) than the NG soils.

When C and N were added during each of the three wetting and drying
cycles, delta MWD values for aggregates, 2 - 4.75 mm across, from CT soils
decreased, Table 5. Additions of water caused soil aggregates to be less stable
than additions of C and N. Additions of C and N caused aggregates in the
rhizosphere to become more stable than those in bulk soil Fig. 11.

The percentages of water stable aggregates (ﬂSA), 2 - 4.75 mm across,
for bulk soils were signiﬁcantly greater for NT soils subjected to three cycles of
wetting and drying when water and C and N compounds were added by the
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signiﬁcant increase in the WSA of rhizosphere aggregates from CT soils.
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the Rhizos microtubes, declined signiﬁcantly. WSA for 2 - 4.75 mm in all regions
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79
water control among the water and C + N treatments during three cycles of

wetting and drying, Fig. 14. The absence of increased soil aggregation stabilities
among tillage treatments of these soils, agrees with Degens and Sparling (1995)
who reported that two wld cycles reduced aggregate stabilities by 25 - 35%.
These data suggest that movement of clay, cations, and other soil components,
associated with soil aggregation processes, did not seem to become effectively
organized in the tilled soils to a degree which enhances aggregate stability

during the ﬁrst three cycles of wetting and drying.

80

 

 

 

 

 

 

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8]
Nine wetting and drying (w/d) cycles

Experiment one of the two nine-wld cycles was completed without a water
control, which contained no Rhizos tubes nor TDR probes. The bulk densities of
the Kalamazoo loam soils for experiment one and two were 1.35 and 1.15 g/cc.
Another difference between the two 9-w/d cycle experiments was that the
“rhizospheres” of these experiments were deﬁned as soil regions 0-5 and 0-10
mm from the Rhizos microtubes. Nine wld cycles produced larger aggregates
(ie., 4.75 - 6.30 mm) than did three wld cycles for all three managed soils. W
of aggregates, 4.75 - 6.30 mm, were significantly increased by the nine cycles of
wetting and drying when C and N compounds were naturally higher, as in the NG
soils, or when C and N compounds were added to the rhizospheres by simulated
roots in CT, NT, and NG soils, Table 7. Root C and N stimulation of aggregate
stabilities appeared to diminish as aggregate sizes decreased from 4. 75 - 6.30
to 2 - 4.75 and 1 - 2 mm across, Tables 7, 8 and 9.

Experiment two of the 9-cycle wld treatment demonstrated drammatic and
signiﬁcant increases in the stabilities of aggregates, 4.75 - 6.30 mm across,
especially in the rhizospheres of soils when water was removed and either water
and/or C plus N compounds were injected into soils from CT management
programs, Fig. 15 and Table 7. C and N treatments also caused substantial, 5-
fold, increase in the stability of aggregates from rhizospheres of NT soils.
Greater, 2-fold, MWDs of aggregates in the rhizospheres of root water

treatments suggest that the withdrawal of water by roots may concentrate clay

82

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Soil management groups

simulated root C and N compounds and nine wetting and
n=3

9
drying cycles of a Kalamazoo loam soil subjected to 10 years of CT

and NT and 40 of NG.

Fig.15. Mean weight diameter for soil aggregates 4.75-6.30 mm across,
followin

86
and other ﬁne textured soil particles in regions surrounding plant root systems.

Additions of C and N compounds by root systems futher augment stabilization
processes of macroaggregates adjacent to roots. Although MWDs were greater
for all treatments of the NG soils, additions of C and N compounds still
demonstrated signiﬁcant increases in MWDs of rhizosphere soils, located within
10 mm of the root, Fig. 15 and Table 7. Similar trends of increased aggregate
stabilization by root-based C and N compounds continued among smaller soil
aggregates from tilled soils, Figs. 16 and 17. Withdrawal of water and injections
of C and N compounds by roots caused signiﬁcant, 120 to 130%, increases in
the MWDs of aggregates, 2-4.75 mm, taken from rhizosphere regions of CT
soils, Fig. 16. Similar trends for increased values in rhizosphere MWDs of CT
soils, over those from bulk soils of the same treatment, were also observed for
smaller, 1-2 mm aggregates, Fig. 17. MWDs were the greatest in the rhizosphere
and bulk soil regions of NG for all three aggregate sizes ranging from 1 - 6.30
mm, Figs. 15, 16, and 17. Values for the delta MWDs of treatments containing
simulated C and N exudations for aggregates 2 - 4.75 and 4.75 - 6.30 mm,
Tables 7, 8, and 9, suggest that macroaggregates are more responsive to root
modiﬁcations of soil aggregates than are smaller, 1 - 2 mm, aggregates.

Soil aggregate stabilities in this study appear to be greater in soils from
native grasslands (NC). This information is supported by Haynes and Swift
(1990). The injection or withdrawal of water and C plus N compounds from the

Rhizos SSS simulations of roots signiﬁcantly increased aggregate stabilities of

 

 

 

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Soil management groups

following simulated root C and N compoungs and nine wetting and
T and 40 of NG. n=3

drying cycles of a Kalamazoo loam soil subjected to 10 years of CT

Fig.16. Mean weight diameter (MWD) for soil ag regates 24.75 mm across,
and

 

 

   

 

 

 

 

 

 

    

   
 

 

 

 

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Soil management groups

Fig.17. Mean weight diameter for soil aggregates 1

- 2 mm across,
following simulated root C and N compounds and nine wetting and

drying cycles of a Kalamazoo loam soil subjected to 10 years of CT

and NT and 40 years of NG. n=3

89
CT and NT soils more than for N6 soils. Nine wetting and drying cycles

increased the stability of macroaggregates, 4.75 - 6.30 mm across, in the
rhizospheres of CT soils nearly 5-fold above the background water control while
MWD of the bulk soils approximately doubled. MWD of NT aggregates in the
rhizosphere increased more than 4-fold while aggregates in the bulk soils
increased 60%. Nine w/d cycles slightly decreased the MWD of smaller
macroaggregates, 1-2 mm across, for CT soils. These data suggest ways in
which water and solutes move in aggregated soils may depend on the mode of
saturation of the pore space which is made up of the micropore region within the
aggregates and the macropores surrounding them (Youan and Leeds-Harrison,
1990). The nine wld cycles of this study enhanced soil particle aggregation which
formed larger and stronger soil aggregates. Utomo and Dexter (1982) reported
that six wetting and drying cycles of a ﬁne sandy loam soil increased the stability
of remoulded aggregates by up to 4 times. Others have noted that soils become
completely structureless after 60 wld cycles (Jager and Bruins, 1975) suggesting
that the mode of wld cycling as well as the losses of ﬁne particles and associated
important cations associated with soil aggregation processes may cause

reductions in soil aggregation sizes and strengths of different macroaggregates.

90
Soil microbial activities

Dates of sampling

Soil microbial biomass estimations from three management practices on
three different dates varied considerably, Table 10. Generally, microbial biomass
values for CT and NT soils were lower than those for the native grassland. In
November 1996 the soil was frozen and the top 3 to 5 cm were removed during
sampling. This may be one of the reason the soil microbial biomass is low
compared to November 1995.
Three wetting and drying cycles

Three wetting and drying cyclesincreased the microbial biomass in the
rhizosphere soils of NT when root exudates were added. Increases of microbial
biomass were 15 % when root water was added and 51 % when root exudates
were added, Table 11. Root additions of water or C plus N had little affect on
microbial biomass values of the bulk soil. Neither root exudates nor additions or
withdrawals of water had any affects on the microbial biomass activities in N6
soils, Table 12.
Nine wetting and drying cycles

In contrast to bulk soils, additions of root C and N compounds signiﬁcantly
increased the microbial biomass values for rhizosphere soils from all
management practices tested in this study, Table 13. Nine wetting and drying
cycles of soils receiving simulated root exudates increased, the microbial

biomass values for rhizosphere soils over the control values of NG soils.

91

Table 10. Background microbial biomass from samples collected from O - 20 cm
in the three management practices in Kalamozoo loam soil subjected to
10 years CT, NT and at least 40 years of NG.

 

Microbial biomass

 

 

 

 

Date CT NT NG
. pg 0 g“l soil
November 1995 269 277 412
May 1996 275 296 357
(68) (81) (8)
November 1996 175 210 310
(22) (12) (16)

 

Values in parentheses are standard deviations of the means. n= 3.

92

 

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93

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94

 

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95
These data suggest that rhizosphere of 0 - 10 mm, certainly less than 20 mm,

were established by the Rhizos SSS microtubes of this study. Root exudates
increased microbial activities by 16 % in CT, 14 % in NT, and 69 % in N6 for
rhizosphere soils above control values, Table 14. Greater microbial biomass in
the rhizospheres of NG soils may be due to greater quantities of microorganisms
in this soil, Table 13. The negligible and negative affects of 9 wetting and drying
cycles, without additions of root C and N compounds, on microbial biomass
estimates of soil microbial activities, Table14, suggest that soil wetting and
drying of these soils does not liberate more C and N for mineralization and
contradicts the conclusions by Degens and Sparling (1995).

Decreases in the microbial biomass of bulk soils is expected after nine wetting
and drying cycles because most ions, including C and N are diluted with each
wld cycles resulting in greater competition for the limited C and N compounds.
As available C and N compounds in the soil are used up microorganism numbers
decline. Microbial biomass after nine wetting and drying cycles for the samples
collected on three different dates are variable with consistently higher treatment
responses observed for soils sampled on November 1995, Table 11, 13 and 14.
One of the reason is because the soil was frozen and the top 3 to 5 cm were
removed when samples were collected. Also microbes are more active when it
starts to be warm. Microbial biomass values, after 9 WM cycles, for samples
collected on November 1995, Table 15, had higher values than those collected

on November 1996, Table 13, because the surface 3 - 5 cm were removed due

96

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97

 

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98
to their frozen condition at the time of sampling. This suggests higher

concentration of microorganisms in the top 0 — 5 cm than in the 5 - 20 cm.

In conclusion, microbial biomass in rhizosphere soil increased with
additions of root exudates. The increase was signiﬁcant of the nine wetting and
drying cycles in CT, NT, and NG. However, there were no signiﬁcant differences
in bulk soil. In three wetting and drying cycles, only root exudates in rhizosphere
of NT shows a signiﬁcant difference. These differences show that few of the C
and N compounds added, actually migrated into the bulk soil. No signiﬁcant
increases in microbial biomass occurred during the 9 wld cycles, for the root
water controls. These data also contradict Van Gestel et al., (1990) who
suggested that greater amounts of organic C becomes available to
microorganisms as a result of the disruption of micro-aggregates or soil pores
during drying and re-rewetting. This phenomenon may have occurred in NG soil,
but not in historically tilled agroecosystems (eg., CT, NT). Changes in aggregate
stability in the rhizosphere have often been attributed to the presence of root
exudates (Oades, 1984; Reid et al., 1982; Turcheneck and Oades, 1978). Due to
its polysaccharidic nature, root exudates play a signiﬁcant role in the
‘cementation’ of soil particles (T urcheneck and Oades, 1978). It was shown to
bind with the surface of minerals present in natural or artiﬁcial soils (Guckert et
al., 1975; Tisdall and Oades, 1982). The use of intact root mucilage, collected
from maize plants , produced evidence of the adsorption of exudates on clay

minerals(Habib et al., 1990; Morel et al., 1987).

99
Summary

The results obtained during this experiment demonstrated the inﬂuence of
artiﬁcial roots, root exudates, wetting and drying modiﬁcations on soil aggregate
stability, previously subjected to different management practices. The presence
of artiﬁcial roots used to removed water from the soil during the wetting and
drying cycles has an impact on the reorganization of clay particles inside the
cyclinder. The presence of artiﬁcial roots with no C and N increased aggregate
stability by 2.3 times. Application of root C and N as exudates improved
aggregate stability by increasing the quantity of larger aggregates and their
mean weight diameter. Aggregate stability increased in the rhizosphere 4.9 times
greater when compared to the baseline. The number of wetting and drying cycles
affect aggregate stability. More stable aggregates were expected after three
wetting and drying cycles. Soil management practices signiﬁcantly affected the
formation and stabilization of soil aggregates when subjected to root C and N.
Larger and more stable aggregates were formed in treated NG soils. The
greatest improvements in soil structure were obtained through the application of
root exudates in NT, and CT. This means that aggregates in tilled plots are more
responsive to improvements when changes are made in plant and soil

management practices.

100

References

Buyanovsky, GA, and Wagner, G.H. 1997. Crop residue input to soil organic
matter on Sanbom ﬁeld. In Soil Organic Matter in Temperate
Agroecosystems. Long-Term Experiments in North America. Eds by Paul,
E.A., Elliot, E.T., Paustian, K., and Cole, C.V.

Dalton, F.N. 1992. Development of time-domain reﬂectometry for measuring soil
water contents and bulk soil electrical conductivity. In: Advances in
measurements of soil physical properties: Bringing theory into practice
pp 140-167. (SSSA Special Publication no. 30).

Dalton, F.N., Herkelrath, W.N., Rawlins, D.S., and Rhoades, JD. 1984. Time-
domain reﬂectometry: Simultaneous measurement of soil water content
and electrical conductivity with single probe. Science 244:989-990.

Emerson, W., and Grundy, G.M.F. 1954. The effect of rate of wetting on water
uptake and cohesion of soil crumbs. J.Afric.Sci.44:249-253.

Degens, BR, and Sparling, GP. 1995. Repeated wet-dry cycles do not
accelerate the mineralization of organic C involved in the macro-
aggregation of a sandy loam soil. Plant and Soil 175:197-203.

Emerson, W. 1977. Physical properties and structure. In Soil Factors in Crops
Production in a Semi-Arid Environment (eds J.S. Russell and EL.
Greacen). pp. 78-104. Queensland: University of Queensland Press.

Guckert, A., Chavanon, M., Mench, M., Morel, J.L., and Villemin, G. 1991. Root
exudation in Beta vulgaris: a comparison with Zea mays. Plant roots and
their environment. Eds McMichael, B.L., and Person, H. Elsevier. pp.
449-462.

Habib, l.., Chenu, C., Morel, J.M., and Guckert, A. 1990. Adsorption de
mucilages racinaires de mais sur des argiles homoioniques;
Consequences sur la micro-organisation des complexes forrnés. CR.
Acad. Sci. 310:1541-1546.

Haynes, R.J., and Swift, RS. 1990. Stability of soil aggregates in relation to
organic constituents and soil water content. J.Soil Sci. 41:73-83.

Hillel, D. 1982. Introduction to soil physics. Academic Press, Inc. San Diego,
California

Jager, G., and Bruis, EH. 1975. Effect of repeated drying at different
temperature on soil organic matter decomposition and characteristics, and
on the soil microﬂora. Soil Biol. Biochem. 7:153—159.

Jenkinson, D.S., and Powlson, D.S. 1976. The effects of biocidal treatments on
metabolism in soil-V. A method for measuring soil biomass. Soil Biology
and Biochemistry 82209-213.

Kemper, W.D., and Rosenau, RC. 1986. Aggregate stability and size
distribution. Methods of Soil Analysis, Part 1. Physical and Mineralogical
Methods-Agronomy Monograph No. 9 (2nd eds.) Am. Soc. Agon.
Madison WI.

Lynch, J.M., and Bragg, E. 1985. Microorganisms and soil stability. Advances in
Soil Science. Volume2. pp 133-170

101

McCalla, TM. 1945. Inﬂuence of micro-organisms and some organic substances
on soil structure. Soil Sci. 59:287-297.

McCalla, T.M., Haskins, F .A., and Frolik, E.F. 1957. Inﬂuence of various factors
on aggregation of Peorian loess by microorganisms. Soil Sci. 84:155-161

Morel, J.L., Andreux, F., Habib, L., and Guckert, A. 1987. Comparison of the
adsorption of maize root mucilage and polygalacturonic acid on
montmorillonite homoionic to divalent lead and cadmium. Biol. Fertil. Soils
5: 1 3-17.

Morel, J.L., Mench, M., and Guckert, A. 1986. Measurement of Pb2+ Cu2+ and
Cd” binding with mucilage exudates from maize (Zea mays L.) roots. Biol.
Fertil. Soils 2:29-34.

Oades, J.M. 1984. Soil organic matter and structural stability; mechanisms and
implication for management. Plant and Soil 76:319-337.

Paul E.A., and Clarck PE. 1996. Soil Microbiology and Biochemistry (2nd
Academic Press, Inc., New York.

Powlson, D.S., and Jenkinson, D.S. 1976. The effects of biocidal treatments on
metabolism in soil. ll. Gamma irradiation, autoclaving, air-drying and
fumigation. Soil Biol. Biochem. 8: 179-188.

Reid, J.B, Gross, M.J., and Robertson, PD. 1982. Relationship between the
decrease in soil stability effected by the growth of maize roots and
changes in organically bound iron and aluminium. J. Soil Sci. 33:397-410.

Rovira, AD, and Greacen, D.J. 1957. The effect of aggregate disruption on the
activity of microorganisms in the soil. Aust. J. Agron. Res. 82315-322.

Sissoko, F., and Smucker, A.J.M. 1997. Simulated root exudates enhanced soil
aggregation during wetting-drying cycles.(Agronomy J. submitted).

Sorensen, L.H. 1974. Rate of decomposition of organic matter in soil as
inﬂuenced by repeated air drying-rewetting and repeated additions of
organic material. Soil Biol.Biochem. 6:286-292.

Tisdall, J.M., and Oades, J.M. 1982. Organic matter and water-stable aggregates
in soils. J.Soil.Sci. 33:141-163.

Topp, G.C, Davis, J.L., and Annan, AP. 1982. Electromagnetic determination of
soil water content using TDR: l. Application to wetting fronts and steep
gradients. Soil Sci. Soc. Am. J. 46:672-678.

Turcheneck, L.W., and Oades, J.M. 1979. Fractionation of organo-mineral
complexes by sedimentation and density techniques. Geoderma.

21 :31 1343.

Utomo, W.H., and Dexter, A. R. 1982. Changes in soil aggregate water stability
induced by wetting and drying cycles in non-saturated soil. J. Soil Sci.
33:623-637.

Van Gestel, M., Ladd, J.N., and Amato, M. 1991. Carbon and nitrogen
mineralization from two soils of contrasting texture and micro-
aggregate stability: inﬂuence of sequential fumigation, drying and storage.
Soil Biol. Biochem. 23:313-322.

102

Youngs, E.G., and Leeds-Harrison, PB. 1990. Aspects of transport processes in
aggregated soils. J. Soil Sci. 41:665-675.

5.0 CONCLUSIONS

The interactions among plant roots and their exudates, soil water, soil
ions, and microbial biomass activities affect soil aggregate development,
function, and stability. Carbon and nitrogen compounds from plant roots
stimulate rhizosphere microorganisms which contribute to soil aggregation
processes. Frequent soil wetting and drying cycles appeared to contribute
directly to the positioning of soil particles. Interactions of root exudates, soil ion
and water contents, and augmented soil cohesion improved soil aggregate
stability. This experiment has demonstrated the inﬂuence of artiﬁcial roots, root
exudates, wetting and drying modiﬁcations on microbial biomass and soil
aggregate stability. The presence of artiﬁcial roots used to remove water from
the soil during the wetting and drying cycles has an impact on the reorganization
of clay particles and microbial activity inside the cyclinder. Application of root C
and N compounds which simulate exudates (composed of different organic
acids, carbohydrates, and amino acids) by Rhizos SSS microtubes improved
aggregate stability by increasing the number of microorganisms in the
rhizosphere soil, and the quantity of larger aggregates as well as their mean
weight diameters. Microbial biomass decreased when samples were collected far
from the Rhizos SSS microtubes. Aggregate stability increases in the
rhizospheres were 4 to 5 times greater when compared to the water controls for

the aggregate sizes 4.75 - 6.30. The number of wetting and drying cycles

103

104
affected the aggregate stability. The three wetting and drying cycle has no

signiﬁcant effect on the improvement of soil aggregate stability. However, the
nine wetting and drying cycles has a great impact on the formation and
stabilization of soil aggregates. The management history has a very signiﬁcant
affect on the formation and stabilization of soil aggregates. The addition of root
exudates accompanied by an adequate number of wetting and drying cycles
improved soil aggregates in no tilled (NT) and conventionally tilled (CT) more
than the native grassland (NG) which was already well aggregated.

One of the powerful components of soil aggregate formation is the soil
clay content. It’s orientation has a direct impact on soil aggregate formation. In
this study, it appeared that removal of water, by Rhizos vacuum extractions
accumulated some of the dispersed clay on soil aggregates located in the
rhizosphere regions adjacent to the simulated roots. Additional experiments need
to be conducted to better understand the mechanisms associated with clay
movement around the Rhizos SSS, and the interactions of bacteria in the
rhizosphere soil which combined with other C and N compounds act as a

cementing agents which form stable soil aggregate formation.

6.0 References

Allison, F.E. 1973. Soil organic matter and its role in crop production. Eds In
Elsevier scientiﬁc publishing company.

Anderson, J.P.E. and Domsch, K.H. 1978. A physiological method for the
quantitative measurement of microbial biomass in soils. Soil Biology and
Biochemistry 10:215-221 .

Anderson, J.P.E. and Domsch, K.H. 1989. Ratios of microbial biomass carbon to
total organic carbon in arable soils. Soil Biol. Biochem. 10:215-221.

Angers, D.A., And Mehuys, GR. 1989. Effects of cropping on carbohydrates
content and water-stable aggregation of a clay soil. Can. J. Soil Sci.
69:373-380.

Arnold, PW. 1978. Surface-electrolyte interactions. In The Chemistry of Soil
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