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V. Biomass and Bioactivity Measurements

V. Biomass and Bioactivity Measurements

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404



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



405



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

following section.

TABLE XIV

Biomass of Fungi and of Bacteria and Actinomycetes in

Matador Grassland Soil"

Fungi



Bacteria plus actinomycetes'



Month



0-10 cm



0-30 cm



0-10 cm



0-30 crn



April

May

June

July

August

September

October



51

35

40

63

56

53

65



I24

84

109

169

I53

153

177



12.3

23.4

27.2

19.4

25.9

14.7

20.6



31.9

70.9

76.2



58.2

67.0

42.3

54.7



Values are expressed as grams per square meter, dry weight.

"D. Parkinson (personal communication).

Babiuk and Paul ( 1970).



B. BIOMASSI N RELATION

TO MICROBIAL

ACTIVITY

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



406



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.



C. MEASUREMENT

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



407



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



P

0

00



TABLE X V

Enzyme Activity and Riogenic Nature of Organic Matter in Various Soils Typical of Horizontal Belts"



Polyphenol

oxidase



Catalase



Number of

organisms per gram

of organic matter

in thousands



5.8



2.5



220,000



12.8

11.0

10.6

8. I

7.0

6.4



6.8

5.1

6.3

4.7

3.9

3.0



190,000

47,000



Activity



Soil



Humus

(%)



Invertase



Amylase



Dehydrogenase



9.2b

25.4

45.0

29.7

53.4

66.8

87.2



2.0



2.3

5.8

16.5

8.3

18.4

8.9

11.3



0



5m

x



Dark brown

Chestnut

Leached chernozem

Brown

Meadow-steppe

Brown mountain-meadow

Mountain-meadow peat



1.7

3.0

7.2



7.8

18.1

15.3

21.6



5.6

9.0



10.2

10.2



12.1

14.8



~



50,000

12,000

6000

4000



m



I-



b



s

?

z

C



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).



r



THE MICROFLORA OF GRASSLAND



409



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

prove useful.

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.

A. CHARACTERISTICS

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.



410



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.



B. THEBIODEGRADABILITY

OF SOILHUMUS

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



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