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V. Biomass and Bioactivity Measurements
FRANCIS E. CLARK A N D ELDOR A. PAUL
per gram of soil amounts to 450 g live weight per m2 to a depth of 15 cm.
Taking the water content of the bacterial cell as 80% (Camp, 1963), the
live-weight estimate can be restated on a dry weight basis as 90 g/m2to a
depth of 15 cm.
Individual soils of course vary greatly in their microbial content. Published estimates of live-weight bacterial biomass in soil are usually expressed as a range of values. Alexander ( 196 1) has stated this range to be
33-330 g/m2/15 cm; Jensen (1963), 100-1OOOg; Russell (1950), 170-390
g; Krasil’nikov ( 1 944),67-720 g; Latter and Cragg ( 1967), 2 1 - 135 g; and
Stockli (1956), 160-380 g. Averaging the above estimates gives a value
of 270 g live weight, or 54 g dry weight, for the bacterial biomass per m2 to
a depth of 15 cm.
In the older literature concerning estimates of the fungal biomass in
soil, the fungi were commonly stated to possess a biomass ranging from
equivalence to twice that of the soil bacteria. Recent measurements
(Stockli, 1956; Jackson, 1965, Latter et al., 1967) of hyphal lengths in
soil by direct microscopy range from 100 to 2000 mlg. The mean value
obtained by Jackson (1965) for seven soil samples was very close to 120
m of hyphal length per gram of soil. Transposing this measurement to
biomass, using an average hyphal diameter of 3 p , gives a value of 38 gl
m2/15 cm, or somewhat less than the 54 g estimate derived above for the
bacterial biomass. It is interesting to note that Latter and Cragg ( 1 967)
found the biomass of fungi in a Juncus community to be approximately
two-fifths that of the bacterial biomass.
Data collected by Babiuk and Paul (1970) and Parkinson (personal
communication) in their studies on the magnitude of the microbial biomass in grassland at the Matador site in Saskatchewan are shown in
Table XIV. Their data were calculated on the basis that the average size
of the bacteria and actinomycetes was 0.6 by 1 p, and the average diameter of the fungi, 2.5 p. The bacterial diameter of 0.6 p, determined by
microscopic examination using fluorescein isothiocyanate, may be somewhat low, but even the use of 0.8 p as the diameter would still indicate
that the fungal biomass was roughly twice that of the bacteria, if viewed
on a mean seasonal basis. The data indicate that the ratio between biomass estimates of the bacteria plus actinomycetes and that of the fungi
at the 0-10 cm depth is similar to the ratio between the two groups at the
0-30 cm depth. There did not appear to be any positive correlation between the changes in biomass of bacteria and fungi during the season.
Whereas the bacteria approached their highest value in May, the fungi
were at their lowest. The biomass values tabulated are perhaps the best
current estimates for bacteria and fungi in grassland soil, and accordingly
THE MICROFLORA OF GRASSLAND
the averages, namely, 57 g/m2/30cm for the bacteria and actinomycetes,
and 138 g for the fungi, are combined to give 200 g as the microfloral
biomass in grassland. This value is used for some calculations in the
Biomass of Fungi and of Bacteria and Actinomycetes in
Matador Grassland Soil"
Bacteria plus actinomycetes'
Values are expressed as grams per square meter, dry weight.
"D. Parkinson (personal communication).
Babiuk and Paul ( 1970).
B. BIOMASSI N RELATION
The total microbial biomass in soil at any given time does not necessarily reflect microbial activity. An insight into the difference between
these two can be obtained by looking at microbial respiration rates or
by calculating energy requirements of microbial cells. In the laboratory,
actively multiplying bacteria are capable of producing many times their
dry weight of COz during 24 hours, whereas in stationary populations,
the production of CO, during 24 hours is approximately the equivalent
of dry cell weight. The discussion just above indicates a microbial biomass of 200 g/m2/30 cm. This is 35-70 times as much as the weight of
COz usually respired from field soils daily (Domsch, 1962; Monteith et
al., 1964; Krzysch, 1965; Tamm and Krzysch, 1965). The respiration of
the soil fauna and plant roots must also be taken into account for they are
in part responsible for the daily COZproduction from the soil. It is easily
apparent that great discrepancy exists between field measurements of
CO, production and that calculated for the microbial biomass using microbial respiratory rates determined in the laboratory.
Similarly large discrepancies are encountered in other calculations.
Pirt ( 1965) and Veldkamp (1 967) utilized total cell weight, and Oginsky
FRANCIS F . CLARK A N D ELDOR A. PAUL
and Umbreit ( I 9 5 9 , total cell nitrogen, as a basis for expressing the
maintenance energy needed by the living cell. The latter workers have
stated that bacteria require per gram 50 kcal/day of cell nitrogen to maintain their life functions. Accordingly, 60 g of bacteria, containing 6 g of
cell nitrogen, would require 300 kcal of energy daily. The inclusion of
140 g of fungal biomass would raise the requirement to approximately
1000 kcal/day. The net productivity in Matador plant communities rarely
exceeds 5000 kcal/m2/yr.This falls far short of meeting the annual maintenance energy requirement for 200 g of microbial biomass. Similarly,
the process of microbial cell production can be evaluated in terms of
available substrate. Assuming aerobic utilization of substrate, each gram
of dry cell weight synthesized requires the equivalent of 1.4 g of dextrose
(Camp, 1963). Therefore, 200 g of microbial biomass would require 280
g of dextrose, or roughly 1100 kcal of energy, for each generation of
microbial cells produced. Under this requirement, the soil microflora at
the Matador site, as well as that in soil generally, could reproduce only
a very few times per year.
OF BIOACTIVITYI N SOIL
Such calculations as the above, together with dissatisfaction with the
usually large and variable discrepancies between the dilution and direct
counts of soil microorganisms, have led many workers to turn to measurements of soil metabolism as a means of studying the soil microflora. Such
studies are directed primarily at the bioactivity in a given soil and are not
concerned with identifying the individual species, genera, or groups of
bacteria and hngi involved in that activity.
Respiration is one of the earliest and still one of the most frequently
used indices of microbial activity in soil (Fred and Waksman, 1928; Katznelson and Stevenson, 1956; Stotzky, 1965). Measurements of oxygen
consumption or of carbon dioxide production have been used extensively
in studying microbial activity and rates of decomposition of organic
materials in soil. The value of the respiratory approach is well recognized,
and hrther discussion of it will not be undertaken here.
Recently, the recognition of the presence of enzymes in soil and development of methods for their measurement have led to their use for
studying bioactivity in soil. Several recent studies suggest that this approach will be informative (Hofmann, 1963; Hofmann and Hoffman,
1966; Casida et al., 1964; Skujins, 1967: M. G. Myers and McGarity,
1968). Ross (1965) found that significant correlations existed between
sucrase and lipase activity in New Zealand soils and such soil properties
as organic carbon, soil moisture, and air temperature. Ross ( 1 966, 1968)
THE MICROFLORA OF GRASSLAND
has also reported some specific vegetation effects on soil enzyme activity.
Sucrase and lipase activities in soils under forests differed from those
in soils under pastures. In comparisons of pasture plants, it was found
that the ratio of sucrase to lipase was always significantly higher under
Dactylis glomerata and Bromus spp. than under Trifoliurn pratense and
T . repens.
The biological properties and enzyme activities of a range of Russian
grassland soils are shown in Table XV. Mishustin and co-workers (1968)
believed that a direct correlation exists between the activity of invertases
and soil respiration while activity of the oxidases is correlated to the dynamics of nitrate. Their data (Table XV) show that soils high in organic
carbon contain comparatively low numbers of microorganisms per gram
of organic matter. This is not surprising. Soil organic matter is known to
consist of an active and a passive fraction, as will be discussed shortly.
Soils low in humus but having relatively high primary productivity can be
expected to have a greater microbial population per gram of carbon than
soil in which stable organic matter is accumulating.
A direct relation between numbers of soil organisms, activities of enzymes, and soil organic carbon content should not be expected. These
relationships are determined by a number of factors such as persistence
and inactivation of enzymes and production of enzymes by microorganisms and plant roots. Because enzymes are adsorbed and stabilized in a
soil system, certain of them may have to be looked on as potential, or as
representing past activity, just as is the case for numbers of microorganisms determined by the direct count.
Some recent and as yet largely untried methods involving enzyme measurements as a means of estimating either gross microbial activity of the
activity of individual microbial units in soil include measurement of ATP
activity by means of a luciferin-luciferase enzyme system (Doxtader,
1969) and the use of esterified dyes such as fluorescein diacetate or dibutyrate in direct microscopy (Rotman and Papermaster, 1966). In the
latter procedure, fluorescence occurs only after an active esterase has
removed the acetate or butyrate side groups.
Bioactivity studies to date show the necessity for further definitive
work in this field. There are a number of variables interacting to affect
microbial numbers and activity in the field that are difficult to separate
and that make it difficult to determine which factor or combination of
factors is controlling activity. A true understanding of what happens in a
soil system probably will come about only by use of extensive field experiments in which the environmental parameters on all aspects of an ecosystem are carefully measured. Such attempts have been undertaken in
TABLE X V
Enzyme Activity and Riogenic Nature of Organic Matter in Various Soils Typical of Horizontal Belts"
organisms per gram
of organic matter
Mishustin et al. ( 1 968).
"he activity of invertase is expressed in milligrams of glucose per 1 g of soil (during 24 hours); of amylase, in milligrams of maltose per 1
g of soil (during 24 hours); of dehydrogenase, in milligrams of triphenylformasan per 10 g of soil (during 28 hours); of polyphenoloxidase, in 6
mg purpurogalin per 100 g of soil (during 30 minutes); and the activity of catalase, in 0 2 (in milliliters) per 1 g of soil (during 1 minute).
THE MICROFLORA OF GRASSLAND
the laboratory. Some examples are: growth and activity measurements
of soil organisms in microbead cultures (Parr et al., 1967); continuous
culture investigations (Macura, 1966, 1968); and batch, phase, or dialysis cultures of microorganisms (Kurtz et al., 1969). The use of model
systems for analyzing controlling factors and their effects also should
VI. The Humic Component of Grassland Soil
Organic compounds within the soil can be divided into the following
rather indistinct groups: (a) living plant roots, (b) the microflora and fauna
biomass, (c) decaying plant and animal tissues, (d) low molecular weight
compounds found either in the soil solution or adsorbed on particle surfaces, and (e) the humic component. Collectively, the first four groups
constitute the nonhumic materials. They accumulate in arable grassland and mineralize relatively rapidly in soil given cultivation or incubated
in the laboratory (Grant, 1967; Greenland and Ford, 1964; Heard, 1965).
The first three groups are particulate and can in large part be separated
from soil by screening or densimetric fractionation techniques. The nonhumic materials constitute 15-25% of the organic carbon in A horizons
of cultivated soil and a larger proportion in grasslands and forests.
OF THE HUMICCOMPONENT
The dark-colored, amorphous, pol ydispersed and highly polymerized
materials, commonly referred to as humus, comprise 50-85% of the soil
carbon and a slightly higher percentage of the soil nitrogen. These materials show a great deal of similarity even when isolated from widely
different locations (Kononova, 1966; Hansen and Schnitzer, 1966; Hodgson et al., 1968). They cannot be expected to be uniform in chemical
nature, inasmuch as the physical and chemical characteristics of the
environment at the time of formation significantly affect their structure.
Dubach and Mehta (1963) have remarked that probably no two molecules
of humic acid are exactly the same.
The term heteropolycondensate best describes the series of related
compounds in which aromatic structural units are linked by covalent
bonds such as ether linkages, -N H -, - S -, and -CH2-bonds. Hydrogen and cation bridge bonding also play a dominant role in the structure
of humic materials. Simonart et al. (1967) reported that high molecular
weight, amino-rich materials were adsorbed to an aromatic core and could
be removed by phenol treatment. Work of Biederbeck and Paul (1968)
also indicates the adsorption of high molecular weight proteinaceous
materials to an aromatic core.
FRANCIS E. CLARK A N D ELDOR A . PAUL
The involvement of microorganisms, aromatic substrates, and aminonitrogen-containing materials in humus formation has been well demonstrated (Kutzner, 1968; Flaig, 1964; Ladd and Butler, 1966). During
the growth of Epicoccum nigrum on glucose-asparagine medium, Martin
et al. ( 1 967) noted production of a fungus-synthesized humic acid which
was very similar to that of naturally occurring material in soil. Its rate
of decomposition, reaction to acid hydrolysis and proteolytic enzyme
attack, elemental composition, content and distribution of functional
groups, base exchange capacity and molecular weight distribution closely
resembled corresponding properties of natural humic materials. Values
for a number of these characteristics for Epicoccum-synthesized material
and Leonardite humic acid are shown in Table XVI.
Grassland soils are typified by their high content of organic matter
stabilized by the presence of calcium and sesquioxides and by the inherent resistant nature of the organic materials. The classical fractionation technique in which acid pretreatment and alkali extraction and/or
chelating agents are used to remove organic materials makes it possible
to disperse much of the soil carbon for later analyses. The precipitation
of humic acids under acidic conditions, or high concentrations of electrolyte, separates these materials on the basis of molecular weight, functional groups and aromaticity (Posner et al., 1968; Botner, 1967: Lindqvist,
1968). It does not necessarily separate them on the basis of their dynamic
or turnover rate in nature.
Carbon-dating analysis has indicated that the separation of soil into
fulvic and humic acids, although of use from a pedogenic standpoint,
does not result in fractions which are meaningful from a turnover viewpoint (Paul, 1970). The amino-rich materials of chernozemic soils have
a low mean residence time but are closely associated with the resistant
aromatic fractions which comprise the majority of soil carbon. The humic
acid hydrolyzate, containing 7% of the carbon and 15% of the nitrogen
in the humate, was found to be the most easily degradable and thus could
be expected to contribute the largest proportion of nitrogen and carbon
mineralized on an annual basis.
The use of 14C-labeledplant material has shown that undecomposed
plant and microbial components not only appear among the soluble fulvic
acids, but also contribute to the acid-insoluble fractions (Sauerbeck and
Fuhr, 197 1; Hardisson and Robert-Gero, 1966). Similarly, labeled carbon is distributed throughout the organic matter fractions after incubation
of glucose or plant constituents for even short periods of time (Simonart