Microbial community analysis of earthworm burrow wall

by Community Level Physiological Profiles using BIOLOG

“We know more about the movement of celestial bodies than about the soil

underfoot.”

Leonardo DaVinci

Introduction

Soil probably harbors most of our planet’s undiscovered biodiversity. Soils differ from

the above-ground and aquatic ecosystems because of their immense biological and

physicochemical diversity and complexity. One of the main reservoirs of microbial

diversity on the planet, soils sustain an enormous diversity of microbes. The ecological

functions of soil depend on a healthy and dynamic community of soil biota. This could

provide a range of information about the origins of microbial functional diversity as well

as novel genetic resources. Microbiologists have been investigating the impact of

microbial diversity on the stability of ecosystem function since the 1960s (Harrison et al.,

1968). Now there is heightened interest in the effect that the diversity of microbial

communities have on ecological function and resilience to disturbances in soil

ecosystems. Soil microbial communities are often difficult to fully characterize, mainly

because of their vast phenotypic and genotypic diversity, heterogeneity and crypticity.

The fraction of the cells making up the soil microbial biomass that have been cultured

and studied in any detail are negligible, often less than 5% (Borneman and Triplett 1997;

Torsvik et al., 1990). Traditional methods to analyze soil microorganisms have been

based on cultivation and isolation (Van Elsas et al., 2002); a wide variety of culture

media has therefore been designed to maximize the recovery of diverse microbial groups.

New methods have been developed over years to overcome the limitations due to culture

techniques and access a significantly greater soil microbial diversity. Method that allows

observation of the functional potential of microbial communities using the Biolog system

was described by Garland and Mills (1991). This method shows the potential degrading

abilities of a community in culture conditions. Winding (1994) has demonstrated that

these metabolic profiles can reveal differences between microbial communities that come

both from different kinds of soils and from different soil fractions. Known as the ‘Sole

Carbon Source Utilization’ (SCSU) or ‘Community Level Physiological Profiles’

(CLPP), the data can also be used to assess microbial diversity with measures such as the

Shannon Index (Zak et al., 1994). The above information allows examination of natural

variation and diversity of microbial community and offers the potential changes in

microbial diversity spatially and temporally. Turco et al., (1994) have also suggested that

SCSU is one of the several techniques that should be considered for development as

biological indicators to monitor potential changes in soil quality. This can provide an

otherwise hard-to-attain insight into the biology and evolution of environmental micro-

organisms. The application of this type of information is limitless, as are the avenues of

exploration.

Community Level Physiological Profile using BIOLOG

To gain better insight to the microbial processes within an ecosystem, it is essential to

study functional diversity in combination with taxonomic diversity. Most studies of

microbial community structure have involved isolate-based methods that depend on

cultural methods, which can exclude the majority of endogenous microbes from study

due to the selective nature of the media (Trousseilier and Legendre 1981). Recent studies

have attempted to characterize the portion of the microbial community that responds to

nutrient availability by comparing community fingerprints after incubation in individual

BIOLOG wells (Smalla et al., 1998). The application of the community-level approach to

assays of microbial function would provide a more sensitive and ecologically meaningful

measure of heterotrophic microbial community structure and metabolic abilities of the

community.

The Biolog plate method was first used to compare metabolic activity of heterotrophic

microbial communities from different habitats – water, soil and wheat rhizosphere

(Garland and Mills 1991). The technique is a redox system. During incubation at a

constant temperature, soil microorganisms oxidize substrates in the plate wells and,

simultaneously, reduce colourless tetrazolium dye to a violet formazan. Colour

development is measured spectrophotometrically (Kelly and Tate 1998). The rate of

utilization of different substrates by different groups of microorganisms varies, so one

can observe high variability in the rate of colour development and its intensity depending

on the composition of a microbial community. The characteristic pattern, called microbial

“metabolic fingerprint” (Baudone et al., 2001) or “community-level physiological profile

(CLPP)” (Gomeze et al., 2004), is obtained in this way. The diversity of soil microbial

communities can be characterized using the utilization pattern of individual carbon (C)

substrates generated with commercially available 96-well Biolog microtiter plates

(Garland and Mills 1991; Insam 1997).These CLPPs provide a rapid and relatively

inexpensive means of assessing differences in the soil microflora (Campbell et al.,,1997;

Staddon et al.,, 1998). Microbial community analyses based on CLPPs have been

corroborated by other microbial community measures, including plate counts (Wunsche

et al.,1995; Soderber et al.,2002), fatty acid methyl ester and phospholipid fatty acid

analysis [Lawlor et al., 2000; Widmer et al., 2001), API 20NE enzyme and C tests (Truu

et al., 1999), and an array of molecular assays (Widmer et al., 2001, Zvreafis and Torsvik

1998; Di Giovanni et al.,1999). In addition, research has demonstrated that CLPPs are

highly reproducible (Bossio and Skow 1995; Di Giovanni et al., 1999).

Types of Biolog plates

Commercially available microplates from Biolog, Inc. (Hayward, CA, USA) allow for

simultaneous testing of 95 separate carbon sources which include carbohydrates, amino

acids, carboxylic acids, amines, amides and polymers. Additionally, each well contains a

colourless tetrazolium dye. Color produced from the reduction of tetrazolium violet is

used as an indicator of respiration of sole carbon sources. Direct incubation of whole

environmental samples in BIOLOG plates, produces patterns of metabolic response

suitable for the rapid classification of heterotrophic microbial communities (Garland and

Mills 1991). There are various types of plates available for this purpose. The plates used

in this study are described below.

Biolog GN/GN2 plates

The most popular of the biolog plates, these have been developed for Gram- negative

organisms and contain 95 different carbon substrates in wells and no substrate in one well

which is used as control. These plates have also been used for ecological research of soil

microorganisms. Stefanowicz (2006) has reported that many scientific publications

related to microbial community analyses on the Biolog plates, deal with the use of GN

plates. The Biolog GN2 plates, which are a modification of the GN plates, have been

developed which have partly different substrates than original GN plates. These changes

have been introduced to restrain colour development, which is not connected with

microbial activity (O’connel and Garland 2002).

Biolog Eco plates

ECO plates were specifically developed for bacterial community analyses of

environmental samples and were first described by Insam (1997). They have been

typically designed for ecological study of whole microbial communities and not for

identification of microbial strains (Preston-Mafham et al., 2002). Selected especially for

ecological research these plates contain 31 unique C substrates that are purportedly more

relevant to the ecological functions these organisms perform within ecosystems and one

well with water (Choi and Dobbs 1999). Nine of the 31 substrates are known as com-

ponents of exudates of plants roots (Campbell et al., 1997). To help account for

variability in inoculum densities derived from environmental samples, substrates are

replicated three times within each ECO plate (Choi and Dobbs 1999; Gamo and Shoji

1999).

Biolog MT plates

The Biolog MT plates contain redox dye and no substrates. These plates enable the

choice of any substrate according to the researchers’ needs (Preston-Mafham et al., 2002;

Haack et al., 1995).

Data collection and Interpretation

Colour development due to the reduction of tetrazolium dye is read using a

spectrophotometer. Raw optical density data are corrected by blanking each response

well against its own first reading (immediately after inoculation). This not only avoids the

intrinsic absorbance of the carbon sources but also the negative values when compared to

subtracting the control well from the response well. The Average Well Colour

Development (AWCD) is calculated from each plate at each reading time. The data

obtained can also be used to calculate the Simpson, Shannon and McIntosh diversity

index. Each of the indices used is influenced by the data in different ways. The Simpson

index is weighted toward the abundances of the most common species, whereas the

Shannon index is influenced more by species richness (Magurran 1988). In contrast, the

McIntosh index is a measure of uniformity (Atlas 1984).Principal Component Analysis

(PCA) of the Biolog color responses allows for comparison of microbial samples on the

basis of differences in the pattern of sole-carbon-source utilization. The principal

components (PC) score plots describe the characteristics of the samples and help to

clarify their distribution and clustering. The PC score plot (PC-I and PC-II) shows the

spatial distribution of the samples. The relationships among different samples on the

basis of either the raw-difference data or the transformed data can be determined by PCA,

This technique projects the original data onto new axis (PCs) which reflect any intrinsic

pattern in the multidimensional data swarm (Pielou 1984). Each PC extracts a portion of

the variance in the original data, with the greatest amount of variance extracted by the

first axis. By correlating PCs with the original variables, the axes can be "labeled" with a

subset of the original variables. Relationships among samples are observed by plotting

samples in two dimensions on the basis of their scores for the first two PCs. PCA also

calculates the proportion of variance in each variable explained by a given PC. The

comparisons are a posteriori in nature; PCA determines how the samples are different,

but does not test among samples for specific differences selected a priori.

Microbial communities have great potential for temporal or spacial change and hence

represent a powerful tool for understanding community dynamics in both basic and

applied ecological contexts (Mishra and Nautiyal 2009). The present study describes

changes in microbial activity and community composition of burrow walls and their

respective control soils of two species of earthworms viz., L. mauritii and P. corethrurus

at different time intervals and depth. The Biolog plates apart from the estimation of the

metabolic diversity of the community in the burrow wall and control soil can also be used

to gather information of the natural variation and diversity of microbial communities in

these areas. The data from these experiments can be used in two ways (1) to quantify

differences among the lower and upper burrow wall soils and their controls and (2) to

assess the functional diversity of microbes in these areas.

Materials and Methods

Generation of Soil Samples

The soil sample was generated as described in chapter 2.

Analysis using BIOLOG plates

Biolog GN2, Eco plates and MT (Biolog, Inc., Hayward, CA, USA) were used to

determine the carbon source utilization pattern (Campbell et al., 1997) of the earthworm

burrow wall soil and control samples. Substrate, dye, and nutrients were supplied in each

well in a dried-film form which was reconstituted upon addition of sample (Bochner

1989).

The Biolog GN2 plates used included 6 amines/amides, 20 amino acids, 28

carbohydrates, 24 carboxylic acids, 5 polymers and 12 miscellaneous substrates totaling

95. The details of various substrates used are shown in Table 7.1.

Table 7.1 Substrates used in Biolog GN2 plates

Water

5 Maltose

10 α-Keto Glutaric Acid

AMINES/ AMIDES

6 D-Raffinose

11 α-KetobutyricAcid

1 2-Aminoethanol

7 L-Rhamnose

12 α-Keto Valeric Acid

2 Glucuronamide

8 Sucrose

13 β-Hydroxybutyric Acid

3 L-Alaninamide

9 N-Acetyl-DGlucosamine

14 D-Galactonic Acid Lactone

4 Phenyl ethyl amine

10 D-Arabitol

15 D-Galacturonic Acid

5 Putrescine

11 D-Cellobiose

16 D-Gluconic Acid

6 Succinamic Acid

12 i-Erythritol

17 D-Glucuronic Acid

AMINO ACIDS

13 L-Fucose

18 D-Saccharic Acid

1 γ-Amino Butyric Acid

14 Gentiobiose

19 Formic Acid

2 D,L-Carnitine

15 m-Inositol

20 Quinic Acid

3 D-Alanine

16 α-D-Lactose

21 Sebacic Acid

4 D-Serine

17 Lactulose

22 Itaconic Acid

5 Glycyl-L Aspartic

Acid

18 D-Mannitol

23 γ- Hydroxybutyric Acid

6 Glycyl-L Glutamic

Acid

19 D-Mannose

24 p-Hydroxy Phenylacetic Acid

7 Hydroxy L Proline

20 D-Melibiose

MISCELLANEOUS

8 L-Alanine

21 β-Methyl-D Glucoside

1 2,3-Butanediol

9 L-Aspartic Acid

22 D-Psicose

2 Bromosuccinic Acid

10 L-Asparagine

23 D-Sorbitol

3 D,L-α-Glycerol Phosphate

11 L-Alanylglycine

24 D-Trehalose

4 α-D-Glucose-1-Phosphate

12 L-Glutamic Acid

25 Turanose

5 Inosine

13 L-Histidine

26 Xylitol

6 Glycerol

14 L-Leucine

27 N-Acetyl-D

Glalactosamine

7 Thymidine

15 L-Ornithine

28 Adonitol

8 Uridine

16 L-Phenylalanine

CARBOXYLIC ACID

9 Urocanic Acid

17 L-Proline

1 Cis-Aconitic Acid

10 Pyruvic Acid Methyl Ester

18 L-Pyroglutamic Acid

2 Acetic Acid

11 Succinic Acid Mono-Methyl-

Ester 19 L-Serine

3 Citric Acid

12 D-Glucose-6-Phosphate

20 L-Threonine

4 D-Glucosaminic Acid

POLYMERS

CARBOHYDRATES

5 D,L-Lactic Acid

1 α- Cyclodextrin

1 L-Arabinose

6 Malonic Acid

2 Dextrin

2 D-Fructose

7 Propionic Acid

3 Tween 40

3 D-Galactose

8 Succinic Acid

4 Tween 80

4 α-D-Glucose

9 α-Hydroxybutyric Acid

5 Glycogen

Biolog Eco Plates contained 31 unique C substrates 9 of which are known as components

of exudates of plants roots (Campbell et al., 1997). The substrates used in the Eco plates

is shown in Table 7.2

Table 7.2: Substrates in Biolog Eco plates

1 2- Hydroxy Benzoic acid 17 L- Asparagine

2 4- Hydroxy Benzoic acid 18 L- Phenylalanine

3 D- Glucoaminic acid 19 L- Serine

4 D, L - α – Glycerol Phosphate 20 L- Threonine

5 D- Cellobiose 21 N- Acetyl- D- Glucosamine

6 D- Galactonic acid γ- Lactone 22 Phenylethylamine

7 D- Galacturonic acid 23 Putrescine

8 D- Malic acid 24 Pyruvic acid Methyl Ester

9 D- Mannitol 25 Tween 80

10 D- Xylose 26 Tween 40

11 Glucose -1- Phosphate 27 α – Cyclodextrin

12 Glycogen 28 α – D- Lactose

13 Glycyl-L- Glutamic acid 29 α – Ketobutyric acid

14 i- Erythritol 30 β – Methyl- D- Glucoside

15 Itaconic acid 31 γ- Hydroxybutyric acid

16 L- Arginine

The Biolog MT plates used had the following 10 substrates.

1. Casein 2. Oxalic acid

3. Cellulose 4. p-nitrophenol

5. Chitin 6. Pectin

7. Gelatin 8. Phytic acid

9. Lignin 10. Urea

Sample preparation

Procedure

Plates were prepared using the manufacturer’s instructions (Biolog Inc., Hayward, CA,

U.S.A).

1. Individual burrow wall and control soil samples (1.0 g) were shaken in 9.0 ml of

sterile 0.85% saline for 2 hours and then made up to a final dilution 10-2

.

2. 150 µl of sample was inoculated in each well of Biolog GN2, Eco and MT plates

and incubated at 300C.

3. The rate of utilization was indicated by the reduction of tetrazolium, a redox

indicator dye, which changes from colorless to purple.

4. Data were recorded for days 1-15 at 590 nm (Mishra and Nautiyal 2009).

Data analysis

1. Microbial activity in each microplate, expressed as average well color

development (AWCD) was determined (Garland, 1996).

2. Diversity and evenness indices were calculated as described by Mishra and

Nautiyal (2009). Formulae used for diversity calculations are given in Table

7.3

3. Principal component analysis (PCA) was performed on 10th

day data divided

by AWCD (Garland and Mills, 1991).

4. Statistical analyses were performed using SPSS 16.0 for significance and

Statistica 7.0 for PCA.

Table 7.3: Description of the diversity calculations used

Index Definition Formula Definitions

Shannon

diversity

Measure of richness or

evenness ∑

Pi = Proportional

color development

of the ith well over

total color

development of all

wells of a plate

Shannon

evenness

Evenness calculated from

Shannon index

S= number of well

with color

development

Simpson

index

Dominance index sensitive to

the abundance of the most

common species

∑ ( ( ))

( ( ))

ni= color

development of the

ith well. N= total

color development.

Values shown as

1/D

McIntosh

diversity

Measure of diversity based

upon the Euclidian distance

of the community of the

assemblage in an S

dimensional hypervolume

(Index of uniformity)

√(∑ )

McIntosh

evenness

Evenness calculated from

McIntosh index

√ ⁄

Results

Microbial metabolic activity using Biolog ECO Plates

In L. mauritii highest metabolic activity was seen in 45 days upper burrow wall soil with

a log phase from day 2 which continued up to day 15 (Figure 7.2). The 45 days upper

control soil showed less microbial activity with increase only from day 4. Lower burrow

wall soil at 45 days too showed a higher microbial metabolic activity from day 3 and kept

increasing whereas 45 days lower control showed the least activity. An opposite result

was seen in 30 day trials. Both upper and lower burrow wall soil showed lesser metabolic

activity compared to control. The lower burrow wall showed a higher activity compared

to upper burrow wall soil. In P. corethrurus (Figure 7.1) the highest microbial activity

was seen in 45 days lower burrow wall soil which was much higher than all other

samples. This was followed by 30 days lower burrow wall soil and 45 days upper burrow

wall soil both of which were higher than their respective control soil samples. The 30

days upper burrow wall soil showed lesser microbial activity compared to control soil.

All samples showed log phase after day 3 except for 45 days upper burrow wall soil

where there was no lag phase seen.

Principal Component Analysis (PCA) based on carbon utilization

The PCA based on carbon utilization in L. mauritii (Figure 7.4) showed a clustering of 45

days lower and upper burrow wall samples. The 30 days upper and lower burrow wall

samples showed a drift in microbial community and were placed distantly from each

other. A similar result was also seen with 30 and 45 days lower burrow wall sample and

control soil. The non earthworm worked soil was observed to cluster with 45 days lower

and upper burrow wall soil. In P. corethrurus (Figure 7.3) the 30 days upper and lower

burrow wall soil and 45 days upper control soil was observed to be in a cluster. All the of

30 and 45 days trials in both upper and lower burrow wall soil were found to be placed

distantly from their respective control soil. Non earthworm worked soils was placed

distinctly away from any of the clusters.

Diversity and evenness index based on carbon utilization

The diversity and evenness index in L. mauritii samples showed a significant difference

between the 30 day burrow wall and control soil both in the upper and lower half of the

setup (Table 7.5). Here the diversity was significantly higher in the control soil. In the 45

day trials the lower burrow wall soil showed significantly higher values compared to

control but the upper burrow wall though showed a higher value does not show a

significant difference. It is important to note that the non earthworm worked soil was

significantly different from the burrow wall soil of both 30 and 45 days and did not show

a significant difference when compared to the control soils in all trials and intervals.

In P. corethrurus the different diversity indices did not show significant difference

between the burrow wall and control soils (Table 7.4). It was significant though that the

non earthworm worked soil was different from the earthworm worked soils.

Categorized carbon substrate utilization pattern

Substrate utilization profiles of the microbial community at various intervals and depths

differed significantly in soil worked with L. mauritii. The categorized carbon substrate

utilization pattern in L. mauritii is indicated in Figure 7.6. The utilization of amines and

carboxylic acid shows an increase in 30 days upper and lower burrow wall soils and 45

days upper burrow wall soils whereas a decrease was seen in 45 days lower burrow wall

soils compared to control soil. In P. corethrurus (Figure 7.5) utilization of amines

increased in 45 days upper and lower burrow wall samples and 30 days upper burrow

wall samples but decreased in 30 days lower burrow wall soils. Amino acid utilization

increased in all burrow wall samples except 45 days upper burrow wall soils in L.

mauritii and also showed an increase in 30 and 45 days upper and lower burrow wall

soils compared to control soil. In L. mauritii the carbohydrate utilization pattern showed

an increase in 30 and 45 days upper burrow wall soils and a decrease in 45 days lower

burrow wall soils. No difference was seen in the utilization pattern in the 30 days burrow

wall and control soil. In P. corethrurus carbohydrate utilization pattern was seen to

increase in both 45 and 30 days upper burrow wall soil and decrease in 30 and 45 days

lower burrow wall soils. In P. corethrurus carboxylic acid utilization pattern decreased in

30 and 45 days upper burrow wall soils and increased in 30 and 45 days lower burrow

wall soils compared to control soils. Polymer utilization patterns showed a decrease in 30

days upper burrow wall soil and 45 days lower burrow wall soil and increased in 30 days

lower burrow wall soil and 45 days upper burrow wall soil in L. mauritii. In P.

corethrurus polymer utilization was seen to decrease in 30 days burrow wall soil and

increase in 45 days burrow wall soil compared to control. It is interesting to note that the

45 days lower burrow wall sample had microbes with highest polymer degrading ability.

The miscellaneous carbon utilization pattern showed an increase only in 30 days lower

burrow wall soil sample in L. mauritii whereas in P. corethrurus an increase was seen in

both 30 and 45 lower burrow wall samples compared to control. Except for utilization of

amines the non- earthworm worked control showed less utilization of all other carbon

sources.

Utilization pattern of carbon substrates in MT plates

The results of the utilization pattern of carbon substrates in MT plates indicate the

following in L. mauritii. The utilization of urea, cellulose and p- nitro phenol is seen to

decrease in the 30 day lower and upper burrow wall soil compared to control and increase

in the 45 days upper and lower burrow wall soil. The utilization of p- nitro phenol was

seen to be the highest in all samples. Lignin utilization increased in all burrow wall soil

samples studied (Figure 7.8). In P. corethrurus too, a similar pattern of utilization was

seen as far as urea, cellulose and p- nitro phenol is concerned (Figure 7.7). The pattern of

utilization of pectin showed that except in the 30 days lower burrow wall soil where there

was a decrease compared to control, in all other samples an increase was seen in L.

mauritii worked soils. In P. corethrurus an increase in the pectin utilization was seen in

the 45 days upper and lower burrow wall soil and a decrease in the 30 days upper and

lower burrow wall soil. The pattern for the utilization of chitin showed an increase in the

burrow wall soil in the 45 day trials compared to control in both P. corethrurus and L.

mauritii. Phytic acid utilization increased in both the species of earthworms studied in

the 45 day trials. In L. mauritii an increase in the utilization of casein and gelatin was

seen in the 45 day trials of both upper and lower burrow wall soil. In P. corethrurus the

utilization of casein showed a decrease in the 30 and 45 days upper burrow wall soil and

no change in the 30 and 45 days lower burrow wall soil compared to control. Pattern for

utilization of gelatin showed a decrease in the upper and lower burrow wall 30 day trials

and in the 45 days upper burrow wall compared to control. In the 45 days upper burrow

wall soil there was an increased utilization of gelatin. In L. mauritii the utilization pattern

of oxalic acid decreased in the 30 and 45 days upper burrow wall soil and increased in the

30 and 45 days lower burrow wall soil. In P. corethrurus there was increased utilization

of oxalic acid only in the 45 days upper burrow wall soil.

Discussion

Attempts to understand diversity at the microbial level are hindered by the fact that

microbial communities typically consists of diverse microorganisms of which only small

percentage can be cultured and taxonomically identified. Analyzing sole carbon source

utilization patterns by microbial samples using Biolog microplates which rely on

measurements of utilizing different carbon substrates by microorganisms is a means of

assessing soil microbial community structure. This examines the functional capabilities

of the microbial population and the resulting data can be analyzed using multivariate

techniques to compare metabolic capability of communities (Garland and Mills 1991;

Zak et al., 1994).

Effects of earthworms on the microbial community depend, in part, on the timing of the

measurement. Some effects of earthworms may become apparent only after an extended

period of time because changes that affect microbial community composition and trophic

interactions, such as diffusion of nutrients beyond the burrow walls and development of

pore structure in the burrow walls, may occur gradually. The present study shows high

microbial metabolic activity in the burrow wall soil especially in the 45 day samples of L.

mauritii and P. corethrurus reflecting that with the increase in time the effect of these

species of earthworms increases. It was interesting to note that in the 30 day trials the

burrow wall soil showed a decreased activity compared to control in both species of

earthworms studied. This report is the first of this kind as far as CLPP is concerned.

There have been contrasting effects on microbial biomass, with microbial biomass

increasing, decreasing,

or showing no net change relative to soil unaffected by

earthworms (Brown 1995). Clegg et al., (1995) found that total bacterial counts in burrow

and bulk soil were initially no different, but increased through time in casts and remained

elevated compared with bulk soil. Also at greater depths the contrast between drilosphere

and bulk soil and the relative contribution of burrow walls to the total soil microbial

activity probably increase strongly (Stehouwer et al., 1993; Joergensen et al., 1998).

Principal Component Analysis (PCA) based on carbon utilization in the present study

shows a clustering of the 45 days burrow wall soils while the 30 day soil were distantly

placed in L. mauritii. The non earthworm worked soils was found to cluster with the

burrow wall soil. In P. corethrurus the burrow wall soil, their respective control and the

non earthworm soils were placed distantly from each other indicating that burrow wall

soils are different from the controls. Tiunov and Dobrovolskaya (2002) observed that

both bacterial and fungal communities in the burrow wall soil differed significantly from

the control soil. Idowu et al., (2006) in a related study also reported that the total aerobic

and anaerobic counts of microflora were higher in casts than in surrounding soil. The

diversity and evenness index reflect the fact that in L. mauritii the burrow wall soils are

different from the control soils whereas in P. corethrurus there is no difference though it

is different from the non earthworm worked soils. In L. mauritii in all the samples the

burrow wall soil showed a significant difference in the diversity compared to control.

This reiterates the results of many studies by traditional methods (Tiunov and Scheu

1999; Barois 1992; Daniel and Anderson 1992; Edwards and Fletcher 1988; Kristufek et

al., 1992). The values of diversity indices in L. mauritii and P. corethrurus were on the

higher side indicating that most of the carbon sources were utilized by the microbial

community. The values were found to be higher in L. mauritii worked soils compared to

P. corethrurus. The categorized carbon substrate utilization pattern clearly indicates that

the burrow wall soil microbes have better ability to degrade various carbon sources and

could increase the soil fertility.

The utilization pattern of urea and cellulose in L. mauritii and P. corethrurus was seen to

follow the same pattern with a decrease in the 30 day lower and upper burrow wall soil

compared to control and increase in the 45 days upper and lower burrow wall soil

demonstrating that as the time increases these specific organisms show an increase.

Lampito mauritii worked soils could be a source of lignin, pectin, p- nitro phenol, casein,

gelatin and oxalic acid degrading organisms. Chitin and phytic acid degrading organisms

were increased in 45 day trials indicating that the longer the soil was worked with

earthworms better the soil fertility. It is interesting to note that the cellulose degrading

organisms were not activated by L. mauritti but also decreased as time progressed.

Pontoscolex corethrurus worked soils could be a source of p nitrophenol, lignin, pectin,

cellulose, phytic acid degrading organisms. Chitin degrading organism increased with

time in the burrow wall while gelatin utilizing organisms decreased with time. P-

Nitrophenol is recognized as an environmental contaminant; it is used primarily for

manufacturing medicines and pesticides (Spain and Gibson 1991). Species known to

degrade p- nitrophenol include species of Arthrobacter, Pseudomonas, Acinetobacter,

Alcaligenes and Moraxella. Oxalic acid a naturally occurring, highly oxidized organic

compound with powerful chelating activity is a widely occurring natural product of

animals, plants and other organisms. It sometimes occurs as a free acid, but more

commonly as a calcium salt. Fungi, bacteria, and actinomycetes are able to metabolize

oxalate. Common species include Ralstonia, Alcaligenes, Pseudomonas,

Methylobacterium, Pandoraea and Streptomyces. Microbial phytate-degrading activity is

detected in numerous micro-organisms. Extracellular phytate-degrading activity was

observed nearly exclusively in A. niger strains (Gargova et al., 1997). Phytate-degrading

enzymes have also been detected in species of Pseudomonas (Richardson and Hadobas

1997), E. coli (Greiner et al., 1993) and Enterobacter (Yoon et al., 1996).

There is a great deal of information suggesting that earthworm activity changes the

microbial community structure of soil. It has been proposed that earthworms have a

mutualistic relationship with microorganisms (Barois and Lavelle 1986; Lavelle et al.,

1995; Trigo and Lavelle 1993) and may contribute to the maintenance of soil fertility and

soil microbial diversity. The present study emphasizes the concept that earthworms, as

ecosystem engineers, play an important role in many soil ecosystems and improve soil

fertility and in turn impact plant growth. These two species studied do have significant

impacts on microbial diversity in soil.

Legend: 1- UCS (30 d), 2- LBWS (30 d), 3- UBWS (30 d), 4- LCS (30 d) , 5- LBWS (45 d), 6- LCS (45 d), 7- UCS (45 d) , 8- UBWS (45 d)

Figure 7.1: Microbial metabolic activity measured by optical density and expressed by average well color development (AWCD) i.e.,

substrate utilization pattern of burrow wall and control soil samples from P. corethrurus using Biolog Eco Plates. Error bars indicate

Standard errors

0

0.05

0.1

0.15

0.2

0.25

0.3

0.35

0.4

0.45

1 2 3 4 5 6 7 8 9 10 12 15

OD

590

nm

1

2

3

4

5

6

7

8

Legend: 9- UBWS (30 d);10-UCS (30 d); 11-LBWS (30 d);12- LCS (30 d);13- UBWS (45 d); 14- UCS (45 d); 15- LBWS (45 d);16- LCS (45 d);17- Non earthworm worked

soil

Figure 7.2: Microbial metabolic activity measured by optical density and expressed by average well color development (AWCD) i.e.,

substrate utilization pattern of burrow wall and control soil samples from L. mauritii using Biolog Eco Plates. Error bars indicate Standard

errors

0

0.2

0.4

0.6

0.8

1

1.2

1.4

1.6

1.8

1 2 3 4 5 6 7 8 9 10 12 15

OD

59

0 n

m

Time (days)

9

10

11

12

13

14

15

16

17

Projection of the variables on the factor-plane ( 1 x 2)

Active

1

23

4

56

7

8

9

-1.0 -0.5 0.0 0.5 1.0

Factor 1 : 20.43%

-1.0

-0.5

0.0

0.5

1.0

Facto

r 2 :

15.5

1%

Legend: 1- UCS (30 d), 2- LBWS (30 d), 3- UBWS (30 d), 4- LCS (30 d) , 5- LBWS (45 d), 6-

LCS (45 d), 7- UCS (45 d) , 8- UBWS (45 d) , 9- non earthworm worked soil

Figure 7.3: Principal component analysis based on carbon source utilization by

microflora of burrow wall and control soil samples from P. corethrurus

Projection of the variables on the factor-plane ( 1 x 2)

Active

9

10

11

12

13

14

15

16

17

-1.0 -0.5 0.0 0.5 1.0

Factor 1 : 61.53%

-1.0

-0.5

0.0

0.5

1.0

Fa

cto

r 2

:

9.6

9%

Legend: 9- UBWS (30 d);10-UCS (30 d); 11-LBWS (30 d);12- LCS (30 d);13- UBWS (45 d);

14- UCS (45 d); 15- LBWS (45 d);16- LCS (45 d);17- Non earthworm worked soil

Figure 7.4: Principal component analysis based on carbon source utilization by

microflora of burrow wall and control soil samples from L mauritii

Table 7.4: Diversity and Evenness index calculated based on substrate utilization by microflora of burrow wall and control soil

samples from P. corethrurus

LBWS

(30 days)

LCS

(30 days)

UBWS

(30 days)

UCS

(30 days)

LBWS

(45 days)

LCS

(45 days)

UBWS

(45 days)

UCS

(45 days)

Non

Earthworm

worked

soil Shannon

Diversity 3.67±0.08

abc 3.96 ±0.06

c 3.54±0.09

abc 3.48 ±0.17

ab 3.90±0.02

bc 3.80 ± 0.10

bc 3.82 ± 0.14

bc 3.36±0.21

a 4.65±0.03

d

Mcintosh

Diversity 0.88 ± 0.01

ab 0.92 ±0.01

b 0.85 ±0.01

ab 0.85 ±0.03

ab 0.93 ± 0.01

b 0.87 ± 0.03

ab 0.89 ± 0.03

ab 0.82±0.04

a 0.98±0.00

c

Simpson

Diversity 0.97 ± 0.00

ab 0.98 ±0.00

b 0.95±0.01

ab 0.95 ±0.01

ab 0.98 ± 0.00

b 0.96 ± 0.01

ab 0.97 ± 0.01

ab 0.94±0.02

a 1.00±0.00

c

Shannon

Evenness 0.81±0.01

abcd 0.86±0.01

cd 0.76 ±0.02

ab 0.77±0.04

abc 0.88 ± 0.01

d 0.83±0.02

abcd 0.86±0.03

bcd 0.74±0.05

a 0.96±0.01

f

McIntosh

Evenness

0.89 ±

0.01abc

0.93±0.01

bc 0.85 ±0.01

ab 0.86 ± 0.03

ab 0.94 ± 0.00

c 0.88 ±0.03

abc 0.91 ±0.03

abc 0.83±0.04

a 0.99±0.00

d

Different letters showing significant difference at p = 0. 05 using Waller Duncan test

Legend: LBWS- Lower Burrow Wall Soil; LCS- Lower Control Soil; UBWS- Upper Burrow Wall Soil; UCS- Upper Control Soil

Table 7.5: Diversity and Evenness index calculated based on substrate utilization by microflora of burrow wall and control soil

samples from L. mauritii

L.

mauritii. UBWS (30 d) UCS (30 d) LBWS (30 d) LCS (30 d)

UBWS (45

d) UCS (45 d)

LBWS (45

d) LCS (45 d)

Non

earthworm

worked

soil Shannon

Diversity

4.03 ±0.03b 4.51±0.03

d 4.38±0.06

c 4.54±0.01

de 4.63±0.03

de 4.55±0.02

def 4.52±0.03

d 3.72±0.02

a 4.65±0.03

f

Mcintosh

Diversity

0.92 ±0.01b 0.97±0.00

d 0.96±0.01

c 0.98 ±0.00

d 0.98±0.00

d 0.98±0.00

d 0.97±0.00

cd 0.89±0.01

a 0.98±0.00

d

Simpson

Diversity

0.98 ±0.00b 1.00±0.00

cd 0.99±0.00

c 1.00±0.00

d 1.00±0.00

d 1.00±0.00

d 0.99±0.00

cd 0.97±0.00

a 1.00±0.00

d

Shannon

Evenness

0.88±0.01a 0.94±0.00

bc 0.92 ±0.01

b 0.96±0.00

c 0.95±0.00

c 0.95±0.00

c 0.95±0.01

bc 0.90±0.01

a 0.96±0.01

c

McIntosh

Evenness

0.94 ±0.01a 0.98 ±0.00

bc 0.97 ±0.01

b 0.98±0.00

c 0.98±0.00

c 0.98

±0.00bc

0.98±0.00bc

0.94±0.00a 0.99±0.00

c

Different letters showing significant difference at p = 0. 05 using Waller Duncan test

Legend: LBWS- Lower Burrow Wall Soil; LCS- Lower Control Soil; UBWS- Upper Burrow Wall Soil; UCS- Upper Control Soil

Legend: 1- UCS (30 d), 2- LBWS (30 d), 3- UBWS (30 d), 4- LCS (30 d) , 5- LBWS (45 d), 6- LCS (45 d), 7- UCS (45 d) , 8- UBWS (45 d)

Figure 7.5: Categorized carbon substrate utilization pattern by microflora of burrow wall and control soil samples from P.

corethrurus

0

0.5

1

1.5

2

2.5

3

3.5

4

1 2 3 4 5 6 7 8 9

OD

59

0 n

m

AMINES/AMIDES AMINO ACIDS CARBOHYDRATES CARBOXYLIC ACID POLYMERS MISCELLANEOUS

Legend: 9- UBWS (30 d);10-UCS (30 d); 11-LBWS (30 d);12- LCS (30 d);13- UBWS (45 d); 14- UCS (45 d); 15- LBWS (45 d);16- LCS (45 d);17- Non

earthworm worked soil

Figure 7.6: Categorized carbon substrate utilization pattern by microflora of burrow wall and control soil samples from L. mauritii

0

0.5

1

1.5

2

2.5

9 10 11 12 13 14 15 16 17

OD

59

0 n

m

AMINES/AMIDES

AMINO ACIDS

CARBOHYDRATES

CARBOXYLIC ACID

POLYMERS

MISCELLANEOUS

Legend: 1- UCS (30 d), 2- LBWS (30 d), 3- UBWS (30 d), 4- LCS (30 d) , 5- LBWS (45 d), 6- LCS (45 d), 7- UCS (45 d) , 8- UBWS (45 d), 9- Non earthworm

worked soil

Figure 7.7: Utilization pattern of carbon substrates in MT plates by microflora of burrow wall and control soil samples from P.

corethrurus

0

0.5

1

1.5

2

2.5

3

3.5

1 2 3 4 5 6 7 8 9

OD

59

0 n

mUrea Lignin Pectin Cellulose Phytic acid

Chitin Para-nitrophenol Casein Gelatin Oxalic acid

Legend: 9- UBWS (30 d);10-UCS (30 d); 11-LBWS (30 d);12- LCS (30 d);13- UBWS (45 d); 14- UCS (45 d); 15- LBWS (45 d);16- LCS (45 d);17- Non

earthworm worked soil

Figure 7.8: Utilization pattern of carbon substrates in MT plates by microflora of burrow wall and control soil samples from L.

mauritii

0

0.5

1

1.5

2

2.5

3

3.5

4

9 10 11 12 13 14 15 16 17

OD

590

nm

Urea Lignin Pectin Cellulose Phytic acid

Chitin Para-nitro phenol Casein Gelatin Oxalic acid

Plate 10