Soil Bid. Biochem. Vol. 17, No. 4, pp. 565-569, 1985
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Copyright 0 1985 Pergamon Press Ltd
TURNOVER OF ROOT-DERIVED MATERIAL AND
RELATED MICROBIAL BIOMASS FORMATION
IN SOILS OF DIFFERENT TEXTURE
R. MERCKX, A. DEN HARTOG and J. A. VAN VEEN
Research
Institute
ITAL,
P.O. Box 48, 6700 AA Wageningen,
The Netherlands
(Accepted 20 December 1984)
Summary-Wheat
plants were grown on two soils of different texture, a sandy soil and a silty clay loam,
in an atmosphere
containing
“C0,. The 14C and total C content of the shoots, roots, soil rhizosphere CO,
and soil microbial
biomass were measured
21, 28, 35 and 42 days after germination.
There was a
pronounced
effect of soil texture on the turnover
of root-derived
C through the microbial
biomass.
Turnover was relatively fast and at a constant rate in the sandy soil but slowed down in the clay soil,
following an initial high assimilation of root products into the microbial biomass.
Four percent of the total fixed 14C was retained in the clay loam after 6 weeks compared with a
corresponding value of 1.2:4for the sandy soil. The proportion of fixed 14Crecovered as rhizosphere CO,
at each of the sampling times was relatively constant for the sandy soil (ca 19%) but decreased from 17%
at day 28 to 11% at day 42 in the clay soil. The proportion of total fixed ‘%I in the soil biomass as measured
by a fumigation
technique increased to a maximum value of 20% after 6 weeks in the sandy
decreased in the clay soil from 86% at day 21 to 26% after 42 days plant growth.
INTRODUCTION
the related
turnover
of C through
the microbial
biomass. To do this, we measured the distribution
of
14C within shoots, roots, soil, rhizosphere
CO, and
soil microbial biomass during the growth of wheat in
an atmosphere
containing 14C0, of constant specific
activity. The wheat was grown on two soils differing
in clay content.
Plant roots are essential sources of energy and carbon
for heterotrophic
soil microorganisms.
Earlier studies
on root exudates from plants growing in nutrient
solutions showed the great diversity of organic materials that were released from roots and, potentially,
could serve as substrate for microbes (Hale et al.,
1978). The use of ‘%02 in closed growth chambers
allowed for a more realistic measurement
of the
transfer
of photosynthesis
products
to roots and
subsequently
to the soil (Martin, 1975, 1977; Barber
and Martin,
1976; Helal and Sauerbeck,
1983). A
large proportion
of the photosynthesis-products,
translocated
to roots, are mineralized to CO,, either
by root respiration or by microbial degradation.
The
latter process results in a marked proliferation
of the
rhizosphere microflora, as predicted by the model of
Newman and Watson (1977).
Microbial
biomass formed through
the decomposition of labelled compounds
can be estimated by
the chloroform
fumigation incubation method (Ladd
et al., 1977; Amato and Ladd, 1980; Kassim et al.,
1981; Cerri and Jenkinson,
1981). Such studies
confirmed the key role of microbial biomass in the
turnover
of elements such as C and N in soil.
Although the biomass C generally constituted
only
l-5% of the soil C, 20 to even 60% of residual organic
14C and “N could be found in the microbial biomass,
several weeks after addition
of labelled organics
(Kassim et al., 1981; van Veen et al., 1984).
Microbial
biomass formation
is influenced by a
number of variables such as temperature,
moisture
and the presence of clay minerals. More microbial
biomass is formed as clay contents increase; likewise
more organic matter is stabilized (Martin et al., 1976;
Ladd et al., 1981).
Our objectives were to assess the influence of soil
texture on the production
of root-derived
C and on
S.BB. 17,4---L
soil but
MATERIALS AND METHODS
Soils
Two soils were collected from the Ap-horizon
of
arable land; a sandy soil at Horst and a silty clay
loam at Lelystad, in the newly reclaimed Lake Yssel
polders (Table 1). Soils were stored at constant
field moisture content before use in the incubation
experiments.
Prior to filling columns
(dia 90 mm; height
250 mm), soils were sieved (< 5 mm) and the moisture
level adjusted to 60% of the field capacity.
Plant growth
Soaked seeds of summer wheat (Triticum aestivum
var. Sicco) were transferred to soil columns (one per
column). The columns contained approx. 1500 g of
Table 1. Characteristics of the soils from Horst (NL) and Lelystad
(NL)
pH H,O
organic C content (%)
inorganic C content (%)
H,O content at f.c.
(% w/w)
% clay <2flm
‘% silt 2-50pm
% sand > 50 fi m
565
Horst
(sandy soil)
Lelystad
(silty clay loam)
5.6
1.0
0
19.1
7.8
1.3
1.2
33.3
2
2
96
31
50
13
566
R.
MERCKX el al.
either soil, packed to a final density of 1.2g cm p3 by
gentle vibration. The soil columns were closed with
a lid, with openings
for the stem and for water
addition.
After growth for 10 days in a normal atmosphere,
the opening around the plant shoots was sealed with
a mixture of lanolin: paraffin (19: I), warmed to 45°C.
The seals were tested against a positive air pressure
(80 kPa) before transferring
the columns
to the
growth chambers.
The growth chambers
of the Experimental
SoilPlant Atmosphere
System (ESPAS) allow for a separate control of the climatic conditions of shoots and
roots.
The atmospheric
14C02 concentration
was kept
constant at 330 + 5 cm’ mm3, with a specific activity
of 5-6 kBqmg
’ C throughout
the growth period.
During
the day period
(16 h, 20 klx: P.A.R.
54 W rn-‘) the temperatures
in the shoot and root
compartments
were 21°C and 20°C respectively.
A
30 min period with a light intensity of 10 klx (P.A.R.
29 W m-*) simulated twilight. During the dark period
the temperatures
of the shoot and root compartments
were 16” and 18°C respectively. The relative humidity
was constant at 6&70:,/,. Soil moisture was adjusted
daily by top irrigation.
Carbon dioxide produced in the soil column was
trapped in 300 ml of a 0.5 M NaOH solution by
flushing the columns
for 15 min every 6 h with
COz-free air. Total CO, was determined by titration
of excess NaOH with 0.5 M HCI after addition of
BaCl, to precipitate HCO; and CO,‘- ions; 14C0, by
liquid scintillation counting using Instagel (Packard).
At each sampling
(3, 4, 5 and 6 weeks after
germination)
three columns were taken out of the
growth chamber. Shoots were clipped at soil level and
roots and soil were separated by gently shaking the
root--soil core. Root fragments were removed from
the soil that fell off by handpicking.
Root material
was washed with tapwater to remove adhering soil
Table 2.
RESULTS
Growth
of wheat on the silty clay loam was
considerably slower than on the sandy soil (Table 2).
It seemed that the plants on the Lelystad soil were
about one week behind the plants on the Horst soil,
since plant growth rate in the Horst soil over the
21-35-day
period
was similar to that over the
28-42-day period in the Lelystad soil. Although the
specific activities of shoots of the plants grown on
both soils equalled the specific activity of the 14C0,
atmosphere,
the specific activity of the roots and that
of the 14C0, respired in the root--soil compartment
were lower, in particular with plants grown on the
silty clay loam (Table 2).
The pattern of distribution
of ‘“C within plant and
soil components
was similar in both soils (Table 3).
About 5(r60”/, of the fixed 14C was in the shoots, and
the remaining “C-photosynthates
were transferred to
the below-ground
parts. A significant proportion
(up
Dry weight and ‘%-activity of plant and soil component! 3 after growth of wheat plants m a “C01 atmosphere (specific activity
5%6kBqnlgg’
C) on (A) a sandy soil and (B) a silty clay loam. Values are means of measurements on three plants
Time” (days after germination)
28
35
21
Shoots
Roots
particles.
Dry weights of shoots and roots were
determined
after drying at 70 ‘C for 24 h. The dry
material was ground and homogenized
for total C
and 14C content determination,
using a modified
wet-combustion
procedure
(Dalal,
1979). Plant
(30 mg) or soil (1 g) mat.erial was digested in 5 ml or
a solution of 5.0 g K2CrZ07 in 100 ml of a mixture of
concentrated
H,SO, and H3POd (3:2 v:v) at 120 C
and 105 kPa for 1 h. The CO2 evolved was trapped in
10 ml 0.5 M NaOH. After correction for evaporatiorl
Iosses, total CO, and 14C0, concentrations
in the
NaOH solution were determined as described abo\*c.
Total biomass-C and biomass-“C
were determined
using the chloroform
fLlmigation-incubation
method
(Jenkinson and Powlson. 1976b). We used the CO,
evolved from the fumigated sample during the period
IO-20 days after fumigation as control, as suggested
by Chaussod and Nicolardot
(1982).
A /&-factor of 0.45 was used to convert net CO,-C
production
to biomass-C
(Jenkinaon
and Ladd.
1981).
~~~
~~~~ .._~_._. --
Shoots
Roots
Soil-.root respiration
Soil residue
813
952
..--.
853
463
196
40
-...-- ..-.-
Activity
{3.2)
(1.8)
(1.3)
(<O.Ol)
Shoots
Roots
75
ND”
Shoots
Roots
Soil-root respiration
Soil residue
Activity
45 (2.6)
ND”
12 (0.4)
7 (c10.01)
(A) Horst sandy soil
Dry weight (mg)
2250
2498
-~~~“. ~~~ ~~
(kBq) and specific activity
4102 (4.7)
2301 (3.6)
1616 (3.7)
393 (0.03)
-
(kBq) and specific activity
677 (4.4)
260 (1.5)
204 (1.4)
58 (<O.Ol)
“Prior to growth in the 14C02 atmosphere plants were grown in a normal
bRoots could not be separated from soil at this harvest.
I I.h5i!
7057
7341
10.59 1
(kBq mg ~’ C) (in parentheses)
13.479 (5.8)
7643 (4.4)
4940 (4.51
42 1 (0.03)
(B) Lelystad silty clay loam
Dry weight (mg)
565
3x0
(kBq mg‘
atmosphere
47
1640
2327
24,029 (5.5)
14.377 (4.5,
044 1 (4.X)
SX? (0.04,
J 540
7620
’ C) (in parentheses)
2853
1501
858
265
(5.6)
(2.9)
(3.2)
(0.02)
for IO days.
10.331
4295
1993
684
(6.1)
(3.5)
(3.2)
(0.05)
Root
Table 3. Distribution
of ‘%Z amongst plant and soil components after growth of wheat in a 14C0,
atmosphere on (A) a sandy soil, and (B) a silty clay loam
Time” (days after germination)
28
35
21
Shoots
Roots
Soil-root
Soil
561
material and microbial biomass formation in soil
respiration
42
(A) Horst sandy soil
Distribution
(% of total fixed-W)
48.8
50.9
49.6
21.3
28.9
29.1
‘1:;
1;:;
19.5
15.2h
23.9b
20.2b
1.2
55.0
29.8
12.6
2,6
20.7b
(B) Lelystad silty clay loam
Shoots
Roots
Soil-root
Soil
respiration
Distribution (“/:,of total fixed-“‘C)
56
52
60
22
21
25
NDC
ND’
ND’
ND’
1:
22b
“Prior to growth in the ‘%ZO, atmosphere plants were grown in a normal atmosphere
‘Sum of soil-root respiration and soil.
‘Root biomass and ‘%Z-activity could not be determiwd
at this harvest.
to 40%) of the latter compounds
was respired, either
by root metabolism
or by microbes.
Of the total
amount of 14C fixed, a larger proportion
was respired
below-ground
in the sandy soil than in the silty clay
loam, but the proportion
which remained in the soil
was considerably higher in the silty clay loam than in
the sandy soil.
Large differences were observed between soils in
the turnover
of Y-labelled
products
through the
microbial biomass (Fig. 1). At the beginning of the
experiment
the microbial biomasses were 980 pg C
gg’ and 660 pg C gg’ for the sandy soil and the silty
clay loam respectively,
as measured by the chloroform fumigation
incubation
method. The absolute
amount of 14C in the biomass of the two soils as
measured by the chloroform
fumigation-incubation
method steadily increased during plant growth, but
to a larger extent in the silty clay loam than in the
sandy soil, in spite of the much larger quantities of
the roots in the latter soil (Table 2 A and B). Very
different proportions
of residual 14C were incorpo-
for 10 days.
rated into microbial biomass in the two soils. In the
silty clay loam, biomass-‘%
as a proportion
of soil14C decreased from a remarkably high 86% at day 21
to 26% at day 42. In the sandy soil biomass-‘4C
increased from 8% of the residual 14C at day 21 to
20% at day 42. When calculated on the basis of the
specific activity of roots and microbial biomass, 14Clabelled biomass comprised approx. 4% of the total
microbial biomass in the silty clay loam and approx.
6% in the sandy soil at the end of the experiment.
DISCUSSION
The distribution
of 14C-carbon between shoots,
roots and soil-root
components
(including
rhizosphere 14C02) in our experiments
with wheat is
roughly similar to the distribution
patterns observed
by Helal and Sauerbeck (1983) for maize and Martin
(1975, 1977) for wheat, ryegrass and clover. Thus,
this study confirms observations
that considerable
amounts of root-material
are lost from actively grow-
kBq “?
o-o
A-A
250.
1
1
E” 150.
.o
0
0
.a
e IOO.I!
E
.,o
50-
21
28
Fig. 1. Soil microbial 14C content during 6 weeks growth
on a silty clay loam
from Lelystad
35
42 days
of wheat on a sandy soil from Horst (A) and
(0); and percentages
of soil ‘T in microbial biomass in either soil
(x and l respectively).
568
R.
MERCKX rt ul
ing plants, supplying the soil organisms with energyrich substrates. However, we also observed a marked
difference in the production
and the fate of rootderived materials between soils of different texture. In
the silty clay loam (a) a greater proportion
of fixed
C was retained in soil at harvest than in the sandy
soil; (b) larger amounts of soil lJC were incorporated
in the microbial biomass than in the sandy soil; (c) the
proportion
of soil ‘% in the microbial
biomass
decreases with time, while it increased in the sandy
soil and (d) the production
of rhizosphere
‘“CO? as a
percentage
of total fixed ‘% decreased
with time,
while it was constant in the sandy soil.
A relative decrease in soil-root
respiration
in the
silty clay loam soil could be caused by decreased root
respiration or by decreased microbial respiration.
In
our experiment
these components
could not be distinguished. The decline of the shoot-to-root
ratios at
the end of the growth period of the plants grown in
the clay soil might involve a decrease
in rootrespiration,
following the decline in carbohydrate
supply from shoot to root.
Lower nutrient
concentrations
in the silty clay
loam may also have a negative influence on root
respiration (Veen, 1981). In both soils, but to a larger
extent in the silty clay loam, the specific activities of
the roots and of the soillroot
respiration
are lower
than those of the shoots. The presence of unlabelled
roots that developed during the 10 days before the
plants were put in the 14C0, growth room may
partially
account
for this. Equilibration
between
14C0, evolved by the root-microbial
biomass system
and HCO,/CO:present in the calcareous silty clay
loam soil would also lower the specific activity of the
evolved CO, (Jacoby and Laties, 1971).
Nevertheless, we propose that the overall soil-root
respiration
decline in the silty clay loam must be
ascribed to a decrease in microbial respiration.
This
hypothesis is based on the observations
that a higher
proportion of root-originating
products is fixed in the
clay-rich soil and that a decreasing proportion
of soil
14C is incorporated
in soil microorganisms.
Both
point to an accumulation
of root-derived
material,
presumably
due to a reduction
in the microbial
degradation.
We consider the high content of clay minerals the
principal factor restricting microbial degradation
of
root-derived
material in the silty clay loam. Other
factors,
such as pH and the quantitative
and
qualitative
differences
in microbial population
are
considered to be of secondary importance.
Clay minerals may act as an adsorption
sink for
organic materials in soils, thus reducing their decomposition rate (Marshman
and Marshall,
1981). The
finer texture of clay soils may also hamper
fast
turnover of organic products (Jenkinson and Rayner,
1977; Elliott et ul., 1980; van Veen and Paul, 1981) by
entrapment
in aggregates or by other mechanisms
that restrict biological degradation.
However, stimulatory effects of clay addition
have also been reported. Clay minerals can buffer changes in pH and
thus allow breakdown to continue under favourable
conditions
(Stotzky
and Rem, 1966; Kunc and
Stotzky, 1974).
Immobilization
of toxic metabolites
or inhibitors
by adsorption
on clays was suggested (Martin et al.,
1976) to be more important
than pH stabilization.
Furthermore,
clay minerals have also been claimed to
exert a protective
action on bacterial cells by the
formation of so called “clay-envelopes”
around them
(Marshall, 1971; Lahav, 1962; Kilbertus et al., 1977).
Such clay coatings
are effective barriers
against
predation (Marshall,
1975). Different processes may
dominate in different cases.
We suggest that in our study the soil microflora
initially assimilated root-derived
organics to a larger
extent in the silty clay loam than in the sandy soil.
However, as the plants developed, the composition
of
the root released products gradually changed from
soluble exudates to less-soluble polymers (mucilage,
sloughed-off cells), which were less accessible to the
microbial biomass in the silty clay loam than in the
sandy soil. A shift from soluble to less soluble
(non-diffusible)
root-released
products is of greater
importance
in the silty clay loam than in the sandy
soil because diffusion
of nutrients,
necessary
for
bacterial degradation
of compounds
lvhich do not
diffuse themselves,
is more restricted
than in the
sandy soil. Together
with the slower turnover
of
microbial
cells in the silty clay loam, this could
explain the high, but decreasing proportion
of soil14C in the microbial biomass of the silty clay loam as
compared to the lower, but increasing proportion
of
‘% in the microbial biomass of the sandy soil.
Soil microbial biomass estimates by the fumigation
method should, however, be interpreted
with great
care when fresh substrate has recently been added
(Jenkinson
and Powlson.
1976a, b; Voroney
and
Paul, 1984). When using the original chloroform
fumigation
incubation
method
of Jenkinson
and
Powlson (1976b) it is highly probable that native soil
and recolonizing
soil populations
decompose
the
fresh material at different rates in unfumigated
and
fumigated samples. It is common knowledge that in
the presence of fresh substrate the CO? evolution rate
of the control often exceeds the flush of CO2 from the
fumigated samles, which will lead to unacceptable
negative biomass values. This also happened in the
present experiments.
The difficulty in defining an
appropriate
control arises because of the different
(micro)biological
conditions
that occur in the fumigated and the non-fumigated
soil (Chaussod
and
Nicolardot.
1982: Voroney and Paul. 1984). Some
authors omit the use of a control to overcome these
problems (Voroney and Paul. 1984). We considered
the omission of a control as incorrect in our experiments since the decomposition
of the continuouslyproduced fresh root material should not be neglected.
We therefore used the CO? evolved from the fumigated soil sample in the I&20-day
period as the
control, as suggested by Chaussod and Nicolardot
(1982). This version of the chloroform
fumigation
incubation
method has the, at least theoretical,
advantage, that the control CO, production comes from
the same (and microbiologically
comparable)
sample
as the CO? evolved after fumigation.
We are aware
that when continuous additions of fresh root material
occur, the quantity of material decomposed
in the
fumigated
soil during the O&IO days incubation
period may not be the same as during the lO_20-day
period and that this difference could well affect the
“C-biomass
determinations.
For these reasons we
Root
material and microbial biomass formation in soil
569
preferrred this version of the chloroform
fumigation
method and found that it gave meaningful
results
which otherwise could not be obtained.
We adopted a kc-factor of 0.45 for the calculation
of total biomass-C and biomass-‘% of both soils. The
almost complete absence of fungi (J. A. van Veen,
unpublished
results) in the silty clay loam may,
however, necessitate the use of a smaller kc-factor. A
kc-value of 0.33 was suggested
by Anderson
and
Domsch to be more appropriate
for measurement
of
bacterial biomass (Anderson
and Domsch,
1978).
However, the same kc-value was used throughout this
study, because values for the individual soils were not
avaiiable.
Until recently, k, was assumed to be the same in
soils of different texture. However, the results of this
and other studies on the effect of soil texture on
microbial biomass turnover processes (van Veen et
al., 1984) raises questions as to the constancy of k,
and k, in different soils.
for measuring soil biomass. Soil Biology & Biochemistry
8, 209-213.
Jenkinson D. S. and Rayner J. H. (1977) The turnover of
soil organic matter in some of the Rothamsted classical
experiments. Soil Science 123, 298-305.
Jenkinson D. S. and Ladd J. N. (1981) Microbial biomass
in soil-measurement
and turnover. In Soil Biochemistry,
Vol. 5 (E. A. Paul and J. N. Ladd, Eds), pp. 415471.
Dekker, New York.
Kassim G., Martin J. P. and Haider K. (1981) Incorporation of a wide variety of organic substrate carbons into
Acknowledgements--We thank Mr H. Roelofsen and Mr W.
van Lienden for their technical assistance and Dr S. C.
van de Geijn (ITAL, Wageningen) and Dr J. K. Martin
(CSIRO, Glen Osmond, Australia) for their critical comments and suggestions.
derived from the soil biomass. In Soil Organic Matter
Studies, Proceedings of the IAEA/FAO/SSF
Symposium,
Braunschweig, 1976, Vol. 1, pp. 301-310.
Ladd J. N., Oades J. M. and Amato M. (1981) Microbial
biomass formed from ‘“C, ‘5N-labelled plant material
decomposing
in soils in the field. Soil Biology & Biochemistry 13, 119-126.
Lahav N. (1962) Adsorption
of sodium bentonite particles
on Bacillus suhtilis. Plant and Soil 17, 191-208.
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