Biochar and the Soil Nitrogen Cycle: Unravelling the ... › files › ...Biochar and the Soil Nitrogen Cycle: Unravelling the Interactions Daniel Norman Dempster BSc (Agric, Hons);

Biochar and the Soil Nitrogen Cycle:

Unravelling the Interactions

Daniel Norman Dempster

BSc (Agric, Hons); BComm

This thesis is presented for the degree of

Doctor of Philosophy

to The University of Western Australia

School of Earth and Environment

Faculty of Natural and Agricultural Sciences

2013

ii

iii

Table of Contents

Table of Contents ....................................................................................................... iii

Acknowledgements ..................................................................................................... ix

Declaration .................................................................................................................. xi

Publications arising from this Thesis ..................................................................... xiii

List of Abbreviations .............................................................................................. xvii

Abstract ..................................................................................................................... xxi

1. General Introduction and Literature Review ................................................. 1

1.1 General Introduction .................................................................................. 1

1.2 Western Australian Agriculture ................................................................. 1

1.3 Biochar ....................................................................................................... 4

1.3.1 Introduction to biochar ..................................................................... 4

1.3.2 Biochar in the agricultural system: focus on broadacre agriculture in

Western Australia ............................................................................. 8

1.3.3 Soil biological fertility changes with biochar addition ..................... 9

1.4 The Nitrogen Cycle .................................................................................. 13

1.5 Sorption of nitrogen to biochar ................................................................ 18

1.5.1 Mechanisms of sorption and properties influencing sorption to

biochar ............................................................................................ 18

1.5.2 The role of nitrogen sorption to biochar in ecosystem processes ... 21

1.6 Desorption ................................................................................................ 21

1.7 The influence of biochar on nitrogen leaching......................................... 22

1.7.1 Leaching of nitrogen decreases due to sorption.............................. 22

1.7.2 Leaching of nitrogen decreases due to increasing water holding

capacity ........................................................................................... 23

iv

1.7.3 Leaching of nitrogen changes due to nitrification and mineralisation

......................................................................................................... 23

1.7.4 Leaching of nitrogen changes due to microbial immobilisation ..... 25

1.8 Changes to nitrogen immobilisation induced by biochar application ...... 25

1.9 Biochar influences on nitrogen mineralisation ......................................... 27

1.10 The influence of biochar on nitrification .................................................. 30

1.11 The influence of biochar on plant nitrogen uptake ................................... 32

1.12 Summary of key research areas highlighted within this review ............... 34

2. Decreased soil microbial biomass and nitrogen mineralisation with

Eucalyptus biochar addition to a coarse textured soil .................................. 39

2.1 Abstract ..................................................................................................... 39

2.2 Introduction............................................................................................... 40

2.3 Materials and methods .............................................................................. 43

2.3.1 Biochar characterisation .................................................................. 43

2.3.2 Soil preparation and description ...................................................... 44

2.3.3 Pot experiment ................................................................................ 45

2.3.4 Plant analysis ................................................................................... 46

2.3.5 Microbial biomass ........................................................................... 47

2.3.6 Soil carbon dioxide evolution ......................................................... 48

2.3.7 Community level physiological profiles (CLPP) ............................ 48

2.3.8 DNA extraction and terminal restriction fragment length

polymorphism (T-RFLP) profiles ................................................... 48

2.3.9 Net nitrogen mineralisation ............................................................. 50

2.3.10 Organic carbon sorption experiment ........................................... 50

2.3.11 Nitrogen sorption experiment ..................................................... 50

2.3.12 Statistical analysis ....................................................................... 51

2.4 Results ...................................................................................................... 52

2.4.1 Biochar characterisation .................................................................. 52

2.4.2 Plant growth analysis ...................................................................... 53

2.4.3 Microbial biomass ........................................................................... 55

v

2.4.4 Microbial community function (CLPP) and structural diversity

(amoA T-RFLP) .............................................................................. 58

2.4.5 Carbon dioxide evolution and nitrogen mineralisation .................. 60

2.4.6 Sorption isotherms .......................................................................... 64

2.5 Discussion ................................................................................................ 67

2.5.1 Microbial biomass .......................................................................... 67

2.5.2 Carbon and nitrogen mineralisation ................................................ 69

3. Clay and biochar amendments decreased inorganic nitrogen leaching but

not dissolved organic nitrogen leaching in soil ............................................. 75

3.1 Abstract .................................................................................................... 75

3.2 Introduction .............................................................................................. 76

3.3 Materials and Methods ............................................................................. 78

3.3.1 Soil, clay and biochar ..................................................................... 78

3.3.2 Experimental design and setup ....................................................... 81

3.3.3 Nitrogen sorption characteristics .................................................... 82

3.3.4 Water retention ............................................................................... 82

3.3.5 Statistical analysis ........................................................................... 82

3.4 Results ...................................................................................................... 83

3.4.1 Nitrogen leaching ............................................................................ 83

3.4.2 Nitrogen sorption characteristics .................................................... 85

3.4.3 Water retention capacity ................................................................. 87

3.5 Discussion ................................................................................................ 89

4. Organic nitrogen mineralisation in two contrasting agro-ecosystems is

unaffected by biochar addition ....................................................................... 93

4.1 Abstract .................................................................................................... 93

4.2 Introduction .............................................................................................. 93

4.3 Materials and Methods ............................................................................. 95

4.3.1 Experimental design and characterisation of soils and biochars .... 95

4.3.2 Statistical Analysis .......................................................................... 97

4.4 Results .................................................................................................... 101

vi

4.5 Discussion ............................................................................................... 107

5. Minimal interaction between wheat chaff biochar and N fertiliser in a

broadacre field experiment ........................................................................... 111

5.1 Abstract ................................................................................................... 111

5.2 Introduction............................................................................................. 112

5.3 Materials and Methods ........................................................................... 114

5.3.1 Experimental site and design ........................................................ 114

5.3.2 Sampling times and analysis ......................................................... 118

5.3.3 Statistical analysis ......................................................................... 118

5.4 Results .................................................................................................... 119

5.4.1 Soil microbial biomass .................................................................. 119

5.4.2 Soil ammonium and nitrate ........................................................... 119

5.4.3 Plant biomass and nitrogen content ............................................... 120

5.4.4 Grain yield and nitrogen content ................................................... 122

5.5 Discussion ............................................................................................... 124

5.5.1 Interaction between nitrogen fertiliser and biochar....................... 124

5.5.2 Yield influence of biochar ............................................................. 125

5.5.3 Biochar application method .......................................................... 126

5.5.4 Conclusion ..................................................................................... 126

6. General Discussion ......................................................................................... 129

6.1 Sorption of low molecular weight nitrogen to biochar ........................... 131

6.1.1 Applying findings to other experiments ........................................ 133

6.1.2 Implications of the minimal impact of sorption of low molecular

weight nitrogen to biochar ............................................................ 134

6.2 Sorption of high molecular weight organic nitrogen compounds to

biochar .................................................................................................... 135

6.2.1 Implications of the sorption of high molecular weight compounds to

biochar ........................................................................................... 138

6.3 The addition of labile carbon contained in biochar ................................ 139

6.4 Increasing the supply of phosphorus and potassium in biochar ............. 141

vii

6.5 Water supply and water holding capacity .............................................. 142

6.6 No effect of biochar................................................................................ 143

6.7 Conclusion .............................................................................................. 145

7. References ....................................................................................................... 147

viii

Acknowledgements

ix

Acknowledgements

I acknowledge the Grains Research and Development Corporation (GRDC) for

funding my scholarship (GRS153) and for the travel awards received. I also

acknowledge the School of Earth and Environment within the University of Western

Australia for providing further operating expenditure.

I thank my supervisors Associate Professor Deirdre Gleeson, Professor Daniel

Murphy, and Winthrop Professor Lyn Abbott, especially Dan Murphy for facilitating

my collaboration with Professor Davey Jones. I also thank Professor Davey Jones for

supervising my research component at Bangor University, Wales.

My family deserve special thanks for providing me with many opportunities

throughout my life. Emily Laing has provided immense support and friendship. I also

thank many other friends for their discussion and thoughts.

x

Declaration

xi

Declaration

I, Daniel Norman Dempster, declare that this thesis was composed by me and that

research detailed within was designed, conducted and interpreted by myself, except in

instances where the work and contribution of others has been acknowledged.

Daniel N. Dempster

xii

Publications arising from this Thesis

xiii


Peer Reviewed Publications

Dempster, D.N., Gleeson, D.B., Solaiman, Z.M., Jones, D.L., Murphy, D.V. (2012)

Decreased soil microbial biomass and nitrogen mineralisation with Eucalyptus

biochar addition to a coarse textured soil. Plant and Soil 354, 311 – 324

(Chapter 2).

Dempster, D.N., Jones, D.L., Murphy, D.V. (2012) Clay and biochar amendments

decreased inorganic but not dissolved organic nitrogen leaching in soil. Soil

Research 50, 216 – 221 (Chapter 3).

Dempster, D.N., Jones, D.L., Murphy, D.V. (2012) Organic nitrogen mineralisation

in two contrasting agro-ecosystems is unchanged by biochar addition. Soil

Biology & Biochemistry 48, 47 – 50 (Chapter 4).

Conference Presentations


Biochar changed microbial dynamics and nitrogen mineralisation. 19th World

Congress of Soil Science, Brisbane, August 2010 (poster).


xiv


Biochar changed microbial dynamics and nitrogen mineralisation. UK Biochar

Conference, Edinburgh, Scotland, May 2011 (poster).

Dempster, D.N., Jones, D.L., Murphy, D.V. (2011) Sporadic effects of biochar on

organic nitrogen decomposition. Australian Society of Soil Science Inc, WA

Branch Conference, Busselton, September 2011 (oral presentation).

Dempster, D.N., Jones, D.L., Murphy, D.V. (2011) Sporadic effects of biochar on

organic nitrogen decomposition. 3rd International Rhizosphere Conference,

Perth, September 2011 (oral presentation).

Other Publications and Presentations

Dempster, D.N. (2009) Biochar attracts attention. Ground Cover: Soil Health

Supplement, May-June 2009, pp 18 – 19.

Dempster, D.N., Wherrett, A. (2010). Biochar trial update. Liebe group “Spring on

the Sandplain” field day, Buntine, October, 2010. Oral given by A. Wherrett

and D. Dempster.

Dempster, D.N. (2011). Biochar saving fertiliser? Results from the first year of the

Liebe group trial. Liebe group 2011 crop updates. Paper and presentation by D.

Dempster.


xv

Dempster, D.N. (2011). Biochar: Still much to uncover about its agronomic impact.

New Frontiers in Agriculture: The Official Journal of the Western Australian

No-till Farmers Association. June edition, pp 53-55.

Dempster, D.N., Gleeson, D.B., Abbott, L.K., Murphy, D.V. (2011). Biochar and the

nitrogen cycle: My investigation. Biochar and Soil Biology Workshop,

Mingenew (25th), August 2011. Oral presentation by D. Dempster.

Dempster, D.N., Gleeson, D.B., Abbott, L.K., Murphy, D.V. (2011). Biochar and the

nitrogen cycle: My investigation. Biochar and Soil Biology Workshop,

Dandaragan (26th), August 2011. Oral presentation by D. Dempster.

Dempster, D.N. (2012) Biochar and the nitrogen cycle: Unravelling the interaction.

Soil and Water Seminar. March 2012. Oral presentation by D. Dempster.

Dempster, D.N. (2012) Biochar and the nitrogen cycle: Unravelling the interaction.

Postgraduate Showcase; UWA Institute of Agriculture. June 2012. Oral

presentation by D. Dempster.

xvi

List of Abbreviations

xvii


ANOVA

AOB

BC

C

CO2

CO32-

CLPP

DMRT

DOC

DON

EC

FAA

HMW

IRGA

K

LMW

MBC

MBN

MRT

N

NA

analysis of variance

ammonia oxidising bacteria

black carbon

carbon

carbon dioxide

carbonate

community level physiological profile

Duncan’s multiple range test

dissolved organic carbon

dissolved organic nitrogen

electrical conductivity

free amino acids

high molecular weight

infra-red gas analyser

potassium

low molecular weight

microbial biomass carbon

microbial biomass nitrogen

mean residence time

nitrogen

not assessed


xviii

NH3

NH4+

NO3-

NS

P

PAHs

PCR

PERMANOVA

SIR

SOM

TN

T-RFLP

T-RFs

VM

VOCs

WA

WHC

ammonia

ammonium

nitrate

not significant

phosphorus

polycyclic aromatic hydrocarbons

polymerase chain reaction

permutational analysis of variance

substrate induced respiration

soil organic matter

total nitrogen

terminal-restriction fragment length polymorphism

terminal-restriction fragments

volatile matter

volatile organic compounds

Western Australia

water holding capacity

xix

xx

Abstract

xxi

Abstract

Methods of sequestering carbon (C) are being investigated to mitigate the current

trend of increasing atmospheric carbon dioxide concentration. Furthermore, since

the onset of intensive farming practices soil organic matter (SOM) content has

declined in many soil types throughout the world. The application of pyrolysed

organic matter, or biochar, to soil can maintain higher soil C. The application of

biochar to soil has also been shown to potentially improve soil fertility and enhance

yields of crops such as maize, wheat and peanuts. However, biochar amendment

changes many soil properties but the mechanism by which biochar alters soil

processes and plant growth is not always clear. This thesis focuses primarily on

those changes that are directly linked to the soil nitrogen (N) cycle.

Biochar has been reported to have a large nutrient and water retention capacity;

which may cause decreases in N leaching with its application to soil. Biochar can

contain labile C and is also highly porous, potentially providing a substrate for

microbial utilisation and a habitat for microbial colonisation. Biochar addition to

soil can increase microbial abundance and activity. Previous research has

hypothesised that this may be responsible for enhanced N mineralisation rates that

contribute to the demonstrated improvements of N fertiliser use efficiency (up to

90%) in glasshouse-based agronomic research.

The sandy soils that are widespread throughout the broadacre agricultural region of

Western Australia (WA) naturally have low water (2-10% v/v) and nutrient

Abstract

xxii

retention capacities and low SOM content. Thus the sandy soils of WA are an

environment in which biochar amendment is likely to influence soil N cycling. This

provided the premise for the investigation into how biochar interacts with the soil N

cycle (mineralisation, nitrification, immobilisation, N sorption, leaching, and plant

N uptake) in coarse-textured soils of Western Australian broadacre agriculture.

The influence of Jarrah (Eucalyptus marginata Donn ex Sm.) biochar on N

mineralisation and nitrification in a Tenosol (95% sand; 3.5% clay) was determined

in a glasshouse experiment combined with subsequent laboratory incubations. Net

N mineralisation and nitrification were highest in the control soil and significantly

decreased with increasing addition of biochar (Chapter 2). These decreases could

not be attributed to immobilisation of N, because microbial biomass N did not

change with biochar addition. Decreased mineralisation and nitrification was also

not caused by the sorption of amino acids, or ammonium (NH4+) to the biochar. It is

possible that toxic compounds (for example benzene) present on biochar might have

inhibited N mineralisation; however such compounds were not detected.

Biochar has been hypothesised as an alternative soil amendment (historically clay

has been used) for coarse textured soils to decrease N leaching. To test this

hypothesis Jarrah biochar (25 t ha-1) was added to a Sodosol (94% sand, 5% clay at

0-10 cm depth) in a glasshouse lysimeter experiment to determine its influence on

N leaching. Biochar amendment decreased leaching of NH4+ and nitrate (NO3

-) by

14% and 28%, respectively (Chapter 3). Sub-soil clay addition at 25 t ha-1 also

Abstract

xxiii

caused similar decreases in NH4+ and NO3

- leaching (18% and 16% respectively)

despite the large differences between their sorption capacities, as determined by

sorption isotherms. The greater decrease in NO3- leaching with biochar addition was

likely to be due to a decrease in nitrification (observed in Chapter 2). Therefore

sorption is unlikely to be the primary mechanism causing decreases in inorganic N

leaching with biochar addition. In the absence of other evidence it may be possible

that improved water holding capacity as a result of biochar addition might have

decreased inorganic N leaching.

To investigate whether organic N sorption to an alternate biochar type similarly

influenced mineralisation of organic N, wheat (Triticum aestivum L.) residue

biochar was applied to a Kandosol (93% sand) at 4 t ha-1. In a laboratory incubation,

the mineralisation rate of 14C-labelled peptides and amino acids did not change with

biochar addition (Chapter 4). Thus, the sorption of low molecular weight (LMW)

organic N to biochar is unlikely to influence N mineralisation upon addition to

sandy soil and the addition of wheat residue biochar at 4 t ha-1 is unlikely to

influence intrinsic dissolved organic N turnover.

To assess the net effect of the interaction between biochar and the soil N cycle

wheat residue biochar was applied to a Kandosol (93% sand) at 4 t ha-1 and soil

(inorganic N and microbial biomass N) and plant measurements (biomass, yield and

N) were taken over the growing season. Biochar decreased wheat yield from 2.06 t

ha-1 to 1.8 t ha-1 where biochar was spread and significantly decreased to 1.7 t ha-1

Abstract

xxiv

where biochar was banded (Chapter 5). Of the sampling periods, only one (terminal

spikelet) showed any significant differences in soil NH4+ or NO3

- with biochar

application. In the first year of application, wheat residue biochar was an ineffective

amendment for improving N cycling and synergistically interacting with N

fertiliser.

Although biochar decreased N leaching in a high rainfall controlled environment,

there were no other benefits observed from the addition of two types of biochar to

the tested coarse-textured soils used for broadacre agriculture in WA. This may be

due to the low rates of application possible in this agro-ecosystem. Results may

vary with contrasting biochar types, different soil types or higher application rates

of biochar. The sorption of low molecular weight N to biochar is unlikely to

influence N mineralisation, nitrification or be the main factor decreasing N leaching

with biochar addition to coarse-textured soil. Overall biochar is unlikely to

influence N cycling for the improvement of crop productivity on coarse-textured

soils in broadacre agriculture of WA.

General Introduction and Literature Review Chapter 1

1

Chapter 1

General Introduction and Literature Review

1.1 General Introduction

This thesis examines how biochar amendment to coarse-textured soils used for

broadacre agriculture in Western Australia (WA) impacts the terrestrial nitrogen (N)

cycle. This chapter introduces both broadacre agriculture in WA and biochar, and

examines research assessing the interaction between soil N cycling and biochar

application to soil. The examined aspects of the N cycle are: sorption of N; desorption

of N; leaching; mineralisation; nitrification; and plant N uptake. Where possible, this

review extrapolates current research to a broadacre agricultural context for WA.

1.2 Western Australian Agriculture

Broadacre agriculture, within WA and other parts of Australia, is characterised by

large scale, low productivity and infertile soils. From 2000-2001 to 2010-2011 the

total area of winter crop in Australia averaged 21.4 m ha, of which WA contributed

7.5 m ha (ABARES, 2011). In this agro-ecosystem, wheat is the dominant cropping

enterprise, occupying 12.6 and 4.6 m ha in Australia and WA, respectively

(ABARES, 2011). Broadacre agriculture within WA occurs in a mediterranean-type

climate (Rovira, 1992), with average winter-dominant rainfall ranging from less than

300 mm to around 500 mm (1996-2005 records; Bureau of Meteorology, 2012).

Average maximum and minimum temperatures range from approximately 34 and


2

17˚C in January and 17 and 3 ˚C in July (Bureau of Meteorology, 2012). Wheat

yields average 1.5 t ha-1 in this area (2000-2001 to 2010-2011; ABARES, 2011).

Soil types within the broadacre agricultural region of WA vary from deep sands and

texture contrast (duplex) soils, to loamy earths and cracking clays (Schoknecht, 2002)

but are dominated by coarse-textured top soils, which occupy over 14.2 m ha in the

region (Tennant et al., 1992). The depth of the coarse textured A horizon varies from

5 cm to over 100 cm, and generally has a low clay content (often less than 5%;

Tennant et al., 1992). Consequently, water holding capacities for these soil types are

low, ranging from 20-50 mm m-1 for a medium sand to 90-120 mm m-1 for a loamy

sand (Tennant et al., 1992). Soils within this region tend to be acidic; a recent survey

found that over 80% of top soil (0-10 cm) samples had a pH (CaCl2) of less than 5.5

(Gazey & Andrew, 2009). Low organic carbon is also common, typical values may

range from 1.5-2% (Hoyle et al., 2011), but varies with farming practices (Murphy et

al., 2011).

Due to the inherent low fertility of soils within this region, fertilisers, such as N and

phosphorus (P), are an important component of agricultural production. For crop N

nutrition, 0-80 kg N ha-1 is applied depending on the expected rainfall (Anderson &

Hoyle, 1999), but typically 40-80 kg N ha-1 for an expected 2 t ha-1 wheat (Triticum

aestivum L.) yield (Hoyle & Murphy, 2011). For economic reasons, many farmers

seek methods to improve productivity (Kingwell & Pannell, 2005) and decrease input

costs (Keating & Carberry, 2010) by increasing N fertiliser use efficiency (Chen et

al., 2008).


3

Within Australia, estimates of N fertiliser use efficiency in wheat production vary

from 0 to 43 kg grain for each kilogram of N applied with a recovery rate of applied

N ranging from 6-78% (Mason et al., 1972; McDonald, 1989; Ladha et al., 2005).

Generally, this is greater than the global estimate of 33% N fertiliser use efficiency

for cereal production (Raun & Johnson, 1999).

To improve N fertiliser use efficiency in crop production, the synchrony (both in

terms of time and location) between crop N demand and N supply (mineralised or

fertiliser) must be improved (Chen et al., 2009a). During the dry summer fallow

period, episodic rainfall events enable N to be mineralised (McNeill et al., 1998),

which, provided it is not leached, could be available for plant N uptake during the

following growing season. At the commencement of the growing season, plant (crop

and pasture) N demand is small and plant roots have a low surface area for N capture.

Coarse-textured soils are particularly prone to N leaching and acidification (Dolling

& Porter, 1994; Dolling et al., 1994; Anderson et al., 1998). Nitrate (NO3-) leaching

has been reported to range from 17-59 kg N ha-1 yr-1, with greater NO3- leaching

under crops (40-59 kg N ha-1 year-1) than pastures (17-28 kg N ha-1 yr-1; Anderson et

al., 1998). Fine-textured amendments, such as ‘red mud’ bauxite residue or subsoil

clay from the B horizon of Sodosols and Chromosols, can ameliorate nutrient

leaching (Vlahos et al., 1989) and water repellence in coarse-textured soils

(McKissock et al., 2000; McKissock et al., 2002). Biochar addition can also decrease

N leaching (Lehmann et al., 2003; Ding et al., 2010), thus may aid increased N

fertiliser use efficiency on coarse-textured soils of WA.


4

1.3 Biochar

1.3.1 Introduction to biochar

Current concerns about climatic change, have provided impetus for investigations

into a range of carbon (C) sequestration options such as increasing organic C in soils

(Jackson & Schlesinger, 2004). The organic matter content of many soil types

globally has declined within recent history (Dalal & Chan, 2001; Bellamy et al.,

2005). In Australia, losses of organic C can be in the range of 0.04-1.21 g C g-1 soil

yr-1, varying with soil type (greater losses in coarse textured soils; Dalal & Mayer,

1986). Consequences of declining soil organic matter (SOM) include increases in

bulk density (Dalal & Chan, 2001) and decreased energy for heterotrophic soil

microbes to catalyse nutrient transformations (Rosswall, 1982; Killham, 2006).

One method to increase SOM is by adding biochar to soil (Lehmann et al., 2006;

Sohi et al., 2010). Biochar has been defined as thermally decomposed (or pyrolysed)

organic matter, created by heating to temperatures from 250°C to 700°C in low

oxygen conditions (Lehmann & Joseph, 2009). The application of biochar to soil

provides a method to sequester C (Lehmann et al., 2006; Ogawa, et al., 2006; Sohi et

al., 2010) because biochar contains a large portion of aromatic C (Schmidt & Noack,

2000) which can be recalcitrant for thousands of years (Preston & Schmidt, 2006).

The length of time over which biochar C is stored in soil and the influence of biochar

on soil fertility vary with the properties and characteristics of the biochar (Singh et

al., 2010). Some properties of biochar that enable it to alter soil fertility are porosity


5

(Fig. 1.1), surface area (Downie et al., 2009; Table 1.1), pH (Laird, 2008; Sohi et al.,

2010; Table 1.1), bulk density (Laird, 2008; Sohi et al., 2010; Table 1.1), surface

functional groups (Laird, 2008; Clough & Condron, 2010; Sohi et al., 2010) and the

sizes of the fractions of recalcitrant biochar C and mineral ash (Baldock & Smernik,

2002; Bruun et al., 2008; Singh et al., 2010; Table 1.1). Biochar production variables

(e.g. pyrolysis temperature and pyrolysis time) influence biochar properties like the

size of the recalcitrant fraction of biochar C (Baldock & Smernik, 2002; Bruun et al.,

2008; Singh et al., 2010), the functional groups present on the surface of the biochar

(Boehm, 1994; Antal & Grønli, 2003) and the surface area and porosity of the biochar

(Antal & Grønli, 2003; Bornemann et al., 2007). Consequently, biochar is inherently

heterogeneous.

Changes in all production variables such as feedstock (type, water content, particle

size), peak pyrolysis temperature, type of pyrolysis equipment (batch or continuous)

and length of pyrolysis will result in biochar with varying characteristics (Antal &

Grønli, 2003). Increasing peak pyrolysis temperature will increase C concentration of

the biochar (Antal & Grønli, 2003; Table 1.1), altering its influence on C

sequestration and soil fertility (Singh et al., 2010). Contrasting feedstocks will

produce biochars with different surface areas, and hence varying sorption capacity

(Bornemann et al., 2007). The effect of biochar also varies with soil type and crop

type (Van Zwieten et al., 2010b). Therefore, to isolate the mechanisms controlling

changes to soil processes induced by biochar addition, it is important to characterise

experimental biochars. Seven properties for characterisation of biochars have been


6

proposed by Sohi et al. (2010): pH; the content of volatile compounds; ash content;

water holding capacity (WHC); bulk density; pore volume; and specific surface area.

Consistent characterisation of biochar could aid with determining which biochar

characteristics influence soil fertility across a range of ecosystems. The goal of

consistent and thorough characterisation of biochar could be to predict the agronomic

effectiveness of each biochar type as an amendment for varying soil or crop types.

Fig. 1.1: A scanning electron microscope illustrating the porosity of biochar. Image

source: D. Dempster and P. Clode.

Tab

le 1

.1:

Sele

cted

pro

per

ties

of a

ra

nge

of

bioc

har

s. *

No

te:

pH

fo

r th

e g

reenw

aste

bio

char

wa

s m

eas

ure

d in

Ca

Cl

2.

Fe

ed

sto

ck

Pyr

oly

sis

Te

mp

erat

ure

(°C

)

pH

(H2O

)

Bul

k

De

nsity

(g c

m-3)

Tot

al C

(%)

Tot

al N

(%)

Su

rfa

ce

Are

a

(m2 g

-1)

Vo

latil

e

Co

nte

nt

(g k

g-1)

Ash

Co

nte

nt

(g k

g-1)

Re

fere

nce

Oil

Ma

llee

- 8

.4

- -

- 8

1 1

6 5

37

Sol

aim

an e

t al

., 2

010

Mix

ed

Ha

rdw

oo

d 4

50

9.7

0

.35

76

0.6

9 3

9 -

- Jo

nes

et

al.,

201

0

Jarr

ah

600

8

.5

0.4

5 7

8 0

.38

4 -

- Jo

nes

et

al.,

201

0

Ba

mb

oo

600

8

.1

0.7

5 6

8 0

.87

330

- -

Din

g et

al.,

20

10

Ra

diat

a P

ine

300

5

.7

- 6

2 0

.04

21

- -

Ta

ghiz

ad

eh-T

oos

i et

al.,

201

2

Ra

diat

a P

ine

500

6

.6

- 8

3 0

.22

56

- -

Ta

ghiz

ad

eh-T

oos

i et

al.,

201

2

Ma

cada

mia

nu

t sh

ell

6

40

8.2

-

89

0.4

5 -

63

42

De

enik

et

al.,

20

10

Ma

cada

mia

nu

t sh

ell

4

30

5.7

-

85

0.4

5 -

22

3 D

een

ik e

t al

., 2

01

0

Gre

en

wa

ste

600

7

.5*

-

78

0.1

4 40

9 -

- V

an

Zw

iete

n et al

., 2

010

a

Gre

en

wa

ste

350

4

.9*

- 6

2 0

.21

- -

- V

an

Zw

iete

n et al

., 2

010

c

Gre

en

wa

ste

550

7

.3*

- 7

5 0

.24

- -

- V

an

Zw

iete

n et al

., 2

010

c

Sw

itch

gra

ss 2

50

9.7

-

52

1.6

0 -

40

0 -

Sm

ith e

t al

., 2

010

Pou

ltry

Litt

er

400

9

.2

- 4

3 0

.51

- -

34

6 S

ingh

et al

., 2

010

Syd

ne

y B

lue

Gu

m 4

00

6.9

-

70

0.0

2 -

- 3

5 S

ingh

et al

., 2

010

Syd

ne

y B

lue

Gu

m 5

50

8.8

-

84

0.0

3 -

- 3

3 S

ingh

et al

., 2

010


8

1.3.2 Biochar in the agricultural system: focus on broadacre agriculture in

Western Australia

Many renewable feedstocks can be utilised as biochar sources such as urban

greenwaste (Chan et al., 2007), sewerage sludge (Bridle & Pritchard, 2003),

agricultural wastes such as manures (Chan et al., 2008), wheat (Triticum aestivum L.)

residue (Chun et al., 2004) and silvicultural wastes such as oil mallee (Eucalyptus

sp.) residue (Solaiman et al., 2010). The most widely available feedstocks for biochar

production in WA are cereal crop residues and Eucalyptus biomass. Biochar

production from Eucalyptus residue may provide another income source for oil

mallees planted to ameliorate salinity and wind erosion. Further, more Eucalyptus

plantations in the wheatbelt of WA could provide some of the reforestation required

to return the rainfall of the south-west of WA to its long term average (Pitman et al.,

2004).

Practical rates of application of biochar within broadacre agriculture of WA are likely

to be less than 5 t ha-1 (Blackwell et al., 2010). This is low compared to other agro-

ecosystems which may use application rates of 25-50 t ha-1 (e.g. Jones et al., 2010).

The lower rates of biochar application in broadacre WA is due to a combination of

factors such as lower net primary productivity causing lower biochar production

volumes and larger acreages of land causing larger distances for transport and higher

transport costs. Due to the lower rates of application in broadacre agriculture within

WA, it is unknown whether the mechanisms suggested in other previous research

(examined in following sections) are applicable to the target agro-ecosystem.


9

1.3.3 Soil biological fertility changes with biochar addition

A meta-analysis examining data assessing the impact of a variety of biochar types

added to a range of soil types on many cultivated plants found that, on average,

biochar improved plant productivity by 10% and crop yield by 5% (Jeffrey et al.,

2011). The increases in crop yield and plant productivity are explained by many

changes in soil physical, chemical and biological properties induced by biochar

addition to soil (Glaser et al., 2002; Atkinson et al., 2010; Clough & Condron, 2010;

Sohi et al., 2010; Lehmann et al., 2011).

In a comprehensive review of the soil biological changes induced by biochar addition

to soil Lehmann et al. (2011) concluded that in most studies (>20) biochar

amendment increased soil microbial biomass. There are at least six mechanisms by

which biochar application can increase soil microbial abundance: (i) introduction of

nutrients and C; (ii) increasing pH in acidic soils; (iii) adhesion of microbes to pores;

(iv) provision of protection from other biota; (v) protection against desiccation; and

(vi) sorption of toxins (Lehmann et al., 2011). This review will discuss the relevance

of each of these mechanisms for broadacre agriculture in WA.

Soils within WA generally have low labile C content (e.g. Holmes et al., 2011) which

restricts the size of the heterotrophic microbial population. Labile C contained in

biochar may partially alleviate microbial substrate deficiencies increasing microbial

abundance (Lehmann et al., 2011), but the longevity of this influence is unclear. The

influence on microbial population is will vary with the quality and quantity of


10

nutrients contained within the biochar, the intrinsic soil nutrition and the species of

microbe (Lehmann et al. 2011). Biochar amendment may also increase microbial

abundance by increasing the nutrient retention capacity of the coarse-textured soils of

WA.

Increasing the pH of agricultural soil can have a marked effect on microbial

populations in terms of an increase in biomass and activity (Aciego Pietri & Brookes,

2008). As acidic soils dominate the broadacre agricultural region of WA (e.g.

Tennant et al., 1992), increasing soil pH through biochar amendment may increase

microbial biomass and activity similarly. The magnitude of the increase in soil pH

with biochar amendment is variable and previous studies have not determined the

proportion of the increased microbial population due to the changes in pH or the

coinciding increased labile C.

The adhesion of microbes to biochar may prevent leaching of microbes through the

soil profile, enhancing microbial abundance (Lehmann et al., 2011). The adhesion of

microbes to biochar may occur via electrostatic forces, hydrophobic interactions or

covalent bonding, causing adsorption of microbes to the biochar surface; alternatively

microbes could become trapped within the biochar matrix (Lehmann et al., 2011).

The biochar matrix could provide a habitat for microbes to avoid grazing from

mesofauna (Wardle et al., 1998). This would require pore sizes larger than the

microbes, such as 0.6 μm for N mineralising bacteria (Strong et al., 1998), but too

narrow for microbial predators to enter. The nature of pore size distribution of


11

biochar has been poorly documented, and is likely to vary with soil type. In soils with

low clay content, such as those in the broadacre agricultural region of WA, these

mechanisms are likely to be expecially relevant.

The protection of microbes from desiccation may not be applicable in broadacre agro-

ecosystems in WA because it is located within a mediterranean-type climate zone,

where drying is a regular occurrence. The only studies in Lehmann et al. (2011)

assessing microbial abundance in a mediterranean-type climate focused on symbiotic

microbes (mycorrhizae; e.g. Blackwell et al., 2010; Solaiman et al., 2010) rather than

free-living microbes. Due to climatic conditions the soil dries to very low water

content (<0.5% w/w) every summer, therefore free-living microbes have adapted to

desiccation. Water contained within biochar may provide microbes with an extended

period of time prior to drying, due to its large surface area and WHC, but desiccation

of microbes through drying will occur in this environment.

Biochar can sorb (adsorb or absorb) compounds that are microbially inhibitive, such

as catechol (Chen et al., 2009b; Kasozi et al., 2011), decreasing their prevalence in

the soil solution, which may increase microbial abundance (Lehmann et al., 2011).

Conventional agricultural practices use pesticides that can be microbially inhibitive,

such as tebuconazole (Muñoz Leoz et al., 2011) and diuron (Prado & Airoldi, 2001).

Whilst biochar can sorb some pesticides, such as simazine (Jones et al., 2010),

atrazine and trifluralin (Nag et al., 2010), decreasing their bio-availability to plants

(Bornemann et al., 2007; Nag et al., 2011); further research is required to determine


12

how biochar amendment influences the net effect of the pesticide on microbial

abundance.

The increased soil microbial biomass induced by biochar addition has a variable

effect on specific microbiological phyla and families (Lehmann et al., 2011;

Anderson et al., 2011). Biochar addition can increase the abundance of

Actinobacteria (Khodadad et al., 2011), Gemmatimonadetes (Khodadad et al., 2011),

Zygomycota (Lehmann et al., 2011) and Glomeromycota (Lehmann et al., 2011).

Similarly, pine biochar increased the abundance of Bradyrhizobiaceae,

Hyphomicrobiaceae, Streptosporangineae and Thermomonosporaceae but decreased

the abundance of Nitrosomonadaceae (nitrosovibrio), Streptomycetaceae and

Micromonosporaceae (Anderson et al., 2011). However in general, less genetically

diverse microbial communities have been found in biochar amended soils (Lehmann

et al., 2011; Khodadad et al., 2011). Further research should determine whether

changes in process rates (e.g. nitrification) induced by biochar amendment correlate

with changes in the microbial population that exhibit the genetic capabilibility of

regulating the changed process rate (e.g. the ammonia oxidiser population). Research

should also correlate changes in both the microbial community and the process rate

with the physical and chemical properties the biochar. This may facilitate the

prediction of microbial population changes and process rate changes based on the

properties of the biochar.


13

1.4 The Nitrogen Cycle

The N cycle is integral for all living organisms. For plants, N is contained within

compounds such as amino acids and proteins. In conventional agricultural systems,

plants have access to two sources of N, biologically derived N and inorganic fertiliser

N. Although many conventional farming systems only apply N in the form of

inorganic fertiliser (e.g. ammonium (NH4+)), the contribution of biologically

mediated plant available N can be up to 90% in Australian soils (Angus, 2001), which

constrasts with temperate systems where approximately 50% of plant N uptake is

from fertilisers (Jenkinson, 2001).

Biochar addition can alter a number of N cycling processes (Clough & Condron,

2010) but due to differences in biochar type, soil type and climatic conditions the

impact of biochar in many agro-ecosystems is unclear. This thesis focuses on the

impacts of biochar addition to soil on the N cycle for broadacre agriculture in WA.

Thus the following sections of this review synthesise research examining how

biochar addition to soil affects N sorption, desorption, N leaching, N immobilisation,

N mineralisation, nitrification, and plant N uptake and where possible extrapolate

results to a WA broadacre agricultural context (Fig. 1.2; Table 1.2).


14

Fig. 1.2 (page 15): A diagram describing how biochar addition to soil can influence

the N cycling processes assessed in this thesis. Note: 1 – Lehmann et al., 2003; 2–

Deenik et al., 2010; 3– Chan et al., 2008; 4– Chan et al., 2007; 5– Prendergast-

Miller et al., 2011; 6– Ding et al., 2010; 7 – Laird et al., 2010b; 8– Berglund et al.,

2004; 9 – DeLuca et al., 2006; 10 – Ball et al., 2010; 11 – Clough & Condron, 2010;

12– Lehmann et al., 2011; 13– Clough et al., 2010; 14– Spokas et al., 2010; 15–

Spokas et al., 2012.


15

Biochar addition to soil may influence Plant N uptake

by: Increasing immobilisation of N1;2; Increasing supply of P and K3;4;

Localising NO3- in rhizosphere biochar5.

NO3-

Biochar addition to soil may influence N leaching

by: Increasing sorption of N1;6;7;

Increasing immobilisation of N1;7; Altering mineralisation rates7;

Increasing soil water holding capacity1. Leached NH4+ Leached NO3

-

Biochar addition to soil may influence Nitrification by:

Sorbing inhibitory compounds8;9; Introducing inhibitory compounds11;13;

Altering immobilisation of N9;10; Emitting ethylene14.

Biochar addition to soil may influence Sorption

by: Adding sorptive functional groups15;

Increasing soil surface area.

Biochar addition to soil may influence Immobilisation

by: Adding labile C compounds1;2;12.

Biochar addition to soil may influence Mineralisation

by: Introducing inhibitory compounds11;13.

Sorbed N

Nitrification

Sorption

Immobilisation

Sorption

Immobilisation

Immobilised N

NH4+

Mineralisation

Organic Nitrogen

Plant N uptake Plant N uptake

Leaching Leaching

Tab

le 1

.2:

Su

mm

ary

of s

ome c

ha

nges

in N

cyc

ling

pro

cess

es in

duce

d b

y th

e a

ddi

tion

of b

ioch

ar

to s

oil.

Ital

icis

ed

sta

tem

ents

are

hyp

oth

ese

s (e

ither

hyp

oth

eses

for

cha

nge

s in

th

e pr

oce

ss r

ate o

r h

ypo

thes

ised

mech

anis

ms)

sug

gest

ed

by t

he c

orr

esp

on

ding

refe

ren

ce.

N c

ycle

pro

cess

es

Net

pro

cess

effe

ct

Me

cha

nis

ms

aff

ect

ing

pro

cess

ra

tes

Re

fere

nce

(s)

- M

iner

alis

atio

n an

d

nitr

ifica

tion

Incr

ea

se: N

itrifi

catio

n ra

tes

we

re a

bo

ut 5

times

hig

her

in f

ore

st (

low

nitr

ifica

tion

act

ivity

) so

ils.

Ch

arco

al s

orb

ed

phe

nol

ic c

ompo

und

s ca

usin

g

eith

er

incr

ease

d ni

trifi

catio

n a

nd/

or d

ecre

ase

d

imm

obili

satio

n.

De

Luca

et a

l., 2

00

6

D

ecr

ease

: At t

he

en

d o

f a 1

4 d

ay

incu

batio

n,

NH

4+ co

nte

nt w

as

25%

low

er

with

a lo

w

vola

tile

mat

ter

(VM

) b

ioch

ar

and

75%

low

er

with

a h

igh

VM

bio

cha

r

Hig

he

r vo

latil

e co

nte

nt b

ioch

ars

ad

de

d p

hen

olic

com

pou

nd

s to

soi

l, re

sulti

ng in

imm

obi

lisa

tion,

de

crea

sing

min

era

lisa

tion.

De

enik

et

al.,

20

10

D

ecr

ease

B

ioch

ar a

me

ndm

ent t

o so

il ca

uses

eth

yle

ne

em

issi

on

whi

ch c

an

dec

reas

e ni

trifi

catio

n.

Sp

oka

s et

al.,

201

0

- Im

mo

bilis

atio

n In

cre

ase

La

bile

C c

onte

nt in

bio

char

is m

icro

bial

ly

ava

ilabl

e; c

au

sin

g im

mo

bilis

atio

n o

f N.

Leh

ma

nn

et a

l., 2

003;

Lair

d e

t al

., 2

010

b;

De

enik

et

al.,

20

10

- P

lan

t up

take

In

cre

ase

d N

fe

rtili

ser

use

effi

cien

cy

Pro

visi

on o

f o

the

r n

utri

ent

s, s

uch

as P

an

d K

,

lea

din

g to

N b

ein

g a

mor

e lim

itin

g nu

trie

nt.

Ch

an

et a

l., 2

00

7;

Ch

an e

t al

., 2

008

In

cre

ase

d N

fe

rtili

ser

use

effi

cien

cy

Loca

lisat

ion

of N

O 3- w

ithin

rhi

zosp

here

bio

cha

r P

rend

erga

st-M

iller

et

al.,

20

11

- Le

ach

ing

De

crea

se -

Cu

mu

lativ

e N

H4+

lea

chin

g

de

cre

ased

by

25%

with

ch

arco

al a

dditi

on

3

we

eks

aft

er fe

rtili

satio

n

Sor

ptio

n of

NH 4

+

Leh

ma

nn

et a

l., 2

003

In

cre

ased

wa

ter

hol

din

g ca

pa

city

Le

hm

an

n et

al.,

200

3

D

ecr

ease

S

orp

tion

of N

O 3-

In

cre

ase

- O

ver

a 4

5 w

eek

incu

bat

ion,

bio

cha

r a

t th

e gr

eat

est a

dded

ap

plic

atio

n

rate

(20

g k

g-1 d

ry s

oil)

incr

ea

sed

NO 3-

lea

chin

g b

y 2

5%

Incr

eas

ed o

rgan

ic N

min

era

lisa

tion

Lair

d e

t al

., 2

010

b

D

ecr

ease

- O

ver

a 45

we

ek

incu

batio

n,

bio

cha

r a

ppl

ica

tion

with

man

ure

de

cre

ase

d

NO

3- leac

hin

g co

mp

are

d to

no

bioc

har

cont

rols

Bio

char

sor

be

d N

H 4+ a

nd

solu

ble

org

ani

c

com

pou

nd

s, in

hibi

ting

org

anic

N m

ine

ralis

atio

n

Lair

d e

t al

., 2

010

b

- S

orp

tion

NH 4

+

Incr

ea

se

Larg

e su

rfac

e ar

ea,

va

riabl

e s

urf

ace

ch

arg

e

- S

orp

tion

NO 3

- In

cre

ase

La

rge

surf

ace

are

a, v

aria

ble

su

rfa

ce c

har

ge

- D

eso

rptio

n S

mal

l de

cre

ase

- 8

0%

of t

he N

H3 so

rbe

d to

bio

cha

r in

a g

lass

hou

se e

xpe

rim

ent

wa

s

bio

ava

ilabl

e

Larg

e s

urfa

ce c

harg

e T

agh

iza

de

h-T

oos

i et

al.,

20

12


18

1.5 Sorption of nitrogen to biochar

Biochar, charcoal and activated C are used in industrial, medicinal and filtration

processes to sorb impurities. Their large surface area and variety of surface

functional groups enable them to sorb a range of chemicals, such as pesticides

(Yang et al., 2004; Bornemann et al., 2007), aromatic hydrocarbons (Zhu &

Pignatello, 2005; Accardi-Dey & Gschwend, 2003), metals (Ma & Rate, 2007)

nitrogenous ions such as NH4+ (Ding et al., 2010), ammonia (NH3; Kastner et al.,

2009; Taghizadeh-Toosi et al., 2012) and NO3- (Mizuta et al., 2004). The role that

sorption interactions play in altering N bioavailability is unclear because N sorption

to biochar after its addition to soil has not been assessed.

1.5.1 Mechanisms of sorption and properties influencing sorption to biochar

Biochar can both accept and donate electrons in pi bonds and electrostatic

attractions (Keiluwait & Kleber, 2009). This is because it can be comprised

primarily of aromatic C rings located within graphene sheets and can exhibit

varying charge (Schmidt & Noack, 2000). Functional groups, such as carboxyl

groups, are located on the edge of graphene sheets (Schmidt & Noack, 2000).

Acidic functional groups, and areas of the graphene sheet with impurites are

electron rich, and react with pi bond acceptors (Zhu & Pignatello, 2005). Negatively

charged functional groups enable NH3 sorption (Seredych & Bandosz, 2007;

Kastner et al., 2009; Clough & Condron, 2010; Spokas et al., 2012) and increase

CEC upon addition to soil (Boehm, 1994; Liang et al., 2006; Clough & Condron,

2010). By contrast, basic functional groups present on biochar react with pi bond


19

donors anabling anion exchange capacity (Boehm, 1994). As the pH of biochar

increases, its negative charge increases proportionally (Cheng et al., 2008) and

although biochar pH will be indicative of the general charge, there may be some

different functional groups of varying charge within the biochar.

The functional groups on the biochar are influenced by a number of factors

including pyrolysis temperature, feedstock and extent of oxidation. Greater

pyrolysis temperatures increase the C content of biochars, decreasing acidic

functional groups and increasing basic functional groups (Antal & Grønli, 2003;

Chun et al., 2004). Greater pyrolysis temperatures also increase the hydrophobicity

of biochar which restricts access of water soluble molecules like NH4+ (Seredych &

Bandosz, 2007) but enables hydrophobic organic matter sorption (Kasozi et al.,

2010). The feedstock used to create the biochar shall influence which functional

groups are present on the biochar surface. Over time, biochar oxidises (Cheng et al.,

2006; Cheng et al., 2008). Through changes in surface functional groups, oxidation

decreases surface positive charge and increases negative surface charge,

correspondingly increasing cation retention, but decreasing anion retention (Cheng

et al., 2008). The main reactive functional groups on aged biochar are carboxyl and

phenol groups (Liang et al., 2006). The extent of oxidation is highly correlated with

temperature (Cheng et al., 2006; Cheng et al., 2008) but the time required for

oxidation to occur is unclear.


20

The total surface area of biochar influences its sorption potential by providing a

larger area for reactions to occur. Although the surface area of biochar varies with

biochar type (ranging from 4-605 m2 g-1; Table 1.1; Table 1.3) it is generally greater

than quartz based sand (~1.3 m2 g-1) or kaolin (4-88 m2 g-1; Hart et al., 2002). The

feedstock used to produce the biochar influences the surface area of biochar (Table

1.3). Analysis method also influences surface area values; Bornemann et al. (2007)

used N2 adsorption on a BET surface area analyser. The surface area of biochar

generally increases with peak pyrolysis temperature (Table 1.3) to a temperature of

around 500-600°C (Antal & Grønli, 2003).

Table 1.3: Specifice surface areas of biochar produced from two feedstocks at three

pyrolysis temperatures; data from Bornemann et al. (2007).

Biochar Feedstock Pyrolysis Temperature

(°C)

Specific Surface Area

(m2 g-1)

Red Gum (Eucalyptus camaldulensis

Dehnh.)

250 8

450 34

850 605

Phalaris (Phalaris tuberosa L.) 250 4

450 20

850 371


21

1.5.2 The role of nitrogen sorption to biochar in ecosystem processes

The sorption of N to biochar has been suggested to play a role in ecosystem

processes (Lehmann et al., 2003), yet has not actually been tested. As discussed, it

is clear that biochar can sorb a range of compounds and that the introduction of

biochar to soil has the potential to increase nitrogen retention through electrostatic

adsorption (Keiluweit & Kleber, 2009). However the quantity of sorption that does

occur is unclear and thus, the role that N sorption to biochar actually plays within

ecosystems, whether broadacre agriculture in WA or not, is currently unknown.

1.6 Desorption

Once sorbed to biochar, for N to be bioavailable it must be desorbed. One study

demonstrated that a portion of sorbed N is bioavailable because up to 45% of N

sorbed to biochar was recovered as soil N (either organic or inorganic) or and up to

35% of sorbed N was recovered as plant N (Taghizadeh-Toosi et al., 2012). The

remaining sorbed N was not recovered. By contrast, Hina et al. (2010) found that

only 14-27% of sorbed NH4+ could be desorbed by extraction (2M potassium

chloride). It is likely that the prevalence of desorption depends on the surface

functional groups and charge properties of the biochar.

In the study of Taghizadeh-Toosi et al. (2012) all NH3 sorbed to the biochar

occurred in vitro. Thus sorption within the agro-ecosystem was not tested. As

desorption can only occur after sorption the potential for N sorption to biochar to

occur after addition to soil should also be assessed.


22

1.7 The influence of biochar on nitrogen leaching

Leaching of inorganic N and dissolved organic N (DON) can contribute to

eutrophication, soil acidification, and agricultural productivity losses. The deep

sands and duplex (or texture contrast) soils that occupy about 60% of the broadacre

agricultural area of WA (Tennant et al., 1992) are prone to leaching due to low

water and nutrient retention capacities (Dolling & Porter, 1994; Dolling et al., 1994;

Ridley et al., 2001). Nitrogen leaching can be decreased by the application of

biochar (Lehmann et al., 2003; Ding et al., 2010; Laird et al., 2010b). The

minimum application rate required to induce a response is unknown, but 0.5% w/w

biochar/soil (Ding et al., 2010) is the lowest applied rate of biochar demonstrated to

decrease N leaching in the published literature. Further, it is also unclear if the rate

of N leaching decreases proportionally with increasing application rate. There are

five main causes of decreased N leaching with biochar application: increased

sorption of nutrients (Lehmann et al., 2003); increased water holding capacity

(Lehmann et al., 2003); changes in organic N mineralisation (Laird et al., 2010b);

changes in nitrification; and changes in immobilisation (Lehmann et al., 2003).

1.7.1 Leaching of nitrogen decreases due to sorption

As discussed previously (Section 1.5), biochar has the ability to sorb N. This has

been hypothesised to be a cause of decreased N leaching with biochar application,

but this hypothesis has not been tested. Consequently, it is unclear how much NO3-,

NH4+ and organic N leaching is decreased due to sorption to biochar. The extent to

which sorption decreases N leaching will change with biochar type and application


23

rate due to varying surface areas and quantity and types of surface functional

groups.

1.7.2 Leaching of nitrogen decreases due to increasing water holding capacity

The application of biochar to coarse-textured soils increases WHC by increasing

total soil surface area, and the density of pores within the soil (Tryon, 1948)

enabling sorption of water. Increasing soil surface area and WHC by applying fine

textured amendents, like bauxite residue mud (the fine fraction of the waste

component from bauxite extraction) can decrease nutrient leaching (Vlahos et al.,

1989). Biochar is likely to decrease nutrient leaching similarly.

1.7.3 Leaching of nitrogen changes due to nitrification and mineralisation

Negatively charged clay electrostatically attracts NH4+. If a greater proportion of N

is found as NO3-, because of increased nitrification, the risk of N leaching increases.

Similarly DON generally exhibits negative charge, thus also is less attracted to clay

than NH4+. Adding biochar to soil can change nitrification and mineralisation rates

(discussed in Sections 1.9 and 1.10) and hence the N form available to leach.

In an Amazonian Ferralsol, Lehmann et al. (2003) found that although NH4+

leaching decreased with biochar addition, NO3- leaching increased. Lehmann et al.

(2003) suggest the change in leaching was due to the increased sorption capacity,

increased WHC or greater microbial immobilisation resulting from biochar

amendment, but the increased NO3 leaching is not explained. However, increased


24

nitrification could explain the results of Lehmann et al., (2003) because it would

result in both increased NO3- leaching and decreased NH4

+ leaching.

In a 45 week leaching experiment using soil (Typic Hapludolls) from the mid-west

of the United States of America, the increased NO3- leaching with the greatest rate

of biochar addition was attributed to increased organic N mineralisation (Laird et

al., 2010b). By contrast, with manure and biochar additions, NO3- leaching

decreased compared to manure additions alone also apparently due, in part, to less

organic N mineralisation (Laird et al., 2010b). With manure addition the decreased

organic N mineralisation was attributed to the sorption of NH4+ and DON to biochar

limiting substrate availability (Laird et al., 2010b). Laird et al. (2010b) disregard

anion retention as an influencing factor, suggesting that active sites on aged biochar

are primarily carboxylic and phenolic groups, as demonstrated in Liang et al.

(2006). The validity of this assumption is not known. Most of the leaching in Laird

et al. (2010b) occurred within the first 12 weeks without manure addition, and

between weeks 11 and 30 with manure addition. The extent of oxidation is highly

correlated with temperature (Cheng et al., 2006; Cheng et al., 2008). This leaching

experiment was conducted at 25°C (Laird et al., 2010b), but Cheng et al. (2006) did

not measure any net change in total acidic functional groups after a four month

incubation at 30°C. Future research should clarify the links between organic N

mineralisation, N leaching and the aging of biochar.


25

1.7.4 Leaching of nitrogen changes due to microbial immobilisation

Microbial immobilisation of N can decrease the total quantity of N available for N

leaching. The manner in which biochar addition to soil changes microbial

immobilisation of N will be discussed in a following section (Section 1.8); this

section will discuss the implications of immobilisation for N leaching. Due to the

high C content and low N content, immobilisation of N has been attributed to

decreased leaching of N (Lehmann et al., 2003; Laird et al., 2010b) yet

immobilisation was not measured in either study for verification.

1.8 Changes to nitrogen immobilisation induced by biochar application

Soil microbes require a certain portion of N for metabolism, cell maintenance and

other functions to maintain their desired C:N ratio (5:1 for bacteria, 15:1 for fungi;

Killham, 2006). For high C substrates, such as wheat residue (C:N ratio of

approximately 100:1) microbial decomposition requires available N, hence

microbial immobilistion of N occurs. Because biochar also contains a large C:N

ratio (>32.5; Table 1.1) many previous studies have attributed decreased N

leaching, or net NH4+ with biochar amendment to microbial immobisation of N (e.g.

Lehmann et al., 2003; Steiner et al., 2008b; Deenik et al., 2010; Laird et al.,

2010b). Despite this the quantity of N immobilised in the microbial biomass is

usually not measured. Literature assessing the change in the nitrogen component of

the microbial biomass could not be found.


26

Work that attributes results to N immobilisation because of widening C:N ratios can

be criticised in two ways. Firstly, it must be the amount of labile carbon that is

assessed, rather than the total carbon. The labile carbon on the biochar is more

likely to directly influence the microbial community, hence for immobilisation it is

more important that the biochar is assessed for a labile C:N ratio, rather than a total

C:N ratio. Secondly, for net immobilisation to occur the microbial biomass must

utilise inorganic N at a faster rate than it is being supplied via mineralisation of

SOM or from external inputs. As stated previously, this has not been reported in

published literature.

There has been assessment of changes in the soil microbial biomass with biochar

additions to soil via either the substrate induced respiration (SIR) method

(Anderson & Domsch, 1978) or the fumigation extraction method (Vance et al.,

1987). Literature mostly states that adding biochar to soil increases the microbial

biomass (Steiner et al., 2008a; Kolb et al., 2009; Lehmann et al., 2011). Caution

must be shown when interpreting the results. If the biochar contains carbonates

(CO32-; e.g. Singh et al., 2010) and is added to an acidic soil, abiotic carbon dioxide

(CO2) evolution may confound results. Despite this many studies have not assessed

biochar for CO32- (e.g. Steiner et al., 2008a). Calculating the microbial biomass

using the fumigation extraction method may require a correction factor to account

for sorption of organic C, released during chloroform induced cell lysis, to biochar

(Liang et al., 2010), but this may not be required for all biochar types (Durenkamp

et al., 2010).


27

This chapter shall not assess the soil biological changes due to biochar addition

further than discussed in Section 1.3.3, except to re-iterate that in a comprehensive

review Lehmann et al. (2011) concluded that in most cases biochar addition has

resulted in greater microbial abundance and that Section 1.3.3 concluded that this is

also likely to occur in the coarse-textured soil of the Western Australian wheatbelt.

If biochar amendment causes immobilisation of N, plant N availability is likely to

decrease. However the magnitude of the change is unclear, but will likely vary with

biochar application rate and type.

1.9 Biochar influences on nitrogen mineralisation

For this chapter, N mineralisation specifically refers to ammonification, the

conversion of organic N to NH4+. Microbes decompose organic matter to obtain the

energy and nutrients required for cell functioning and maintenance (Murphy et al.,

2003). Where the decomposing substrate contains N surplus to the decomposing

microbe requirements, N is released in the form of NH4+. The total production of

NH4+ over time is the gross rate of N mineralisation which contrasts with the net

rate of N mineralisation, which is the change in total NH4+ over time.

There are large deficiencies in our understanding of how biochar influences N

mineralisation because research assessing how biochar influences gross N

mineralisation is lacking. Current understanding of how biochar alters N

mineralisation is based on changes in net N mineralisation. The addition of two

types of macadamia nut shell biochar decreased soil NH4+ (by 25% with a low


28

volatile matter (VM) biochar and 75% with a high VM biochar) when added at 5%

w/w biochar/soil (Deenik et al., 2010). Deenik et al. (2010) attributed the decreased

NH4+ to greater microbial immobilisation of N because soil respiration either

increased or did not change with the addition of biochar; the intimate link between

the C and N cycles suggest that N mineralisation will have either increased or not

changed similarly (Manzoni et al., 2008). Biochar can contain volatile organic

compounds (VOCs) such as acetaldehyde, α-pinene, β-pinene and trans-pinocarveol

(Clough et al., 2010). Α-pinene can increase C mineralisation and decrease net N

mineralisation (Uusitalo et al., 2008). If α-pinene was present on the macadamia nut

shell biochar, decreased N mineralisation and increased CO2 evolution may also

have occurred. This would alter the interpretation of the results of Deenik et al.

(2010) to suggest that decreased N mineralisation may have occurred. As such,

biochar may contain compounds inhibitive to mineralisation and decrease intrinsic

soil N mineralisation rates upon amendment. Thus the influence of VOCs contained

within biochar, and their influence on N mineralisation requires research.

Although biochar may introduce compounds inhibitory to N mineralisation to soil,

biochar may also sorb compounds, such as phenolics that may be inhibitory to

mineralisation (Gundale & DeLuca, 2007). Although such phenolics may not be

present in agricultural soil types (DeLuca et al., 2006), other compounds, such as

pesticides (e.g. tebuconazole and diuron) that inhibit microbial activity (Prado &

Airoldi, 2001; Muñoz Leoz et al., 2011) may be present. Pesticide bio-availability


29

may be lessened with biochar addition (Nag et al., 2011), but whether N

mineralisation is influenced similarly is unclear.

Biochar could also change the N mineralisation rate by sorbing compounds integral

to mineralisation such as enzymes. Sorption of enzymes, like β-glucosidase, to

biochar can decrease their activity (Bailey et al., 2011; Lammirato et al., 2011).

Bailey et al. (2011) showed β-glucosidase, lipase and leucine aminopeptidase

activities decreased upon biochar exposure in vitro, but N-acetylglucosaminidase

activity increased. When biochar was added to three soils types from three different

ecosystems, arable (Ultic Haploxeroll), grassland (Xeric Torripsamment) or

shrubland soil (Xeric Haplocambid), the effect on enzyme activity was highly

variable (Bailey et al., 2011). With biochar addition to the arable soil, lipase activity

doubled; but decreased with biochar addition to the grassland and shrubland soils

(Bailey et al., 2011). Leucine aminopeptidase activity also increased with biochar

addition to the arable soil, but did not change with biochar addition to the grassland

soil, and halved with biochar addition to the shrubland soil (Bailey et al., 2011). As

SOM content of the three soil types used in Bailey et al. (2011) was not presented,

it cannot be assessed how enzyme activity varies with SOM content. Other research

has shown that SOM mineralisation decreases with biochar addition to soil correlate

with increasing SOM content (Keith et al., 2011). Whether the changes in the

activity of enzymes with biochar addition to soil correlate with changes in SOM

content is unknown. Similarly, it is unknown whether sorption of dissolved organic


30

C (DOC) and DON to biochar when added to soil affects mineralisation, and

whether the mineralisation changes correlate with SOM content.

Substrates for N mineralisation may also sorb to biochar restricting their mobility

within the soil solution. The sorption affinity of varying organic matter compounds,

such as humic acids and catechol, is known to vary in vitro (Kasozi et al., 2010),

thus it is also likely to vary upon amendment to soil. Although not determined, the

mineralisation of sorbed organic N could be expected to slow, but not completely

inhibited because the sorption of β-glucosidase to biochar limits its catylitic activity

but does not halt it (Lammirato et al., 2011). This topic, however, requires further

research.

1.10 The influence of biochar on nitrification

In boreal forests, biochar can increase nitrification rates because it may sorb

phenolic compounds (Berglund et al., 2004; DeLuca et al., 2006; Ball et al., 2010).

These volatile monoterpenes can inhibit nitrification through allelopathic inhibition

(Paavolainen et al., 1998). Ammonia monooxygenase activity is inhibited by

phenolic compounds or monoterpenes with a 6 C ring and an unsaturated C-C bond

close to the terminal position (White, 1988; White, 1991; White, 1994). Such

volatile monoterpenes are common in the soils of Ponderosa pine forests. In

contrast to forest soils, nitrification in grassland soils did not increase with biochar

addition possibly due to the higher inherent nitrification rates or lack of phenolic

compounds (DeLuca et al., 2006). In agricultural systems other compounds such as


31

pesticides such as tebuconazole can inhibit nitrification (Muñoz-Leoz et al., 2011).

Muñoz-Leoz et al. (2011) partially attribute the decreased nitrification to a killing

of fungi, resulting in a large pool of nutrients for ammonifying bacteria combined

with concomitant mineralisation of organic N to NH4+, although ammonia-oxidising

archaea are not mentioned, Muñoz-Leoz also suggest ammonia-oxidising bacteria

are severely inhibited by tebuconazole. Biochar can sorb other pesticides (trifluralin

and atrazine) negating their influence on weeds (Nag et al., 2011); whether sorption

of pesticides to biochar negates their influence on nitrification is unclear.

Biochar addition to agricultural soil may decrease nitrification if it contains VOCs

or polycyclic aromatic hydrocarbons (PAHs) (Clough & Condron, 2010).

Nitrification is sensitive to PAHs and VOCs (Svederup et al., 2002; Uusitalo et al.,

2008), with the degree of sensitivity to PAHs correlating with the lipophilicity

(Svederup et al. 2002). Sverderup et al. (2002) suggest that PAHs exhibit toxicity to

nitrifiers in a non-specfic mode of action through narcoticism, where the function

and fluidity of biological membranes are disturbed through dissolution (partial or

otherwise). The manner in which VOCs such as monoterpenes affect nitrification

has already been discussed. In a study examining the effect of biochar addition to a

soil used for dairy production on N2O emissions, nitrification was not affected by

biochar addition, even though the biochar contained pinene and acetaldehyde

(Clough et al., 2010). Thus, the influence of PAHs and VOCs on nitrification

depends on the chemistry and concentration of the compound(s). The concentration

at which PAHs influence nitrification can range from concentrations greater than 22


32

mg kg-1 (Sverdup et al., 2002). Whilst White (1988) suggest that only a few parts

per million of monoterpenes similar to nitrapyrin may inhibit nitrification. The

VOCs (α-pinene and β-pinene) contained in the Radiata pine biochar of Clough et

al. (2010) were not quantified, so it is unclear at which concentration thresholds

certain VOCs and PAHs can be present in biochar before nitrification is inhibited.

An important phytohormone, ethylene can induce fruit ripening, fine root hair

growth, and other pleiotropic effects (Arbeles et al., 1992) as well as inhibiting

microbial processes such as ammonia oxidation (Porter, 1992). Ethylene can be

produced upon biochar addition to soil. It is unknown whether biochar decreases

ethylene oxidation or increases ethylene production but it has been hypothesised to

contribute to decreases in nitrification and CO2 evolution (Spokas et al., 2010). Its

production upon biochar amendment is highly variable, and could not be found to

correlate with any chemical or physical properties of biochar (Spokas et al., 2010).

Thus, the impact of this mechanism on soil processes requires further research.

1.11 The influence of biochar on plant nitrogen uptake

As an agricultural amendment, the manner in which biochar addition to soil

influences plant N uptake is highly important. Within the Amazonian region,

biochar amendment at 191 t ha-1 (20% w/w) decreased plant N uptake by up to 25%

(Lehmann et al., 2003). Plant N content and uptake decreased by up to 50% and

80%, respectively, with macadamia nut shell biochar additions of 5 and 10% w/w in

a glasshouse study using an Andisol (Haplustand) from the Hawaiian Islands


33

(Deenik et al., 2010). Both studies attributed the decreased plant N uptake to the

biological immobilisation of N (Lehmann et al., 2003; Deenik et al., 2010) but as

stated in Section 1.8 the N content of the microbial biomass was not measured.

Biochar application in north-eastern New South Wales, Australia has increased N

fertiliser use efficiency (Chan et al., 2007; Chan et al., 2008; Van Zwieten et al.,

2010a). In particular, Van Zwieten et al. (2010) claimed that greenwaste biochar

could decrease N fertiliser requirements by up to 90% when applied at 20 and 50 t

ha-1. However the wheat in the experiment was not grown to maturity (Van Zwieten

et al., 2010a) and an explanation for the increased N fertiliser use efficiency was

not provided, but increased nitrification and microbial activity were implicated.

Increased N fertiliser use efficiency with another type of greenwaste biochar

addition occurred at rates of 50 or 100 t ha-1, but not the lowest rate of 10 t ha-1

(Chan et al., 2007). Using a poultry manure biochar application increased N

fertiliser use efficiency when applied at 10 t ha-1 (Chan et al., 2008). Biochar is

relatively deficient in N, compared to potassium (K) and P. Thus, at very high rates

of application (>20 t ha-1), N may become the limiting nutrient, resulting in the

greater efficiency of use (Chan et al., 2007; Chan et al., 2008).

An alternate mechanism by which biochar addition to soil could increase N

fertiliser use efficiency is through the localisation of NO3- in rhizosphere biochar

particles (Prendergast-Miller et al., 2011). Mass flow, caused by root nutrient

uptake, may relocate NO3- in biochar particles within the rhizosphere, potentially


34

allowing greater NO3- uptake (Prendergast-Miller et al., 2011). Although the

retained N is likely to be bio-available (Taghizadeh-Toosi et al., 2012), it is unclear

if this mechanism translates into improved N fertiliser use efficiency because

Prendergast-Miller et al. (2011) did not find any difference in plant N uptake.

Within broadacre agricultural areas of WA practical biochar application rates are

less than 5 t ha-1 (Blackwell et al., 2010). This is due to both the low bulk density of

the biochar and the large areas over which the biochar is to be applied. At

application rates of 5 t ha-1 or less it is unknown whether biochar induced

immobilised N, greater K and P or localisation of NO3- within the rhizosphere will

influence plant N uptake. This could be verified in laboratory and glasshouse

experiments, but field experiments are also required to verify the results where

climatic conditions are not controlled.

1.12 Summary of key research areas highlighted within this review

The addition of biochar to coarse textured soil can alter N cycling processes

because of its porous structure, large surface area and reactive surface. In the

broadacre agricultural area of WA, where more than 60% of topsoil is coarse-

textured (Tennant et al., 1992), the interaction between biochar and the N cycle has

not been investigated. As an agricultural amendment, this thesis focuses on the

interaction between biochar addition to soil and the N cycling processes that

directly influence plant N uptake. To addresses some research deficiencies, this

thesis answers questions on how biochar addition to a coarse textured soil


35

influences N sorption to biochar, N mineralisation, nitrification, N immobilisation

and plant N uptake as summarised in Table 1.4.


36

Table 1.4: A summary of the questions addressed within each thesis chapter.

Chapter & Title Research Questions Addressed in this Thesis

Chapter 2: Decreased

soil microbial biomass

and nitrogen

mineralisation with

Eucalyptus biochar

addition to soil

� When added to a coarse textured soil, does biochar

sorb NH4+, NO3

- or amino acids?

� Does biochar influence net N mineralisation when

added to a coarse textured soil?

� Does biochar addition to a coarse textured soil

induce microbial immobilisation of N?

Chapter 3: Clay and

biochar amendments

decreased inorganic but

not dissolved organic

nitrogen leaching in soil

� Does biochar addition to a coarse textured soil

decrease N leaching?

� Is sorption the main mechanism decreasing

leaching when added to a coarse-textured soil?

Chapter 4: Organic

nitrogen mineralisation

in two contrasting agro-

ecosystems in

unaffected by biochar

addition

� Does organic N mineralisation change when

biochar is added to a coarse textured soil?

� Does organic N mineralisation change over time

after biochar addition to a coarse textured soil?

Chapter 5: Minimal

interaction present

between wheat chaff

biochar and N fertiliser

in a broadacre field

experiment

� When added to a coarse textured soil in a

broadacre Western Australian agro-ecosystem,

does biochar interact with N fertiliser to increase N

fertiliser use efficiency?

� Does biochar influence plant N uptake when added

to a coarse textured soil in a broadacre Western

Australian agro-ecosystem?

37

38

Biochar decreased N mineralisation Chapter 2

39

Chapter 2

Decreased soil microbial biomass and nitrogen

mineralisation with Eucalyptus biochar addition to a coarse

textured soil

2.1 Abstract

Biochar has been shown to aid soil fertility and crop production in some

circumstances. The effects of the addition of Jarrah (Eucalyptus marginata) biochar

to a coarse textured soil on soil carbon and N dynamics were investigated. Wheat was

grown for ten weeks, in soil treated with biochar (0, 5, or 25 t ha-1) in full factorial

combination with N treatments (organic N, inorganic N, or control). Samples were

analysed for plant biomass, soil microbial biomass carbon (MBC) and nitrogen

(MBN), N mineralisation, CO2 evolution, community level physiological profiles

(CLPP) and ammonia oxidising bacterial community structure. MBC significantly

decreased with biochar addition while MBN was unaltered. Net N mineralisation was

highest in control soil and significantly decreased with increasing addition of biochar.

These findings could not be attributed to sorption of inorganic N to biochar. CO2

evolution decreased with 5 t ha-1 biochar but not 25 t ha-1. Biochar addition at 25 t

ha-1 changed the CLPP, while the ammonia oxidising bacterial community structure

changed only when biochar was added with a N source. It was concluded that the

activity of the microbial community decreased in the presence of biochar, through


40

decreased soil organic matter decomposition and N mineralisation which may have

been caused by the decreased MBC.

2.2 Introduction

Application of biochar, a by-product of organic matter pyrolysis and a form of black

carbon (BC), to agricultural soils has received considerable attention (Lehmann &

Joseph, 2009; Clough & Condron, 2010; Lehmann et al., 2011). Biochar can be stable

for thousands of years (Preston & Schmidt, 2006). It has been proposed that biochar

can improve soil fertility and enhance C sequestration (Lehmann et al., 2006; Ogawa

et al., 2006). Such effects may be explained by potential improvements in soil

structure, increases in soil pH, soil moisture and nutrient retention and decreases in

nitrous oxide (N2O) and methane (CH4) emissions (Lehmann et al., 2006; Atkinson et

al., 2010; Sohi et al., 2010). In particular, amendment of soils with biochar (in the

range 0.5 to 135 t ha-1) has been shown to increase plant yield, improve chemical and

physical properties (Glaser et al., 2002), and modify the soil physical habitat for

microbial colonisation, thus altering soil microbial activity and community structure

(Pietikäinen et al., 2000; Steinbeiss et al., 2009). However, as there is a wide range of

biochar types, application rates, soil types and plant responses (Glaser et al., 2002)

the effects of biochar in agro-ecosystems are poorly understood. This is primarily

because there have been few reported studies that aim to elucidate the mechanisms

underpinning these reported effects on plant growth and soil quality.


41

Biochar has a large surface area, especially compared to sands, due to its porous

physical structure and highly aromatic microstructure (Schmidt & Noack, 2000).

Liang et al. (2006) report an enhancement in specific surface area of a sandy soil as a

result of biochar incorporation and that as a consequence these soils had a greater

potential cation exchange capacity and greater retention of dissolved organic matter

than adjacent soil (Liang et al., 2010).This potentially leads to enhanced availability

of C substrates and nutrient source for microorganisms (Zackrisson et al., 1996).

Increases in microbial activity and biomass have been reported with biochar addition

(Lehmann et al., 2011), along with changes in microbial community composition and

abundance (Steinbiess et al., 2009). Some of these increases may be due to biochar

providing a habitat for microbial colonisation (Pietikäinen et al., 2000) and protection

from predation by soil microarthropods (Zackrisson et al., 1996) as bacteria can

potentially inhabit the pores larger than 0.6 μm (Strong et al., 1998). Alternatively, in

the short term biochar may stimulate the microbial community by providing organic

C substrates (Smith et al., 2010). More recently, compounds inhibiting microbial

activity have also been found either on biochar (Deenik et al., 2010) or released after

its introduction to soil (Spokas et al., 2010). Regardless, it is unlikely that these

changes are spread equally across different phylotypes or functional groups and very

little is known about how specific microorganisms are affected (Graber et al., 2010).

Many soil processes may be affected by the incorporation of biochar into soil, in

particular biochar addition has been reported to alter nitrification rates (Berglund et

al., 2004; DeLuca et al., 2006) and potentially adsorb nitrogenous compounds


42

(Lehmann et al., 2003; Hina et al., 2010). In forest soils, biochar has generally been

associated with increased N mineralisation (Berglund et al., 2004; DeLuca et al.,

2006), however, influences within agricultural and grassland soils appear less certain

(DeLuca et al., 2006; Deenik et al., 2010). Biochar application to such soils has

resulted in either no change (DeLuca et al., 2006) or decreases (Deenik et al., 2010)

in N mineralisation, hence lower availability of N for plants (Lehmann et al., 2003).

There is currently limited understanding of the influence of biochar on soil N

processes, and in particular the organisms involved in N transformations. Autotrophic

ammonia oxidising bacteria (AOB) are a microbial functional group that mediate

nitrification, and are influenced by a variety of environmental factors (Gleeson et al.,

2010). The addition of biochar to soil can significantly increase the pH of the bulk

soil (Chan et al., 2008) or alter the local microsite pH, potentially providing a more

favourable habitat for nitrifying organisms, in particular AOB as they are sensitive to

pH (De Boer & Kowalchuk, 2001). There have been few studies examining the effect

of biochar additions on the AOB, with only the study of Ball et al. (2010) showing an

increase in AOB abundance after fire, which they suggest may be related to the

charcoal content and resultant alterations in pH.

The impact of amending a semi-arid agricultural soil with Jarrah (Eucalyptus

marginata Donn ex Sm.) biochar was investigated, with particular focus on C and N

dynamics. A pot experiment was designed in which biochar was added at rates

equivalent to 0, 5, and 25 t ha-1 to a coarse-textured soil that was selected for its low

surface area and lack of microbial refuges, in which wheat was grown. N applied in


43

either an inorganic or organic form was used to alter the C and N availability in the

soil so as to broaden the range of conditions under which changes to the soil

microbial community and soil organic matter decomposition were assessed.Three

hypotheses were investigated: that the addition of biochar to a coarse textured soil

would (i) increase the microbial population size due to increased habitable pore

space; (ii) increase microbial activity (assessed by C and N cycling) due to the

increased microbial biomass and; (iii) cause an increase in microbial community

functional diversity (assessed by CLPP) and AOB diversity (measured by terminal

restriction fragment length polymorphism (T-RFLP) of the amoA gene).

2.3 Materials and methods

2.3.1 Biochar characterisation

Jarrah wood (trunks and large branches; Eucalyptus marginata Donn ex Sm.) was

pyrolysed at 600°C for 24 h in a Lambiotte carbonisation reactor (Simcoa Ltd,

Bunbury, WA). The biochar used in the experiment was the fraction that passed

through a 3 mm seive. Total C and N were determined by combustion analysis (vario

Macro CNS; Elementar, Germany). The ammonium (NH4+) and nitrate (NO3

-)

content were determined by extracting with 0.5 M K2SO4 and analysing the extract

colorimetrically for NH4+

(Krom, 1980; Searle, 1984) and NO3- (Kamphake et al.,

1967; Kempers & Luft, 1988) on an automated flow injection Skalar Auto-analyser

(Skalar San plus).


44

The carbonate equivalence of the biochar was assessed using the method of Rayment

and Lyons (2011). The electrical conductivity (EC) and pH of the biochar were

determined in 1:5 (w/v; g cm-3) soil-to-water or 0.01 M CaCl2 mixtures, respectively.

The gravimetric ash content of the biochar was determined by loss on combustion

(750˚C). The elemental composition of the ash was determined by X-ray fluorescence

(Philips PW1404 XRF). Random powder X-ray diffraction pattern of biochar was

obtained for the range 0-70° 2θ (Philips PW 1830 X-ray diffractometer). Biochar and

soil samples were analysed for pore size distribution and surface area using a surface

area analyser (Micrometrics Gemini 2375 instrument with a Vac Prep 061) (Brunauer

et al., 1938). The biochar was sieved to obtain a particle size distribution. The biochar

contained 27% greater than 1 mm, 23% inbetween 1 mm and 425 μm, 19%

inbetween 425 μm and 180 μm and 31% less than 180 μm. The water holding

capacity of the biochar is 64% w/w and the bulk density is 0.45 g cm-3.

2.3.2 Soil preparation and description

Field dry soil (0-10 cm) was collected from a site 20 km north-west of Moora, WA

(30°32´ S 115°49´ E), sieved (<2 mm) and stored for 4 weeks at 4°C. The soil

contained 7.5 g kg-1 total C, 0.6 g kg-1 total N and 2 mg N kg-1 of NH4+-N and NO3

--

N (Table 2.1). Surface area, pore size distribution, pH and EC of the soil and

soil/biochar mixtures (5 and 25 t ha-1 equivalent) were assessed in the same manner as

for the biochar. The soil used contained 95% sand, 1.5% silt and 3.5% clay, and was

classified as a grey orthic Tenosol (Australian Soil Classification; Isbell 1996).


45

Table 2.1: Basic chemical properties of biochar and soil.

Chemical Properties Biochar Soil

Total N (g kg-1) 3.0 1.0

NH4+ (mg kg-1) < 0.1 2.0

NO3- (mg kg-1) < 0.2 2.0

Total C (g kg-1) 750.0 8.0

pH (CaCl2) 7.4 4.8

EC (μS cm-1) 554.3 44.1

Carbonate (mg g-1) 27.0 -

2.3.3 Pot experiment

A pot experiment was conducted to assess the influence of biochar on the soil

microbial community in the presence of growing plants. The experimental design

consisted of two factors: biochar (3 levels, fixed) and N treatment (3 levels, fixed)

with four replicates in a randomised block. The biochar treatments comprised of

biochar incorporated homogenously into soil at rates equivalent to 0, 5 and 25 t ha-1

(0%, 0.45% and 2.27% w/w respectively). The N treatments were: (1) organic N

applied as 500 kg ha-1 of a wheat straw and pig manure compost (Called ‘Balance’;

Custom Composts, North Dandalup, WA), which contained 15 kg N ha-1; (2)

inorganic N, 100 kg ha-1 of NH4NO3 (containing 35 kg N ha-1) at 3 and 6 weeks; or

(3) control, defined here as those pots receiving only basal N, as described below.

Pots were prepared by adding 2.2 kg of soil or soil/biochar mixture into non-draining

pots with a resulting bulk density of 1.4 g cm-3. This soil naturally compacts to a bulk


46

density between 1.4 and 1.6 g cm3. As such biochar application did not cause any

difference in the final bulk density. Basal fertiliser was added to all pots and then

adjusted to 50% WHC, equivalent to 30% water filled pore space (WFPS). Basal

nutrients were supplied at the following rates (mg kg-1 soil or soil/biochar mixture): N

33.3 (as NH4NO3), P 12.5, K 53.2, S 20.8, Ca 41, Mg 2.4, Mn 3.3, Zn 2.05, Cu 0.5, B

0.1, Co 0.1 and Mo 0.1. Due to the low soil N content (2 mg kg-1 of both NO3- and

NH4+), all pots (i.e. all biochar and N treatments) required an initial basal addition of

33.3 mg kg-1 N (45 kg ha-1 N) as NH4NO3 to ensure adequate plant growth. Three

wheat plants were grown per pot. A near constant soil water content of 50% WHC

was maintained; the WHC was measured and adjusted accordingly for each of the

biochar application rates. Over the length of the experiment the glasshouse average

daily maximum and minimum temperatures were 25 and 13°C respectively. Soil and

plants were collected for analysis 10 weeks after planting. Soil sub-samples (5 g)

were collected for DNA extraction and T-RFLP and stored at -20°C, the remaining

soil was used for soil analyses (MBC, MBN, CLPP, CO2 evolution, N

mineralisation).

2.3.4 Plant analysis

Shoots were harvested by cutting at soil level. Roots were then collected, with soil

removed by washing over a 4 mm sieve. Plant roots and shoots were then dried at

70°C for 72 h and weighed.


47

2.3.5 Microbial biomass

Microbial biomass was determined using the fumigation-extraction method (Vance et

al., 1987). To calculate MBC the resulting flush of oxidisable-C (analysed with a

Shimadzu Model 5050, Japan) was adjusted by a factor of 0.45 (Wu et al., 1990).

MBN was determined using the ninhydrin method (Joergensen & Brookes, 1990) and

adjusted by a factor of 6.47 (Sparling et al., 1993). Sample MBC values were divided

by MBN to obtain C-to-N ratios of the microbial biomass.

To test the extraction efficiency of microbial C in the presence of biochar, 5 g soil (at

50% WHC) was incubated with 500 µL of uniformly 14C-labelled glucose (100 µM;

8.3 kBq mL-1) for 3 h at 20°C in the presence of 1 M NaOH (1 mL) to trap evolved

14CO2. After incorporation of the 14C-label into the soil microbial biomass, replicate

samples of 14C-labelled soil were extracted with 0.5 M K2SO4 for 15 min (200 rev

min-1), centrifuged (18,000 g) and the 14C content of the supernatant determined by

liquid scintillation counting (unfumigated controls). The remaining soil samples were

CHCl3-fumigated (24 h) and then extracted with 0.5 M K2SO4 as described above,

except that biochar (0, 0.45 or 2.27% w/w) was added to the soil immediately before

extraction. The amount of 14C immobilised in the microbial biomass was assumed to

equal the total amount initially added minus that recovered as 14CO2 and as 14C in the

unfumigated K2SO4 extracts. The effect of biochar and fumigation on the extraction

of amino acids was also assessed.


48

2.3.6 Soil carbon dioxide evolution

At harvest, soil samples from the pot experiment were incubated in glass jars

containing a gas septum, adjusted to 40% WHC, incubated at 15ºC, and sealed to trap

respired CO2 (Anderson, 1982). Headspace gas was analysed for CO2 with an infra-

red gas analyser (IRGA; The Analytical Development Co., Series 225). Gas samples

were collected over a period of 65 d.

2.3.7 Community level physiological profiles (CLPP)

Catabolic response profiles were measured using the method of Degens & Harris

(1997). Substrates used were those specified by Stevenson et al. (2004) at the

concentrations used by Lalor et al. (2007). Headspace gas (1 mL) was analysed for

CO2 in an IRGA. The richness, evenness and diversity of CLPP profiles were

determined using indices described by Blackwood et al. (2007): richness (S),

Shannon diversity index (H’), Simpson index (D) and Shannon evenness (J’).

2.3.8 DNA extraction and terminal restriction fragment length polymorphism (T-

RFLP) profiles

Total nucleic acids were extracted from 0.5 g soil or soil/biochar mixes using

previously described methods (Griffiths et al., 2000) with a minor modification

(samples were homogenised using a Mini-BeadBeater-8 (Biospec Products Inc.,

Bartlesville, OK, USA) at 3200 rev min-1, 2 min).


49

Polymerase chain reaction (PCR) amplification of the amoA gene was based on the

method of Horz et al. (2004), using the primer set amo-1F (5’-

GGGGTTTCTACTGGTGGT-3’) and amo-2R (5’-CCCCTCKGS

AAAGCCTTCTTC-3’) (Rotthauwe et al., 1997) as previously described (Gleeson et

al., 2010). All PCRs were performed in duplicate and pooled for subsequent

purification, restriction and fragment analysis. Purified PCR products were quantified

and approximately 100 ng of PCR product was used in a restriction digest with the

restriction endonuclease enzyme HaeIII (New England Biolabs Inc.) as per the

manufacturer’s instructions. Terminal restriction fragment lengths (T-RFs) were

determined by electrophoresis using a capillary electrophoresis system (Applied

Biosystems) and analysis of fragment profiles was carried out using Genemapper®.

Profiles were generated for each sample based on relative height of peaks indicative

of relative abundance within the sample. T-RFs were assigned using the program

RiboSort (Scallan et al., 2008) and the statistical software R (R Core Development

Team, 2007). T-RFs in different profiles that differed by less that 0.5 bp were

considered identical (Dunbar et al., 2001). For each sample, bacterial T-RFLP

profiles were standardised by total fluorescence and fragments contributing <0.1% to

the total fluorescence of that sample were removed. The richness, evenness and

diversity of amoA T-RFLP profiles were determined using indices (S, H’, D and J’)

described by Blackwood et al. (2007).


50

2.3.9 Net nitrogen mineralisation

At harvest, soil and soil/biochar mixtures from the pot experiment were adjusted to

40% WHC and incubated at 15ºC in replicate containers that allowed free air

exchange. From this incubation, samples were collected over 70 days. Soil NH4+ and

NO3- was analysed as previously mentioned. Net N mineralisation and nitrification

rates were determined as the changes in the sizes of the inorganic N (NH4+ plus NO3

-)

and NO3- pools, respectively, over time.

2.3.10 Organic carbon sorption experiment

A mixture of 16 individual uniformly 14C-labelled amino acids was added to 0.5 M

K2SO4 to give a total concentration of 40 mg C l-1 (ca. 0.4 mM; 0.6 kBq ml-1). This

solution was then mixed with either; soil; soil/biochar mixture (25 t ha-1 or 2.27%

w/w) or; biochar alone at a soil-to-solution ratio of 1:4 (w/v). The soil suspension was

then shaken for 15 min at 200 rev min-1. After shaking, the suspensions were

centrifuged (16,000 g, 5 min) and the supernatant recovered for 14C determination

using a 1404 liquid scintillation counter (Wallace EG&G Ltd., Milton Keynes, UK).

These tests were both conducted using fresh soil and fresh biochar.

2.3.11 Nitrogen sorption experiment

Sub-samples (10 g) of fresh biochar, soil, or soil/biochar mixtures (5 or 25 t ha-1) and

100 mL of organic or inorganic N (leucine-N, (NH4)2SO4, KNO3 or NH4NO3)

solution were added to 120 mL vials. Vials were shaken for 24 h after which solution

N concentrations were measured for organic N, NO3- or NH4

+ as mentioned


51

previously. To replicate the extraction methodology, this procedure was repeated,

except after shaking for 24 h, K2SO4 salt was added to a concentration of 0.5 M and

shaken for a further 1 h. The N concentrations used were 0, 0.5, 1, 1.67, 2.5, 5, 10,

25, 50 and 100 mg N L-1 for leucine, (NH4)2SO4 and KNO3. The N concentrations for

NH4NO3 were doubled to ensure the same concentrations of NH4+ and NO3

- as the

previous incubations. The dilute concentrations (0, 0.5, 1 and 1.67 mg N L-1) were

equivalent to those in the net N mineralisation incubation (0, 5, 10 and 16.7 mg N

kg-1 soil).

2.3.12 Statistical analysis

A series of general analysis of variance (ANOVA) using GENSTAT 10th Edition

(Lawes Agricultural Trust, 2007) were completed. Multiple comparisons were done

using the Duncan’s Multiple Range Test (DMRT) and significance considered at the

P ≤ 0.05 significance level.

Multivariate statistical analyses were performed on standardised, (Log (X+1))

transformed CLPP data and standardised, untransformed T-RFLP profiles using

Primer 6 (Plymouth Marine Laboratory, 2007) and Bray-Curtis similarity matrices.

Tests of the null hypothesis among a priori defined groups were examined using

permutational multivariate ANOVA (PERMANOVA) for both data sets. All

PERMANOVA tests used 9999 permutations of raw data from residuals under a

reduced model (Anderson, 2001). Pairwise comparisons were performed to determine

whether amoA communities differed significantly (P<0.05) between biochar and N-


52

treatments. Analyses of differences in richness, evenness and diversity measures

between treatments were performed using ANOVA.

Data of solution concentrations were analysed to give 95% confidence intervals for

the means; these were used to judge significance.

2.4 Results

2.4.1 Biochar characterisation

Biochar comprised 75% C and 0.3% N with a C:N ratio of 275:1 (Table 2.1); this

equated to an addition of 75 and 15 kg N ha-1 in the 25 and 5 t ha-1 biochar

treatments, respectively. However, The NH4+ and NO3

- content of the extractions

were below the detection limit (<0.2 mg L-1 NO3- and <0.1 mg L-1 NH4

+; Table 2.1).

The biochar had a pH of 7.4 and an EC of 554.3 μS cm-1 (Table 2.1). Biochar

application increased the soil pH from 4.77 to 4.88 and 5.06 with the addition of 5

and 25 t ha-1 of biochar, respectively. The carbonate content of the biochar was 27 mg

g-1 (Table 2.1). The ash content was 60 mg g-1. The major elements in the ash,

determined by X-ray fluorescence analysis, were Si (206 mg g-1), Al (127 mg g-1), Fe

(91 mg g-1), Ca (54 mg g-1) and Mg (11 mg g-1). X-ray diffraction showed that the

biochar contained amorphous C, quartz and calcite (data not shown).

The amount of pore space surface area potentially habitable by microorganisms,

determined on the proportion of pores >0.60 μm, in the biochar alone was 4.42 m2 g-1,


53

which was six times greater than the soil (0.74 m2 g-1) pore area (Table 2.2).

However, at biochar application rates of 5 and 25 t ha-1 to soil, calculations

demonstrate this equates to an increase of 0.02 and 0.08 m2 g-1 of habitable pore space

(i.e. net increase of 2.7 and 10.8%).

Table 2.2: Incremental and total pore area (m2 g-1) for soil and biochar, along with

calculated estimates for application rates of 5 and 25 t ha-1 of applied biochar, for

the range of pore diameters deemed habitable by bacteria.

Pore diameter (μm) Quantity of biochar added (5 and 25 t ha-1 values calculated)

Soil 5 t ha-1 25 t ha-1 Biochar

4.50-1.90 0.28 0.28 0.28 0.49

1.90-1.20 0.19 0.19 0.20 0.67

1.20-0.90 0.13 0.13 0.15 0.96

0.90-0.70 0.09 0.09 0.11 1.05

0.70-0.60 0.05 0.06 0.08 1.25

Total 0.74 0.76 0.82 4.42

2.4.2 Plant growth analysis

ANOVA showed significant differences due to both biochar (P<0.001) and N

(P<0.001) treatments with no interaction. Post hoc comparisons using DMRT showed

biochar addition at 25 t ha-1 was associated with a significantly lower shoot dry

weight (6.1 g) compared to the 0 and 5 t ha-1 application rates (6.9 and 6.3 g,

respectively; Fig. 2.1a). Inorganic N addition resulted in significantly greater shoot


54

dry weight (8.3 g) compared to the organic N (5.8 g) and control treatments (5.7 g;

Fig. 2.1a). No differences between root dry weights were evident (data not shown);

however the root to shoot ratio was significantly lower with inorganic N application

(P<0.001; Fig. 2.1b).

Sh

oo

t Dry

Wei

gh

t (g

)

Ro

ot:S

ho

ot R

atio

0

1

2

3

4

0 5 25 0 5 25 0 5 25

Control Compost Inorganic N

(b)

0

2

4

6

8

10 (a)

Fig. 2.1: Shoot dry weight (a) and root: shoot ratio (b) results for the control,

organic N (compost) and inorganic N treatments at three added rates of biochar (0, 5

and 25 t ha-1). Error bars represent standard errors (n=4).


55


Overall, 97.4 ± 0.2% of the added 14C glucose-C was taken up by the microbial

biomass after 3 h, of which 81 ± 2% was immobilised and 19 ± 2% was respired as

14CO2 (data not shown). Chloroform fumigation-extraction revealed that 21.0 ± 0.1%

of the added 14C was present in the microbial pool whilst 79.0 ± 0.1% remained non-

extractable (data not shown). Overall, the presence of biochar at both addition rates

had no significant effect on the recovery of 14C from the soil microbial biomass. In

addition, biochar did not affect the recovery of amino acids either before or after

fumigation (Table 2.3). No significant difference in the recovery of leucine-N

standards over the range 0 to 50 mg amino-N kg soil-1 in the presence of biochar

when extracted in 0.5 M K2SO4 was found (data not shown). As such the fumigation-

extraction method was not affected by the presence of the biochar.

Table 2.3: Recovery of 14C-labelled amino acids added to soil in the presence and

absence of biochar either before or after CHCl3 fumigation. Values represent means

± standard errors (n=4). There were no significant differences between any of the

treatments (P > 0.05).

Non-fumigated Fumigated

(% recovery) (% recovery)

Soil 93.5 ±2.6 95.9 ±2.4

Soil + biochar (25 t ha-1) 99.2 ±5.9 103.6 ±4.1

Biochar 102.8 ±4.7 104.6 ±4.8


56

There was a significant effect of biochar rate on MBC (P=0.05), and of N treatment

(P<0.001), but no interaction between the treatments. MBC was greatest without

biochar addition (145 mg C kg-1) but decreased significantly with 25 t ha-1 biochar

addition (116 mg C kg-1; Fig. 2.2a). MBC was also significantly lower in the organic

N (135 mg C kg-1) and inorganic N (93 mg C kg-1) treatments compared to the control

(159 mg C kg-1) (Fig. 2.2a). MBN did not change with either biochar or N addition

and there was no interaction between the variables.

The initial microbial biomass C:N ratio was 8:1. This was changed by the addition of

N (P=0.005), but not biochar addition and there was no interaction. The microbial

C:N ratio was significantly decreased by the inorganic N treatment to 4.5:1 (Fig.

2.2c).


57

Mic

rob

ial B

iom

ass

C

(mg

kg-1

)

Mic

rob

ial b

iom

ass

N

(mg

kg-1

)

M

icro

bia

l Bio

mas

s 44

C:N

rat

io0

50

100

150

200

250 (a)

0

10

20

30 (b)

0

2

4

6

8

10

0 5 25 0 5 25 0 5 25

Control Compost Inorganic N

(c)

Fig. 2.2: Microbial biomass carbon (a), microbial biomass nitrogen (b) and

microbial biomass C:N ratio (c) results for the control, organic N (compost) and

inorganic N treatments at three added rates of biochar (0, 5 and 25 t ha-1). Error

bars represent standard errors (n=4).


58

2.4.4 Microbial community function (CLPP) and structural diversity (amoA T-

RFLP)

Multivariate analysis by PERMANOVA showed that addition of 25 t ha-1 biochar

significantly changed the CLPP (P<0.01), but addition of 5 t ha-1 biochar did not. The

N treatments had no effect on CLPP and there was no interaction between the two

factors (Table 2.4). Analysis of T-RFLP data revealed a significant effect of biochar

(P<0.001) but no significant effect of N (P<0.167) on ammonia oxidiser community

structure. There was a significant interaction (P<0.001) between biochar and N

treatments: without N addition (either compost or inorganic N), there was no effect of

biochar on the ammonia oxidiser community structure. However, in the presence of

compost and inorganic N there was a significant effect of biochar addition (Table

2.4).

The effect of biochar addition on CLPP diversity was only observed in the compost

or inorganic N treatments (P=0.002) where both 1/D and H’ significantly increased

with biochar addition (data not shown).

The amoA T-RFLP richness (S) averaged 64 and evenness, J’, averaged 0.67 across

all treatments; neither were significantly affected by biochar rate or N treatment.

However, in terms of evenness, there was a significant interaction between treatments

(P=0.03); when organic N (compost) was added there was a significant effect on

evenness; this was not the case with inorganic N addition or in the control treatment.


59

Diversity averaged 2.78 (H’) and 9.99 (1/D) with no significant effect of biochar rate

or N-treatment (data not shown).

Table 2.4: Significance of biochar and nitrogen treatments on terminal restriction

fragment length polymorphism (T-RFLP) and community level physiological profiles

(CLPP). NS, *, **, and *** denote not significant (P > 0.10), P < 0.10, P < 0.05,

and P < 0.005 respectively.

T-RFLP CLPP

Overall effects

Biochar *** ***

Nitrogen treatment NS NS

Biochar x Nitrogen treatment *** NS

Multiple comparisons within nitrogen treatment for TRFLP

Biochar rate (t ha-1) 0 5

Compost 5 **

25 * *

Nitrogen 5 *

25 * *

No treatment 5 NS

25 NS NS


60

2.4.5 Carbon dioxide evolution and nitrogen mineralisation

There was no interaction between biochar and N treatments but both biochar and N

had significant effects on CO2 evolution (P=0.003 and 0.001, respectively).

Cumulative CO2 evolution from soil was significantly lower where biochar was

added at 5 t ha-1 (101 mg CO2-C kg-1); with no difference between the 0 and 25 t ha-1

of biochar addition (Fig. 2.3). The addition of compost did not change soil CO2

evolution while the addition of inorganic N caused a significant decrease (data not

shown).

CO

2 ev

olu

tion

(mg

CO 2

-C k

g -1 )

Incubation period (weeks)

0

50

100

150

0 2 4 6 8 10

0 t ha5 t ha25 t ha

-1

-1

-1

Fig. 2.3: Soil cumulative CO2 evolution in the control treatment for biochar amended

at 0 (crosses), 5 t ha-1 (open triangles) or 25 t ha-1 (solid circles). Error bars

represent standard error (n=4).


61

Inorganic N in the incubated soil declined significantly with increasing biochar rate in

all three N treatments (P<0.001; Fig. 2.4). On average across N treatments the

inorganic N concentration was 11 mg kg-1, 7 mg kg-1 and 1 mg kg-1 where biochar

was added at 0, 5, and 25 t ha-1, respectively, after 10 weeks of incubation (Fig. 2.4).

This equated to significantly lower average net N mineralisation rates of 176, 102 and

15 μg N kg-1 day-1 with biochar addition at 0, 5 and 25 t ha-1, respectively (Table 2.5;

P<0.001). Net nitrification rates also decreased significantly with increasing biochar

addition (P<0.001; Table 2.5). Average net nitrification rates decreased from 178 μg

N kg-1 day-1 without biochar addition, to 103 and 12 μg N kg-1 day-1 with biochar

added at 5 and 25 t ha-1, respectively. At week 10, the proportion of N present as

NH4+ was significantly higher with biochar addition at 25 t ha-1 (average of 56%

across N treatments) than at 5 t ha-1 and the control (an average of 3.2% and 2.1%

across N treatments, respectively; P<0.001; Table 2.5).


62

Table 2.5: Final rates of net N mineralisation, net nitrification and the proportion of

NH4+-N at week 10. Values represent means ± standard errors (n=4), letters

represent significant differences (P < 0.05).

Nitrogen

Treatment

Biochar rate

(t ha-1)

Net N mineralisation

rate (μg kg-1 day-1)

Net nitrification rate

(μg kg-1 day-1)

NH4+-N proportion of

net N (%)

Control 0 145.8 ±10.8 cd 144.3 ±9.6 c 2.3 ±0.6

5 78.0 ±12.3 b 76.0 ±13.2 b 3.9 ±1.2

25 29.6 ±12.1 a 25.6 ±11.6 a 33.0 ±16.8

Compost 0 195.6 ±20.4 d 191.2 ±20.8 d 2.9 ±0.5

5 138.0 ±37.9 c 136.2 ±35.0 c 2.1 ±1.2

25 5.3 ±5.1 a 1.8 ±4.6 a 79.6 ±16.7

Inorganic N 0 186.3 ±7.7 cd 198.9 ±7.5 d 1.1 ±0.2

5 90.1 ±7.7 b 98.4 ±8.6 bc 3.5 ±1.2

25 8.7 ±5.3 a 8.6 ±5.6 a 57.2 ±24.3


63

Ino

rgan

ic N

1

(

mg

N k

g-1)

Ino

rgan

ic N

1

(

mg

N k

g-1)

In

org

anic

N

1

(m

g N

kg-1)

Incubation period (Weeks)

0

5

10

15

0 t ha5 t ha25 t ha

(A)-1

-1

-1

0

5

10

15 (B)

0

5

10

15

0 2 4 6 8 10

(C)

Fig. 2.4: Total inorganic N extracted from the control (a), organic N (compost) (b),

and inorganic N (c) treatments for biochar added at 0 (crosses), 5 t ha-1 (triangles)

and 25 t ha-1 (closed circles). Error bars represent standard errors (n=4).


64

2.4.6 Sorption isotherms

Both NH4+ and NO3

- sorbed to the biochar when mixed in water for 24 h (Fig. 2.5b).

In water, the amount of NH4+ sorbed to biochar ranged from 88% at the lowest initial

NH4+ concentration (0.5 mg N L-1) to 50% at higher concentrations (>25 mg N L-1).

Recovery of NH4+ increased upon extraction with 0.5 M K2SO4, where around 35%

of the added NH4+ was sorbed up to a concentration of 5 mg N L-1 (Fig. 2.5a). At

concentrations of 5 mg N L-1 to 25 mg N L-1, 20% of added N was sorbed, with no

significant sorption at greater concentrations (data not shown). In comparison, NO3-

sorption to biochar was much greater ranging from around 95% at concentrations up

to 2 mg N L-1 to 75% at concentrations greater than 5 mg L-1. With 0.5 M K2SO4 it

was possible to recover only 5% of the applied NO3- (i.e. 95% sorption) up to initial

concentrations of 5 mg N L-1 (Fig 2.5a). Above this, NO3- recovery increased

gradually until 12 mg N L-1 after which approximately 50% of the applied NO3- could

be recovered (data not shown). However, at the low rates of biochar addition to soil

(5 and 25 t ha-1; or 0.45 and 2.27% w/w) there was not statistically significant

sorption of either form of inorganic N (Fig. 2.6).


65

N c

on

cen

trat

ion

(m

g k

g-1)

P

ost

incu

bat

ion

so

lutio

n

Initial Solution N concentration (mg kg-1)

0

10

20

30

40

50(a)

0

10

20

30

40

50

0 10 20 30 40 50

(b)

Ammonium

Nitrate

Fig. 2.5: Sorption data for ammonium (diamonds) and nitrate (squares) for biochar

extracted with (a) 0.5 M K2SO4 and (b) water. Error bars represent standard errors

(n=3) and the 45° line represents an unchanged solution concentration.


66

N C

on

c. (

mg

kg-1)

P

ost

incu

bat

ion

so

lutio

n

Initial Solution N Conc. (mg kg-1)

0

10

20

30

40

50(a) (b)

0

10

20

30

40

50

0 10 20 30 40 50

(c)

0 10 20 30 40 50

(d)

0 t ha

5 t ha

25 t ha

(A)-1

-1

-1

Fig. 2.6: The solution concentrations of 0.5M K2SO4 extracted (a) NH4+ and (b) NO3

-

or water extracted (c) NH4+ and (d) NO3

- for soil (crosses), biochar added to soil at

either 5 t ha-1 equivalent (0.45% w/w; triangles), or 25 t ha-1 equivalent (2.27% w/w;

circles) after shaking for 24 h. Error bars represent standard errors (n=3) and the

45° line represents an unchanged solution concentration.


67

2.5 Discussion


Bacteria typically inhabit pores greater than 0.6 μm (Strong et al., 1998) and the

biochar used here had a habitable pore area over 6 times that of the soil; assuming all

pores on the biochar were favourable for microbial habitation (i.e. not hydrophobic).

However, when biochar was mixed with soil the calculated slight increase in

habitable pore space (Table 2.2) did not translate into increased microbial biomass

(Fig. 2.2).

Elevation of soil pH in acidic soils can increase microbial biomass and activity

(Aciego Pietri & Brookes, 2008). The addition of biochar (CaCl2 pH; 7.43) would

have introduced alkaline micro-habitats potentially providing more favourable niche

environments for microbial populations (Lehmann et al., 2011). However, MBC, N

mineralisation and net nitrification decreased in the presence of biochar; thus

indicating that any benefit from the ‘liming effect’ of the biochar was outweighed by

a decrease in the mass and activity of the microbial population. An increase in

alkaline microsites may also change ammonia oxidiser community structure as rates

of nitrification, and in particular ammonia oxidation, are significantly reduced in acid

soils (De Boer & Kowalchuk, 2001). However, T-RFLP of the amoA gene

demonstrated that the ammonia oxidiser community did not change with the addition

of biochar alone to soil. Differences were only observed when biochar was added in

conjunction with inorganic or organic N.


68

The decrease in both MBC and shoot biomass with increasing biochar application

rate contrasts with other studies (Chan et al., 2008; Steiner et al., 2008; Kolb et al.,

2009; Lehmann et al., 2011). This may be explained in part by differences in biochar

application rate and type (e.g. biochar feedstock, pyrolysis temperature, etc.) along

with soil type. Methodological differences may also explain microbial biomass

differences between studies. Previous studies have used the SIR method (Anderson &

Domsch, 1978) to measure microbial biomass (Steiner et al., 2008; Kolb et al., 2009)

where CO2 evolution measurements may be confounded by abiotic release (Jones et

al., 2011; Zimmerman, 2010). By contrast, this study used the fumigation-extraction

method (Vance et al., 1987). Previous studies may not have used the fumigation-

extraction method due to the potential for sorption of cell contents to biochar after

chloroform induced cell lysis. It could be expected that this may confound results and

skew MBC values. However, results in Table 2.3 show there was no effect of biochar

on the recovery of 14C labelled microbial biomass or sorption of amino acids during

the fumigation-extraction process. This is in agreement with the findings of

Durenkamp et al. (2010) who also found that biochar did not change the 0.5 M

K2SO4 extraction efficiency of C (to determine MBC) or ninhydrin-N (to determine

MBN) although activated charcoal did.

Volatile compounds present on biochar have the potential to decrease microbial

biomass (Deenik et al., 2010). Girvan et al. (2005) showed that benzene can decrease

the microbial biomass at concentrations of 40 mg kg-1 soil. In this study, all tested

volatile compounds of the biochar were below the detection limits apart from benzene


69

(0.5 mg kg-1; data not shown). But at such low concentrations the effect is unlikely to

solely be attributed to this.

2.5.2 Carbon and nitrogen mineralisation

Interestingly a decline in soil CO2 evolution occurred when 5 t ha-1 biochar was added

but there was no difference in CO2 evolution between 25 t ha-1 and the control soil.

At the levels of biochar (5 and 25 t ha-1) applied to soil it was assumed that both

biotic and abiotic processes were rate dependent meaning that a larger response

should be observed at the higher rate of addition. Given that the biochar contained

carbonate (2.7%) and the soil was acidic (pH 4.76) some dissolution was likely to

have occurred resulting in the loss of HCO3- as CO2. The rate of dissolution was not

determined but was likely to depend on soil water content, the physical location of

the carbonate within the structure of the biochar (i.e. diffusion limitations) and

possibly localised pH buffering (liming effect) of the biochar (pH 7.43). This was

likely to be significant at micro-sites given the low buffering capacity of this

predominately quartz (95% sand) and kaolinitic clay (1:1 lattice structure) dominated

soil. However, assuming that all the carbonate-C was converted to CO2 during the

incubation then this would equate to 24.3 and 122.7 mg CO2-C kg-1 soil in the 5 and

25 t ha-1 biochar applications respectively; this is of the same order of magnitude as

the CO2 evolved from the control soil. Results indicate there was no difference

between the CO2 evolved from the 25 t ha-1 biochar treatment and the control soil;

this suggests that any CO2 derived from carbonate had to be offset by a negative

priming effect on soil organic matter respiration. Results for the 5 t ha-1 biochar


70

treatment supports the occurrence of a negative priming effect as here addition of

biochar did cause a significant decrease in CO2 evolution compared to the control

soil. This is consistent with previous research showing suppression of biotic CO2

evolution after biochar addition to soil (Jones et al., 2011; Keith et al., 2011).

As C and N mineralisation are closely linked a negative priming effect on the soil

organic matter should also have been expressed in the N mineralisation results.

Indeed, the size of the inorganic N pool was significantly lower in the presence of

biochar (P<0.001). This was associated with significantly lower net N mineralisation

rates as biochar application rate increased (P<0.001). Also the rate of nitrification

significantly decreased with increasing rates of biochar addition in the control and in

the presence of both N (inorganic N treatment) and C and N (compost treatment)

(P<0.001; Table 2.4). However, the ammonia oxidiser community structure remained

unaffected by biochar addition, unless in the presence of an added N source and there

was no change in diversity. Here it is likely that the lower NH4+ level in the presence

of biochar has constrained nitrification through substrate limitation due to the

negative priming effect on the soil organic matter decomposition. As such the

negative priming effect of biochar addition on soil organic matter decomposition was

illustrated when assessed by determination of N mineralisation and partially masked

due to the likely abiotic release of CO2 when assessed by CO2 evolution.

A decline in net N mineralisation could also have been caused by an increase in the

rate of microbial N immobilisation compared to the gross N mineralisation rate.


71

Gundale & DeLuca (2007) suggested that the relative dominance of mineralisation

versus immobilisation in the presence of charcoal was determined by the level of

bioavailable C in their incubation (i.e. glycine). These results do not indicate an

increase in the size of the MBC where compost was applied and in addition MBN

remained constant in all treatments, suggesting that increased microbial

immobilisation of N did not occur.

Sorption of nitrogenous compounds onto biochar is another mechanism that could

explain the lower inorganic N traces in the presence of biochar. Microbial activity

oxidises biochar through time, the sorption and desorption characteristics of biochar

are likely to change. Over time surface positive charge is likely to decrease, whilst

negative charge increases (Cheng et al., 2008). A weakness of this sorption study is

that measurements have only occurred at one time point, rather than at the start and

end of the incubation. However, the extent of oxidation is highly correlated with

temperature (Cheng et al., 2006; Cheng et al., 2008) and the study by Cheng et al.

(2006) did not measure any net change in total acidic functional groups after a four

month incubation at 30°C. Given the temperature range of the glasshouse (mean

maximum of 25°C and mean minimum of 13°C) and that the subsequent soil

incubations were conducted at 15°C the biochar was unlikely to have oxidised

sufficiently over the course of the experiment to warrant a time course study of

sorption. These results did indicate NO3- and to a lesser extent NH4

+ sorption to

biochar (Fig. 2.5). However when biochar was mixed with soil, the very minimal

extent of sorption that occurred could not explain the significantly decreased N


72

mineralisation (Fig. 2.6). This is attributed to the small proportion of biochar within

the soil matrix (2.27 and 0.45 % w/w in the 25 and 5 t ha-1 treatments respectively)

and the slow rate of diffusion of NH4+ in solution.

The decrease in mineralisation may have been caused by the presence of potentially

toxic volatile organic compounds present on the biochar during addition (Deenik et

al., 2010), or the evolution of ethylene (Spokas et al., 2010). The Jarrah biochar was

assessed for volatile organic compounds, but all assessed compounds were below the

detection limit, aside from benzene (0.5 mg kg-1 of biochar; data not shown). At such

low levels, the decreases in mineralisation cannot solely be attributed to the presence

of benzene on the biochar used in this study.

This study has provided evidence that biochar addition to soil decreased MBC but not

MBN. Associated with this was a coincident decrease in soil organic matter

decomposition as indicated by the decline in CO2 evolution and net N mineralisation

rates. The microbial community functional diversity (as measured by CLPP) changed

with biochar addition; however the diversity of the ammonium oxidiser community

only changed when biochar was added in conjunction with an N source (organic or

inorganic). Thus findings did not support the hypotheses that biochar addition to soil

would increase the mass, activity and structural diversity of the soil microbial

community. This highlights the need to further determine the conditions (i.e. biochar

feed stock, production conditions, soil types) under which biochar additions to soil


73

will result in positive or negative effects on the soil microbial community and their

associated functions.

74

Clay and biochar decrease inorganic N leaching Chapter 3

75

Chapter 3

Clay and biochar amendments decreased inorganic nitrogen

leaching but not dissolved organic nitrogen leaching in soil

3.1 Abstract

N leaching from coarse textured soils frequently leads to productivity losses and

negative environmental consequences. Historically, clay amendment has been used

on coarse textured soils to decrease water repellence and nutrient leaching. More

recently biochar has been proposed as an alternative soil amendment to decrease N

leaching while simultaneously storing carbon. As biochar has a greater nutrient

retention capacity, it was hypothesised that Eucalyptus marginata derived biochar

would be a more effective amendment than clay at minimising N leaching. Lysimeter

pots were filled with a coarse-textured agricultural soil and amended with the

following treatments: (1) biochar incorporated homogenously into the 0-10 cm soil

layer, (2) clay incorporated similarly, (3) biochar added as a layer at 10 cm depth, (4)

clay added similarly, or (5) a control. Amendments were added at 25 t/ha and watered

periodically over 21 days and watered with the equivalent to 30 mm. Clay and

biochar amendments significantly decreased cumulative ammonium (NH4+) leaching

by approximately 19% and 14% respectively and NO3- leaching by 16% and 28%

respectively. Biochar decreased nitrate (NO3-) leaching significantly more than clay,

possibly due to decreased nitrification. Dissolved organic N leaching was not

influenced by any treatment. N leaching was unaffected by amendment application


76

method, therefore to decrease N leaching, land managers should apply the most

readily available of the amendments in the most economic and convenient manner.

3.2 Introduction

Improving soil fertility and productivity, while simultaneously decreasing negative

environmental externalities, is one of the greatest challenges in the management of

agricultural and horticultural production systems. This is typically very difficult to

achieve on coarse-textured soils with poor intrinsic soil structure, high water

repellence, low water holding capacity and low nutrient retention capacity (Harper et

al., 2000; Moore, 2004). These soils dominate the agricultural region within the

Mediterranean-type climate zone of Australia (Rovira, 1992). Typical fertiliser

regimes on such soil types range from 400 kg N ha-1 crop-1 in high value horticultural

systems, where 3 crops may be grown per year (Flavel & Murphy, 2006) to typically

40-80 kg N ha-1 for an expected 2 t ha-1 wheat grain yield (Hoyle et al., 2011).

Nutrient leaching from agricultural and horticultural production systems represents

not only an operational inefficiency, but can also cause environmental problems, such

as generating excess soil acidity (Dolling & Porter, 1994), eutrophication (e.g.

Smolders et al., 2010) and pollution of ground water (Power et al., 2001). The risk of

eutrophication in these ecosystems is particularly problematic as fresh and marine

water ecosystems have adapted to low nutrient environments (ANZECC &

ARMCANZ, 2000). One potential intervention strategy to protect these environments

is to minimise the use of soluble fertilisers (Gentile et al., 2009), however, this may


77

decrease land productivity (Quiroga-Garza et al., 2001). The risk of leaching is

greatest early in the growing season when N availability is greater than demand,

hence leaching moves N below the rooting zone (Hoyle & Murphy, 2011). Soil

amendments such as biochar and clay may retain N and decrease N leaching, thus

improving N use efficiency while also minimising eutrophication risks.

Biochar is the solid remains of thermally decomposed organic matter, from plant or

animal sources, under limited oxygen supply at temperatures less than 700 °C

(Lehmann & Joseph, 2009). It has the potential to sequester carbon (Lehmann et al.,

2006; Ogawa et al., 2006) and improve soil fertility (Glaser et al., 2002). One of the

potential mechanisms for improved soil fertility is through enhanced nutrient

retention, resulting in decreased leaching losses (Lehmann et al., 2003). However,

significant uncertainty still surrounds its impact on soil N dynamics. While biochar

application may decrease ammonium (NH4+) leaching (Lehmann et al., 2003; Ding et

al., 2010) it may stimulate nitrate (NO3-) leaching (Laird et al., 2010b; Lehmann et

al., 2003).

Duplex or texture contrast soils, where underlying the coarse-textured topsoil is a

subsoil of much greater clay content, comprise 56% or 14.2 m ha of Western

Australian agricultural land (Tennant et al., 1992). On some farms, these coarse-

textured topsoils are being ameliorated with subsoil clay, termed ‘clay’ in this paper.

Field application rates are in excess of 100 t ha-1 (Harper & Gilkes, 2004) and as

such, clay amendments can ameliorate water repellence (McKissock et al., 2000;


78

McKissock et al., 2002). Furthermore, by increasing the ion retention capacity clay

amendment can increase nutrient holding capacity, hence increase yields (Hall et al.,

2010) and decrease leaching. In a similar manner, biochar can also increase yields

and decrease nutrient leaching (Lehmann et al., 2003). However, the ability of each

amendment to ameliorate these properties will vary between clay and biochar types

and their intrisic properties. For example, published cation exchange capacities for

clay amendments range from 6-10 cmolc kg-1 (Hall et al., 2010) and from 6-25 cmolc

kg-1 for biochar (Lehmann, 2007).

The ability of clay and biochar amendments to ameliorate leaching of inorganic and

DON were tested and compared in lysimeter pots. It was hypothesised that both

amendments would decrease N leaching, due to expected greater ion retention

capacity. Whether the placement of these amendments within the profile, i.e. a

homogenous incorporation versus a horizontal layer, influences amendment

performance was also tested. The hypothesis was that the layer would decrease

leaching to a greater extent due to improved interception of leached ions.

3.3 Materials and Methods

3.3.1 Soil, clay and biochar

Soil was collected from a White Subnatric Sodosol (Isbell, 1996) agricultural soil

under a historical crop (generally wheat (Triticum aestivum))/pasture (cape weed

(Arctotheca calendula), annual ryegrass (Lolium rigidum) and clover (Trifolium


79

subterraneum) dominant) rotation located 15 km north of Meckering, WA, Australia

(31° 30′ S, 116° 59′ E) in 0-10, 10-20 and 20-35 cm horizons. Soil was sieved to 2

mm to remove buried plant residue (i.e. stubble) as no gravel or stones were present.

Lysimeters were reconstructed using these soil horizons. Amendments were either

added as a layer at 10 cm depth or incorporated homogenously into the 0-10 cm soil.

Subsoil ('clay') was collected from a clay pit 400 m from the soil sampling site.

Particle size analysis, determined using the pipette method (Bowman and Hutka,

2002), found the clay contained 60.4% clay and 11.9 % silt while the 0-10 cm soil

contained 94.2% sand (Table 3.1). The random powder X-ray diffraction (XRD)

pattern of clay, obtained for the range 0-70° 2θ (Philips PW 1830 X-ray

diffractometer), showed it contained kaolinite and quartz.

Jarrah wood (trunks and large branches; Eucalyptus marginata Donn ex Sm.) was

pyrolysed at 600 °C for 24 h (Simcoa Ltd, Bunbury, WA). This biochar has a specific

surface area of 273 m2 g-1, determined using a surface area analyser (Micrometrics

Gemini 2375 instrument with a Vac Prep 061 and N2 as the adsorbate). The XRD

pattern of biochar showed it contained calcite, quartz and amorphous carbon.

Soil, clay, and biochar were analysed for NH4+, NO3

–, and DON by extracting 20 g

with 80 mL 0.5 M K2SO4 and analysing the extracts colourimetrically for NH4+

using

the salicylate-nitroprusside method (Krom, 1980; Searle, 1984) and NO3-

concentration using the hydrazine reduction method (Kamphake et al., 1967;

Kempers and Luft, 1988) on an automated flow injection Skalar AutoAnalyser


80

(San++, Skalar Analytical, The Netherlands). DON contents of the extracts were

determined after alkaline persulfate digestion (Cabrera and Beare, 1993). CEC of soil,

clay and biochar was determined using the compulsive exchange method, without

pretreatment for soluble salts (Rayment and Higginson, 1992). Total C and N of the

soil, clay and biochar were determined by combustion analysis using an Elementar

analyser (vario Macro CNS; Elementar, Germany). Soil, clay and biochar were

analysed for pH in a 4 g soil and 20 ml 0.5M CaCl2 mixture.

Table 3.1: Basic properties of soil (at various depths), clay and biochar used in this

experiment.

Soil Clay Biochar

0-10 cm 10-20 cm 20-30 cm

NO3- (mg N kg-1) 7.2 3.5 1.0 7.3 <0.1

NH4+ (mg N kg-1) 2.7 1.0 0.4 17.7 0.4

Organic N (mg N kg-1) 18.8 9.9 7.7 20.8 8.7

Total C (%) 1.04 0.47 0.19 0.11 75.0

Total N (%) 0.08 0.03 0.02 0.01 0.3

Cation exchange

capacity (cmolc kg-1)

2.08 1.67 1.59 7.43 10.11

pH (1:5 0.5 M CaCl2) 5.3 4.6 4.3 5.5 7.3

Sand (%) 94.2 92.2 90.9 27.8 -

Silt (%) 0.6 2.1 1.5 11.9 -

Clay (%) 5.2 5.7 7.6 60.4 -


81

3.3.2 Experimental design and setup

The treatments for this experiment were; 1) biochar incorporated homogenously into

the 0-10 cm soil layer, 2) clay incorporated similarly, 3) biochar added as a layer at

10 cm depth, 4) clay added similarly, or 5) a control. Treatments (replicated five

times) were applied at 25 t ha-1 and soil (3.10 kg) or soil plus amendment (3.085 kg

soil and 0.015 kg amendment) was added to each lysimeter pot (8.75 cm diameter, 36

cm depth). The 10-20 cm and 20-35 cm soil layers were added to a bulk density of

1.5 g cm-3 and the 0-10 cm soil to 1.4 g cm-3 to replicate values measured in the field.

Since this soil naturally compacts to a bulk density between 1.4 and 1.6 g cm-3

biochar and clay application did not cause any difference in the final bulk density of

the pots. Nitrogen fertiliser was added in the form of (NH4)2SO4 at a rate of 40 kg N

ha-1 (equivalent to 7 mg N kg-1 soil). To avoid preferential flow all soil was moistened

to 40% WHC during construction of the soil layers in the lysimeters. Glasshouse

mean maximum and minimum temperatures for the experiment were 31 and 18°C,

respectively.

Lysimeters were watered with the equivalent to 30 mm of rainfall equivalent, daily

for the first 10 days, then on days 13, 15, 17 and 20. The day after watering, leachate

was collected and volume assessed, then filtered (Whatman No. 42) and frozen prior

to analysis. Samples were analysed for NH4+

, NO3- and DON as stated previously.


82

3.3.3 Nitrogen sorption characteristics

Sub-samples (10 g) of biochar, soil, or clay and 100 mL of inorganic N solution were

added to a 120 mL vial in triplicate. N solutions were added as NH4NO3 at

concentrations of 0, 5, 10, 20, 30, 50 and 100 μg N L-1 (0, 50, 100, 200, 300, 500,

1000 μg N g-1 soil). All mixtures were shaken for 24 h, filtered (Whatman No 42) and

analysed using the automated flow injection method.

3.3.4 Water retention

Topsoil (0-10 cm depth), and topsoil incorporated with biochar or clay at 25 t ha-1

was firmly packed to bulk density of 1.4 g cm-3 into a core of 53.8 mm diameter and

10 mm height and saturated with water at atmospheric pressure and its gravimetric

water content determined. Triplicates of identical saturated samples were transferred

to porous ceramic plates, placed in a pressure chambers and subsequently equilibrated

at matric potentials of -10, -100 and -1500 kPa. Gravimetric water contents were

measured after equilibration and then adjusted to volumetric values using bulk

density.


Two analysis methods were used to compare the total quantity of leached NO3-, NH4

+

and DON. An ANOVA was conducted using orthogonal contrasts on total cumulative

leached quantities. Orthogonal contrasts were used to test for differences to the mean

(control vs. each of the four treatments), to compare amendments (biochar vs. clay),

to compare incorporation methods (homogenous incorporation vs. a layer) and to test


83

for interaction. A repeated measures ANOVA was also used. Statistical analysis was

conducted in Genstat 10th edition (Lawes Agricultural Trust, 2007). Differences in N

sorption and water retention were tested using ANOVA and multiple comparisons

using Tukey’s 95% confidence intervals. Statistical significance was considered at

5% (P<0.05).

3.4 Results

3.4.1 Nitrogen leaching

Over the duration of this experiment 15.0 ± 0.6 mg pot-1 of NH4+-N was leached from

the control treatment (Fig. 3.1a). Leaching of NH4+-N significantly decreased with the

addition of biochar (an average of 12.9 mg pot-1) and clay (an average of 12.1 mg

pot-1) (P=0.001; Fig. 3.1a), but there was no significant difference between

amendment types (P=0.323). There was also no difference between incorporation

methods (P=0.671) and there was no interaction (P=0.321; Fig. 3.1a).

Cumulative leaching of NO3--N significantly decreased from 19.5 mg pot-1 in the

control treatment, to an average of 16.3 mg pot-1 with the addition of clay and to an

average of 14.0 mg pot-1 with biochar amendment (P<0.001; Fig. 3.1b). Biochar

amendment decreased NO3- leaching to a significantly greater extent than the clay

amendment (P=0.034). There was no difference between incorporation method

(P=0.300) and no interaction (P=0.930).


84

Cu

mu

lativ

e N

H

4+ -N

1

le

ach

ed

(m

g p

ot

-1)

C

um

ula

tive

NO

3- -N

1

l

ea

che

d (

mg

po

t-1

)

Days

0

5

10

15

Control

Clay Homogenous

Clay Layer

Biochar Homogenous

Biochar Layer

(a)

0

5

10

15

20

0 7 14 21

(b)

Fig. 3.1: Cumulative quantity of NH4+ (a) and NO3

- (b) leached over 21 days for the

unamended soil (control; solid squares), clay homogenously incorporated into the top

10 cm of the soil (solid triangle), a clay layer at 10 cm depth (hollow triangle),

biochar incorporated homogenously (solid circle), or a biochar layer at 10 cm depth

(hollow circle). Error bars represent standard errors (n=5).


85

However, by contrast, no significant difference could be found between any of the

treatments with DON leaching (data not shown). The repeated measures ANOVA

showed significant differences in the amount of NO3- (P=0.004) and NH4

+ (P=0.010)

leached but not DON (P=0.501).

3.4.2 Nitrogen sorption characteristics

At all added N concentrations Tukey’s 95% confidence intervals determined that

biochar sorbed the greatest quantity of both NH4+ and NO3

- (Fig. 3.2a; P<0.001 for all

NH4+ and NO3

- concentrations). Biochar sorbed 75% of the NH4+ in solution when

added at the lowest concentrations (2.5 and 5 μg NH4+-N L-1), decreasing to 54%

when added at the greatest concentration (50 μg NH4+-N L-1; Fig. 3.2a). Tukey’s 95%

confidence intervals showed this was always greater than occurred in the soil or clay.

By comparison, the greatest proportion of NH4+ sorption to the clay (32%) occurred

at a concentration of 20 μg NH4+-N L-1 (Fig. 3.2a).

A much greater difference was seen between the sorption characteristics when the

materials were amended with NO3-. While clay exhibited no capacity to sorb NO3

--,

biochar sorbed 80% of the added NO3--N at lower concentrations (2.5- 5 μg NO3

--N

L-1; Fig. 3.2b) decreasing to 38% at the greatest addition rate (50 μg NO3--N L -1; Fig

3.2b).


86

NH

4+ So

rptio

n (μ

g N

H 4+ -N

g-1

so

il)

N

O 3- S

orp

tion

(μg

NO

3- -N g

-1 s

oil)

Initial solution concentration (μg N g-1 soil)

0

50

100

150

200

250

0 100 200 300 400 500

Soil

Clay

Biochar

(a)

0

50

100

150

200

250

0 100 200 300 400 500

(b)

Fig. 3.2: Amount of NH4+ (a) and NO3

- (b) sorbed by biochar (solid circle), clay

(solid triangle) or the unamended topsoil (solid squares). Error bars represent

standard errors (n=3).


87

3.4.3 Water retention capacity

When comparing gravimetric water contents, at atmospheric pressure (0 kPa), both

clay and biochar amendments significantly increased the water holding capacity of

the soil (P=0.003), however, there was no difference between the two amendments

(Table 3.2). At an imposed matric potential of -10 kPa there was no significant

difference in soil water content between the treatments. At an imposed matric

potential of -100 and -1500 kPa, both amendments significantly increased gravimetric

water holding capacity (P<0.001 and P=0.05, respectively). The biochar amended

soil at -1500 kPa contained more water than the same treatment at -100 kPa (Table

3.2). This is attributed to the heterogeneity of the biochar.

The volumetric water retention capacity of the soil increased significantly with

biochar and clay amendment at 0 and -100 kPa (P=0.002; Table 3.3). There was no

significant difference between bulk densities for the top soil, clay amended soil and

biochar amended soil treatments (an average of 1.5 g cm-3).


88

Table 3.2: Gravimetric water holding capacity for the surface soil layer (0-10 cm)

either unamended or amended with clay or biochar (25 t ha-1) at a range of soil

matric potentials. Values (g cm-3) are represented as ± standard errors (n=3). Within

columns, means followed by the same letter are not significantly different at P = 0.05.

0 kPa -10 kPa -100 kPa -1500 kPa

Soil (0-10) 31.4 ± 0.3 a 10.8 ± 2.7 a 3.4 ± 0.1 a 2.9 ± 0.1 a

Soil + Clay 33.7 ± 0.3 b 14.0 ± 0.8 a 4.4 ± 0.0 b 3.8 ± 0.0 b

Soil + Biochar 34.8 ± 0.1 b 15.0 ± 1.0 a 4.8 ± 0.1 c 6.6 ± 1.3 b

ANOVA P = 0.003 P = 0.355 P < 0.001 P = 0.05

Table 3.3: Volumetric water holding capacity for the surface soil layer (0-10 cm)

either unamended or amended with clay or biochar (25 t ha-1) at a range of soil

matric potentials. Values (g cm-3) are represented as ± standard errors (n=3). Within

columns, means followed by the same letter are not significantly different at P = 0.05.

0 kPa -10 kPa -100 kPa -1500 kPa

Soil (0-10) 19.0 ± 0.1 a 6.7 ± 1.6 2.1 ± 0.1 a 2.2 ± 0.0

Soil + Clay 20.1 ± 0.1 b 8.8 ± 0.4 3.4 ± 0.1 b 2.8 ± 0.0

Soil + Biochar 20.9 ± 0.1 c 9.5 ± 0.5 3.6 ± 0.1 b 5.0 ± 1.0

ANOVA P = 0.002 P = 0.297 P = 0.002 P = 0.059


89

3.5 Discussion

Chapter 3 results support the hypothesis that both the addition of biochar and clay

decrease N leaching. Previous studies have shown biochar amendments can decrease

NH4+ leaching (Lehmann et al., 2003; Ding et al., 2010). The main mechanisms to

explain this are an increase in the nutrient and water retention capacities of the soil

(Lehmann et al., 2003). Overall, there was not a difference in cumulative NH4+

leaching between the two amendment types despite the large discrepancy in their

NH4+ sorption capacity (Fig. 3.2). As such, improved gravimetric water holding

capacity resulting from the soil amendment is likely to play a comparatively greater

role in decreasing NH4+ leaching than NH4

+ sorption. The improved gravimetric

water holding capacity occurred largely at 0 and -100 kPa suggesting that this

additional water is also plant available. Although there were no plants in these

experiments, this additional water could support greater plant growth which in turn

would lower soil solution N concentrations and indirectly decrease N leaching.

The amount of NO3- leaching also decreased with both clay and biochar addition,

contrary to previous biochar studies, where leaching of this anion increased (Laird et

al., 2010b; Lehmann et al., 2003). In this study, biochar amendment decreased NO3-

leaching significantly more than clay. One reason for the difference is potentially due

to the large difference in anion retention capacity between the two ameliorants. Clay,

lacking NO3- retention capacity, is likely to have decreased leaching simply by

increasing the soil’s inherent water holding capacity, whereas biochar is likely to

have decreased NO3- leaching due to both sorption processes and an increased water


90

holding capacity. The large anion retention is likely to be due to the positive charge of

biochar (Cheng et al., 2008). Alternatively it may be due to changes in nitrification.

The addition of this biochar to soils is known to decrease N mineralisation and

nitrification (Chapter 2). Changes in nitrification might be the reason for both

decreased nitrate leaching in this experiment, and the increased nitrate leaching seen

in others (Lehmann et al., 2003).

In many agricultural regions significant N losses may also occur in the form of DON

(Jones et al., 2005; Murphy et al., 2000). While much of this DON is not available in

soil it can be subsequently transformed in freshwaters by UV photolysis into

biologically available forms of N and hence contribute to eutrophication. In this

experiment, biochar and clay amendments did not decrease DON leaching. As most

DON carries a net negative charge, the case for decreased NO3- leaching due to

decreased nitrification, rather than sorption differences, is strengthened. However

these results also conflict with the idea that increased WHC should aid decreased N

leaching. Suggesting there may be another mechanism influencing NH4+ leaching.

The hypothesis for this experiment was that a layered application method would

provide a more effective barrier to leaching, due to greater interception of leachable

ions and less chance of bypass flow. Surprisingly, the method of clay or biochar

application made little overall difference to N leaching. This contrasts with the results

of Jones et al. (2011) where horizontal banding of biochar reduced the amount of

herbicide leaching in soil in comparison to when the biochar was homogenously


91

incorporated into the soil. As such, land managers should apply biochar or clay in the

manner that is both most practical and convenient.

It should be noted, however, that over time, as biochar oxidises the net positive

charge is likely to decrease, developing a predominantly negative charge (Cheng et

al., 2006; Cheng et al., 2008). Similarly, the anion exchange sites may become

occupied with stronger sorbing inorganic or organic ligands (e.g. humic substances,

PO43-) reducing its capacity to retain NO3

- or the surface may become blocked due to

biofilm formation. As such biochar application may not provide the same NO3-

leaching amelioration over the long term, but alternatively less NH4+ leaching may

occur. The long term implications on N leaching of biochar in soil therefore requires

further research, especially considering that NO3- is the more mobile of the ions.

Further research is required to ascertain whether the nutrients retained on the soil

amendments are plant available, and whether this is likely to aid N fertiliser use

efficiency in agricultural and horticultural production.

92

Biochar did not change organic N mineralisation Chapter 4

93

Chapter 4

Organic nitrogen mineralisation in two contrasting agro-

ecosystems is unaffected by biochar addition

4.1 Abstract

Biochar additions to soil have been reported to enhance soil fertility whilst

simultaneously storing C. This experiment tested whether either fresh or field-

conditioned (aged) biochar amendment to two contrasting agricultural soils would

alter the mineralisation of organic N compounds. The mineralisation of 14C-labelled

amino acids and peptides were determined over 20 days within each soil. An

exponential kinetic decay model was subsequently fitted to the mineralisation data.

Overall, statistical analysis revealed significant but small differences between the two

biochar treatments and the unamended control treatment. It is concluded that biochar

has very limited impact on the mineralisation rate of low molecular weight dissolved

organic N compounds in these agro-ecosystems.

4.2 Introduction

One method of rapidly sequestering organic C within agricultural systems is the

application of biochar to soil (Sohi et al., 2010). Biochar is the solid products of

thermal decomposition of organic matter, plant or animal, under limited oxygen at

temperatures less than 700°C (Lehmann & Joseph, 2009). Owing to its highly

aromatic structure (Schmidt & Noack, 2000), biochar can remain stable in soil for


94

thousands of years (Preston & Schmidt, 2006) and has proved agronomically

advantageous in some situations (e.g. Chan et al., 2008; Van Zwieten et al., 2010).

Biochar can also impact soil fertility by providing a habitat for microbes (Pietikäinen

et al., 2000), changing soil pH, and sorbing compounds such as pesticides (Yu et al.,

2006; Bornemann et al., 2007; Cao et al., 2009; Jones et al., 2010) and nutrients

(Mizuta et al., 2004; Ding et al., 2010). In forest soils, biochar addition can stimulate

mineralisation and nitrification (Berglund et al., 2004; Wardle et al., 2008) as

phenolic compounds may inhibit these processes (DeLuca et al., 2006). In

agricultural soils this may not be the case as biochar additions have induced decreases

in mineralisation of both C and N (Kuzyakov et al., 2009; Jones et al., 2011; Chapter

3). Mineralisation of organic matter supplies as much as 80% of crop N within

Australia (Angus, 2001), although, globally fertilisers supply a larger proportion on

crop N (50%; Jenkinson, 2001). Nonetheless organic matter mineralisation remains

integral for crop N supply. Despite this, very little research has targeted the

interaction between organic N compounds and biochar.

The implications of this interaction are likely to change over time. By altering

enzyme function, sorption mechanisms are also likely to influence changes in DON

mineralisation (Bailey et al., 2011). As biochar slowly oxidises over time, surface

functional groups change as the particle gains greater negative charge, increasing its

hydrophilicity (Cheng et al., 2006; Cheng et al., 2008). Correspondingly, the

behaviour of the dominant DON compounds in soil (e.g. peptides and amino acids)

may be more likely to be influenced by the sorption properties of biochar after aging.


95

Therefore the hypothesis for this experiment was that in agricultural soils, fresh

biochar addition would not change the mineralisation of small organic N compounds,

but that aged biochar amendment would decrease it.


4.3.1 Experimental design and characterisation of soils and biochars

This experiment used soils from two biochar field experiments contained within two

contrasting agricultural environments. Three soil treatments (control, field-

conditioned “aged” biochar and fresh biochar), and three 14C labelled compound

treatments (an amino acid mix, alanine or the peptide trialanine) were applied. Site 1

was located in Buntine, WA, (30°00’S, 116°21’E) and was where wheat residue

derived biochar (pyrolysed at 450°C for 20 min; Pacific Pyrolysis, Somersby, NSW)

had been applied to an Acidic Yellow Kandosol soil (Isbell, 1996) at a rate of 4 t ha-1

(6 g biochar kg-1 soil; soil bulk density 1.33 g cm-3; 5 cm incorporation depth). This

biochar is comprised of 15% ash, 33% volatile matter, and 44% fixed C and has a

specific surface area of 215 m2 g-1. The replicated field plots (3 × 20 m in size; n = 4)

had biochar applied one year before soil collection. Buntine is located within a

Mediterranean climate zone and the paddock is cropped each winter in a wheat

(Triticum aestivum L.)/wheat/lupins (Lupinus angustifolius L.) rotation, and fallow

over the dry summer. Site 2 was located in Abergwyngregyn, Wales (53°14’N,

4°01’W) where the soil was classified as a Eutric Cambisol (FAO, 1988). Site 2 is a

perennial ryegrass (Lolium perenne L.), crested dogstail (Cynosurus cristatus L.) and


96

white clover (Trifolium repens L.) dominant pasture within a temperate climate.

Mixed hardwood biochar had been applied to the field plots (3 × 6 m in size; n = 4) at

a rate of 25 t ha-1 2.5 years before soil collection (20 g biochar kg-1 soil; soil bulk

density 1.2 g cm-3; 10 cm incorporation depth). The biochar is described in detail in

Jones et al. (2010) and was pyrolysed at 450°C for 48 h (BioRegional Homegrown®;

BioRegional Charcoal Company Ltd, Wallington, Surrey, UK). This biochar is 3%

ash, 16% volatile matter and 77% fixed C, whilst the specific surface area is 39 m2 g-

1. At both sites, batches of biochar were held in storage since field application and

these constituted the fresh biochar used in the experiments described below, whilst

the soil collected from the field biochar plots constituted the aged treatment. Soil

collected from the individual field plots (0-5 cm depth) constituted the replicates in

the experiments.

Soil and biochar characteristics are presented in Tables 4.1 and 4.2. Total C and N

were determined by a combustion analyser. Cation exchange capacity was also

determined (Rayment and Higginson, 1992). Soils and biochar were extracted (0.5 M

K2SO4) and analysed colorimetrically for NO3- (Miranda et al., 2001) and NH4

+

(Mulvaney, 1996), and flourimetrically for free amino-acids (FAA; Jones et al.,

2002) and peptides (Hill et al., 2011). K2SO4 extractable organic carbon and total N

(TN) were measured in a Shimadzu Total Organic Carbon-V-TN analyser. The EC

and pH of the soils and biochar were determined in a 1:5 (w/v) soil-to-water extract.

Soil solution was collected by centrifugal-drainage (Jones et al., 2002) and

characterised as described above.


97

As the Australian soil was dry when collected, prior to use, these soil samples were

moistened to 70% of their water holding capacity and incubated for 7 d at 20°C. Field

moist Welsh soil was used for the incubation. For the mineralisation assays, 10 g soil

or soil/biochar mixture was placed in 50 cm3 polypropylene tubes at 20 °C.

Application rates of the fresh biochar to previously unamended soil corresponded

with field rates. Uniformly labelled 14C-labelled organic N compounds (100 μM; 2

kBq ml-1) comprising either a mixture of 16 amino acids, alanine or trialanine were

then added to the soil. To capture respired 14CO2, a 1 M NaOH trap (1 ml) was added

to each polypropylene tube and the traps replaced after 0.5, 1, 3, 6, 12, 24, 48, 120,

180, 240, 360 and 480 h.

4.3.2 Statistical Analysis

A repeated measures ANOVA was conducted on rates of 14CO2 efflux in GENSTAT

10th edition (Lawes Agricultural Trust, 2007). A double exponential decay model was

also fitted to the mineralization data where

S = Y0 + Y1 × exp(-k1 t) + Y2 × exp(-k2 t) (1)

and where S it the % of the 14C-label remaining in the soil, Y0 is the pool of

recalcitrant microbially synthesised 14C from the added substrate (Hill et al., 2011),

Y1 represents the size of the primary mineralisation pool respired by the microbial

biomass, k1 is the exponential rate co-efficient describing Y1, Y2 represents the size of

the pool initially incorporated into the microbial biomass and subsequently respired,


98

k2 is the exponential rate co-efficient describing the Y2 pool, and t is time. This model

complies with the theory that substrate mineralisation follows a biphasic pattern

(Chotte et al., 1998; Saggar et al., 1999; van Hees et al., 2005; Boddy et al., 2007),

and a certain proportion of added substrate is recalcitrant (Hill et al., 2011). The mean

residence time (MRT) of the labile fraction of the substrate was calculated

numerically where

MRT = (Y1/100)/k1 + (Y2/100)/k2. (2)

Significant differences between model parameters and MRT were assessed by

ANOVA with Tukey’s pairwise comparison with a p < 0.05 threshold for

significance. Due to the contrasting agro-ecosystems, biochar type and rates varied

between soil types; as such statistical comparisons were only made within each soil

type.

Tab

le 4

.1:

Pro

pert

ies

of

soil,

so

il w

ith a

ged

bioc

ha

r am

end

men

t, a

nd

fresh

bio

char

pro

per

ties.

Val

ues

rep

rese

nt

mea

ns ±

SE

M (

n=

4).

S

ite 1

(A

ust

ralia

)

Site

2 (

Wa

les)

S

oil

Age

d b

ioch

ar

am

en

ded

soil

Bio

char

Soi

l A

ged

bio

char

am

en

de

d so

il

Bio

char

Tot

al N

(%

) 0

.08

± 0.

01

0.0

8 ±

0.0

1 2

.32

± 0.

02

0.

26 ±

0.0

1 0

.24

± 0.

01

0.6

9 ±

0.0

1

Tot

al C

(%

) 0

.96

± 0.

07

1.0

3 ±

0.0

8 60

.9 ±

0.4

2.

83 ±

0.4

1 3

.23

± 0.

76

85

.1 ±

0.4

Ext

ract

abl

e C

(m

g C

kg-1)

172

± 1

2 1

75

± 5

40

54 ±

254

70 ±

11

56

± 4

16

53 ±

34

9

Ext

ract

abl

e N

(m

g N

kg-1)

18

.5 ±

1.2

18

.5 ±

0.4

21

1 ±

13

25

.6 ±

14.

1 1

7.5

± 4.

7 8.

5 ±

2.7

Pe

ptid

es (μ

mo

l kg-1

) 5

94

± 1

24

78

5 ±

24

2 2

367

± 5

89

4

12 ±

13

5 7

21 ±

30

7 1

388

± 2

14

Am

ino

aci

ds (

mg

N k

g-1)

1.2

1 ±

0.0

6 1

.31

± 0.

07

1.37

± 0

.12

0.

96 ±

0.0

5 1

.28

± 0.

24

0.1

1 ±

0.0

2

NH

4+ (m

g N

kg-1

) 3

.70

± 0.

38

3.8

7 ±

0.7

6 1

.42

± 0.

21

15

.0 ±

14.

2 5

.16

± 3.

50

0.6

3 ±

0.4

5

NO

3- (m

g N

kg-1

) 2

.02

± 0.

31

1.7

4 ±

0.3

0 1

.05

± 0.

05

26

.8 ±

12.

2 3

0.7

± 1

2.6

0.2

4 ±

0.0

8

pH

(1

:5 H

2O)

6.2

4 ±

0.0

9 6

.55

± 0.

10

9.2

3 ±

0.0

2

6.27

± 0

.13

6.1

4 ±

0.15

8

.41

± 0.

15

EC

(μS

cm

-1)

82

.5 ±

13.

6 8

7.3

± 1

1.6

29

80 ±

27

0

37.4

± 2

.32

65

.1 ±

17.

9 38

8 ±

67.

5

Cat

ion

Exc

ha

nge

Cap

acity

(m

mol

kg

-1)

11

.8 ±

0.3

18

.6 ±

1.0

34.9

± 2

.1

85.

9 ±

2.7

76.

4 ±

5.6

9.5

± 2.

9

So

il re

spir

atio

n (m

g C

O 2-C

kg-1

h-1)

1.8

4 ±

0.3

0 2

.11

± 0.

22

n.d

.

2.55

± 0

.35

3.2

7 ±

0.25

n

.d.

Tab

le 4

.2:

So

il so

lutio

n p

rop

ert

ies

from

th

e s

oil

an

d so

il w

ith

age

d b

ioch

ar

am

end

men

t. V

alu

es r

epre

sent

mean

s ±

SE

M (

n=

4).

Pro

per

ties

Site

1 (

Au

stra

lia)

S

ite 2

(W

ale

s)

S

oil

Bio

char

am

end

ed

soil

S

oil

Bio

char

am

end

ed

soi

l

Dis

solv

ed

org

an

ic C

(m

g C

l-1)

106

± 9

108

± 1

9

82

± 8

98

± 1

3

Tot

al d

isso

lve

d N

(m

g N

l-1)

51

.4 ±

6.8

61

.7 ±

12.

6

21.6

± 1

0.3

19.

7 ±

6.8

Pe

ptid

es (

µm

ol l-1

) 3

6.2

± 1

.4 3

0.6

± 4

.6

42.0

± 1

8.5

11.

2 ±

6.6

Am

ino

aci

ds (

µg

N l-1)

18

3 ±

25

173

± 3

0

102

± 2

4 2

30

± 9

0

NH

4+ (m

g N

l-1)

0.48

± 0

.10

0.4

6 ±

0.1

6

6.59

± 4

.42

1.3

5 ±

0.8

8

NO

3- (m

g N

l-1)

48

.1 ±

6.

3 3

5.8

± 6

.1

7.4

± 4

.5 7

.6 ±

3.9


101

4.4 Results

Repeated measures ANOVA showed that biochar addition only induced a significant

difference in total 14CO2 evolution in the trialanine treatment in the Australian soil

(Fig. 4.1). Similarly, the kinetic model parameters describing mineralisation also

failed to show consistent treatment differences in C partitioning within the microbial

biomass (Y0, Y1, Y2) or rates of C turnover after microbial assimilation (k1, k2) in the

two soils (Tables 4.3, 4.4, 4.5 and 4.6). When significant differences were observed

they tended to be small. For example, in the Australian soil amended with alanine, Y2

was greater in the aged biochar treatment (22.3%) compared to the control (19.2%)

and fresh biochar (19.0%) treatments (p=0.001; Table 4.3). Y2 did not differ in the

corresponding Welsh soil treatment, however, Y1 did (p=0.006; Table 4.3). In the

Welsh soil amended with amino acids, k1 was slightly lower when fresh biochar was

applied to the soil (p=0.049) while no significant effect was seen in the aged biochar

treatment (Table 4.4). In contrast this was not seen in the Australian soil (Table 4.4).

In the Australian soil with peptide amendment the size of the recalcitrant pool (Y0)

and respired pool (Y1) in the aged-biochar treatment changed relative to the control

(p=0.02), however, no effect was seen in the fresh biochar treatment (Table 4.5). The

MRT of the substrate labile pool did not change with biochar addition in either soil

type with any of the added substrates (Table 4.6).


102

Fig. 4.1: Effect of biochar treatment on the percentage of 14C evolved as CO2 in soil

over a 20 d incubation period for the Australian (A, C, E) and Welsh (B, D, F) soils

amended with 14C-labelled alanine (A, B), a mix of amino acids (C, D) or trialanine

(E, F). Data points represent means ± standard errors (n=4). Curves represent fits to

the kinetic model.

Tab

le 4

.3:

Su

bstr

ate

ha

lf lif

e a

nd

par

am

ete

rs fro

m t

he m

odels

fit

ted

to

the

deca

y cu

rves

of

14 C

-la

belle

d al

an

ine; s

ign

ifica

nt d

iffere

nce

s

are

indi

cate

d b

y N

S (

no

t si

gn

ifica

nt)

, *

(p<

0.05

),

** (

p<0.

01)

an

d **

* (p

<0

.001

); m

ultip

le c

om

pari

sons r

epr

ese

nted

by le

tters

. Val

ues

repr

ese

nt m

ea

ns

± st

and

ard

err

ors

(n

=4

).

R

eca

lcitr

ant p

oo

l R

esp

ire

d po

ol Im

mob

ilise

d p

ool

Su

bst

rate

hal

f life

Y

0 (%

) Y

1 (%

) k 1

(%

ho

ur-1

) Y

2 (%

) k 2

(%

ho

ur-1

) S

t 1/2 (h

ou

rs)

Aus

tral

ian

Soi

l

Co

ntro

l 5

6.3

± 1.

5 2

4.2

± 1

.2 0

.38

± 0.

04

19

.2 ±

0.4

0 a

0.0

06

7 ±

0.0

006

6.04

± 0

.71

ab

Age

d bi

och

ar 5

3.2

± 0.

7 2

4.3

± 0

.4 0

.39

± 0.

01

22

.3 ±

0.4

3 b

0.0

05

8 ±

0.0

001

6.6

8 ±

0.06

b

Fre

sh b

ioch

ar

56.

9 ±

1.0

24

.2 ±

0.8

0.4

3 ±

0.01

1

9.0

± 0

.36

a 0

.00

65

± 0.

00

03 4

.80

± 0.

13 a

P v

alu

e N

S N

S N

S **

*

NS

NS

Wel

sh S

oil

Co

ntro

l 6

4.2

± 0.

7 2

1.4

± 0

.7 b

1.0

6 ±

0.03

1

4.1

± 0

.71

0.0

09

3 ±

0.0

017

1.6

5 ±

0.11

Age

d bi

och

ar 6

4.1

± 1.

0 2

0.7

± 0

.5 b

0.8

9 ±

0.08

1

4.8

± 0

.70

0.0

07

1 ±

0.0

003

2.2

0 ±

0.30

Fre

sh b

ioch

ar

63.

8 ±

1.3

18

.4 ±

0.5

a 0

.71

± 0.

15

16

.9 ±

1.5

0.0

10

9 ±

0.0

011

5.1

0 ±

2.19

P v

alu

e N

S **

N

S N

S N

S N

S

Tab

le 4

.4:

Su

bstr

ate

ha

lf lif

e a

nd

par

am

ete

rs fro

m t

he m

odels

fit

ted

to

the

deca

y cu

rves

of

14 C

-la

belle

d am

ino

aci

d m

ixtu

re (

n=

4);

sig

nifi

cant

diff

eren

ces

are in

dic

ate

d b

y N

S (

not

si

gn

ifica

nt)

an

d *

(p<

0.05

); m

ulti

ple

com

pariso

ns r

ep

rese

nted

by

lett

ers

. Val

ues

repr

ese

nt m

ea

ns

± st

and

ard

err

ors

(n

=4

).

R

eca

lcitr

ant p

oo

l R

esp

ire

d po

ol

Imm

ob

ilise

d po

ol S

ubs

trat

e h

alf l

ife

Y

0 (%

) Y

1 (%

) k 1

(%

ho

ur-1

) Y

2 (%

) k 2

(%

hou

r-1)

St 1/

2 (h

our

s)

Aus

tral

ian

soil

Co

ntro

l 6

1.0

± 0.

7 1

6.4

± 0

.6

0.46

± 0

.03

21.9

± 0

.3 0

.00

60 ±

0.0

005

22

.9 ±

3.4

a

Age

d bi

och

ar 6

1.6

± 0.

6 1

4.0

± 0

.6 0.

47 ±

0.0

3 23

.5 ±

0.5

0.0

052

± 0

.00

03 4

2.9

± 2

.0 b

Fre

sh b

ioch

ar

60.

6 ±

1.5

15

.9 ±

0.4

0.39

± 0

.02

22.6

± 1

.2 0

.00

53 ±

0.0

003

29

.9 ±

5.5

ab

P v

alu

e N

S N

S N

S N

S N

S *

Wel

sh s

oil

Co

ntro

l 7

3.9

± 0.

9 1

2.7

± 0

.6 0.

93 ±

0.1

3 b

13.0

± 0

.7 0

.00

81 ±

0.0

003

6.3

7 ±

1.6

1 ab

Age

d bi

och

ar 7

3.8

± 0.

1 1

3.5

± 0

.3 0.

66 ±

0.0

5 ab

12.1

± 0

.2 0

.00

71 ±

0.0

005

4.0

5 ±

0.2

2 a

Fre

sh b

ioch

ar

72.

1 ±

1.1

12

.1 ±

0.2

0.53

± 0

.08

a 15

.0 ±

1.4

0.0

090

± 0

.00

08 1

1.8

± 2.

6 b

P v

alu

e N

S N

S *

NS

NS

NS

Tab

le 4

.5:

Su

bstr

ate

ha

lf lif

e a

nd

par

am

ete

rs fro

m t

he m

odels

fit

ted

to

the

deca

y cu

rves

of

14 C

-la

belle

d pe

ptid

es

(tri

alan

ine; n

=4

);

sig

nifi

cant

diff

eren

ces

are in

dic

ate

d b

y N

S (

not

si

gn

ifica

nt)

, *

(p<

0.05

) o

r **

(p

<0

.01;

mul

tiple

co

mpar

iso

ns

repr

ese

nte

d b

y le

tters

.

Va

lues

rep

rese

nt m

ea

ns

± st

and

ard

err

ors

(n

=4

).

R

eca

lcitr

ant p

oo

l R

esp

ired

poo

l Im

mob

ilise

d p

ool

Su

bst

rate

hal

f life

Y

0 (%

) Y

1 (%

) k 1

(%

ho

ur-1)

Y2

(%)

k 2 (

% h

ou

r-1)

St 1/

2 (h

ou

rs)

Aus

tral

ian

Soi

l

Co

ntro

l 4

6.0

± 1.

8 b

43.

4 ±

1.4

a 0

.71

± 0.

06

10

.9 ±

0.6

0.0

05

3 ±

0.0

005

1.4

2 ±

0.14

Age

d bi

och

ar 4

0.5

± 0.

8 a

48.

2 ±

0.5

b 0

.78

± 0.

06

11

.6 ±

0.2

0.0

05

5 ±

0.0

003

1.2

6 ±

0.10

Fre

sh b

ioch

ar

41.

9 ±

1.1

ab

47.

3 ±

1.0

ab

0.7

9 ±

0.0

6 1

0.9

± 0

.4 0

.00

53

± 0.

00

04 1

.22

± 0.

08

P v

alu

e *

* N

S N

S N

S N

S

Wel

sh S

oil

Co

ntro

l 4

6.1

± 1.

1 4

7.1

± 1.

0 1

.57

± 0.

06

6.7

± 0

.2 a

0.0

13

5 ±

0.0

017

ab

0.5

4 ±

0.02

Age

d bi

och

ar 4

7.0

± 1.

9 4

5.1

± 1.

6 1

.49

± 0.

08

7.8

± 0

.7 a

0.0

10

5 ±

0.0

019

a 0

.60

± 0.

04

Fre

sh b

ioch

ar

42.

9 ±

1.3

45.

6 ±

1.6

1.4

5 ±

0.0

5 1

1.3

± 1

.1 b

0.0

24

4 ±

0.0

039

b 0

.67

± 0.

04

P v

alu

e N

S N

S N

S **

N

S N

S


106

Table 4.6: Mean residence time (hours) of the labile component of 14C labelled

substrate added to the various soil treatments. Values represent means ± SEM

(n=4).

Alanine Amino acid mix Tri-alanine

Australian Soil

Control 34.1 ± 5.1 37.6 ± 3.2 22.1 ± 3.5

Aged biochar 38.9 ± 1.4 45.7 ± 3.2 21.8 ± 1.5

Fresh biochar 30.0 ± 1.7 43.2 ± 4.2 21.6 ± 1.7

P value NS NS NS

Welsh Soil

Control 20.7 ± 2.8 16.4 ± 1.2 5.5 ± 0.7

Aged biochar 21.2 ± 1.4 17.5 ± 1.2 9.7 ± 3.7

Fresh biochar 16.2 ± 2.6 16.9 ± 0.3 5.2 ± 0.7

P value NS NS NS


107

4.5 Discussion

The hypothesis for this experiment was that the addition of fresh biochar would not

change the rate of decomposition of organic N compounds, but that aged biochar

would decrease it. The results from this experiment, however, do not support this

tenet as both short and long term biochar addition failed to reveal major shifts in C

turnover or partitioning within the soil microbial biomass. Based on these results

and those of Kuzyakov et al. (2009), biochar application is unlikely to have a great

effect on mineralisation of organic N compounds. These studies, however, contrast

with studies in boreal forests (Wardle et al., 2008), potentially because these

agricultural soils lack significant quantities of phenolic compounds that suppress N

mineralisation but which can be removed in the presence of biochar (DeLuca et al.,

2006). These results contrast with the findings of Jones et al. (2011), who saw

decreased mineralisation of native organic matter. This is likely to be due to more

recalcitrant nature of the organic matter pool examined and the greater application

rates used. Field aging of biochar also had little effect on mineralisation of N

containing compounds. However, it is known that at least for the aged Welsh

biochar that its chemical properties were very different after being applied 3 years

previously from those of fresh biochar (e.g. pH, nutrient retention; data not

presented). With the large discretion in biochar variables between agro-ecosystems

(e.g. biochar rate and field aging time), differences in DON mineralisation were

expected. For example, the biochar application rate is likely to influence how much

sorption processes will influence DON mineralisation. Furthermore, time may

influence soil-biochar interactions. Initially biochar addition will also introduce


108

some labile C (Smith et al., 2010) and over time biochar surfaces may become

occluded by soil mineral particles (Van Zwieten et al., 2009) both potentially

influencing OM mineralisation. Despite this, few differences were seen with its

addition. As such mechanisms cannot be isolated from this experiment, but it

provides an important insight into application differences actually likely within

contrasting field based situations and their consequences.

Field application rates of biochar in these agro-ecosystems are unlikely to greatly

influence the turnover of the intrinsic DON reserves in soil.

109

110

Minimal biochar and N fertiliser interaction in the field Chapter 5

111

Chapter 5

Minimal interaction between wheat chaff biochar and N

fertiliser in a broadacre field experiment

5.1 Abstract

Biochar, the remains of pyrolysed organic matter, is an agricultural amendment that

can sequester carbon and has been found to synergistically interact with N fertiliser.

The potential for this amendment to improve wheat (Triticum aestivum L.) yields and

N fertiliser use efficiency via two application methods (banded or spread) in a

broadacre, semi-arid agro-ecosystem was assessed. Wheat residue biochar was

applied (4 t ha-1 banded, 4 t ha-1 spread, or control), in full factorial design with N

fertiliser (0, 20 or 40 kg N ha-1). The hypotheses were that (1) biochar application

would increase wheat yields (2) biochar application would improve N fertiliser use

efficiency, and (3) there would be no difference between application methods. Soil

ammonium (NH4+) and nitrate (NO3

-) to a depth of 90 cm, microbial biomass (0-20

cm), plant N and plant biomass were measured through the year, as was grain N and

yield. At harvest there was no effect of biochar on plant biomass, but there was an

increase in biomass with biochar application at terminal spikelet. Banded biochar

application (1.7 t ha-1) significantly decreased wheat yields from the control (2.1 t ha-

1) but spread application was not significantly different (1.8 t ha-1). Biochar did not

influence soil inorganic N at seeding, re-seeding, anthesis or harvest, and had a small

influence at terminal spikelet sampling. In this experiment biochar had very little


112

impact on N fertiliser and its efficiency of use, possibly because of the low rates

application.

5.2 Introduction

An avenue for mitigating increased atmospheric C levels is through sequestration

within managed soils. The application of biochar is one proposed method to store C

without changing current agricultural practices dramatically (Sohi et al., 2010).

Biochar has been defined as the remains from the pyrolysis of organic matter at

temperatures greater than 250°C (Lehmann & Joseph, 2009). In addition to C

sequestration, biochar has also been shown to improve soil fertility resulting in

improved yields (Glaser et al., 2002; Atkinson et al., 2010), with improved fertiliser

use efficiency in some cropping systems (Van Zwieten et al., 2010a).

Interaction between biochar amendment and N fertiliser additions can occur in some

farming systems (Chan et al., 2007; Chan et al., 2008; Van Zwieten et al., 2010a), but

mechanisms inducing these changes are unclear. Adding a porous, alkaline

amendment that contains a diverse range of minerals and nutrients to acid soils adds

many confounding factors when trying to isolate mechanisms. Apart from a fertiliser

impact through nutrients added in biochar, it may provide a habitat for the microbial

community (Peitikäinen et al., 2000). Increases in rates of N mineralisation and

nitrification in soil with added biochar have been measured in pine forest ecosystems

(Berglund et al., 2004; DeLuca et al., 2006) but changes induced in agricultural and

grassland ecosystems are variable (e.g. DeLuca et al., 2006; Chapter 2; Chapter 4).


113

Biochar addition can also decrease N leaching by increasing water holding capacity

(WHC; Lehmann et al., 2003; Chapter 3), and nutrient retention capacity and cation

exchange capacity through increased electrostatic sorption (Lehmann et al., 2003;

Liang et al., 2006).

In the broadacre agricultural environment in semi-arid environments the application

rate of biochar is likely to be less then 5 t ha-1 (Blackwell et al., 2010). Contextualised

globally, these application rates are considered low, with moderate application rates

5-15 t ha-1 and large rates greater than 15 t ha-1 (Sohi et al., 2010). Experiments

establishing mechanisms inducing N cycling rate changes have been conducted at

larger application rates (e.g. 191 t ha-1, Lehmann et al., 2003; 1% w/w, DeLuca et al.,

2006; >1.1% w/w Van Zwieten et al., 2010a). The applicability and impact of these

mechanisms has yet to have been tested for broadacre agriculture in semi-arid

environments but its impact is likely to be less as contact between biochar particles

and N compounds will be less.

Soil amendments must guarantee yield improvements and require minimal time and

effort to be feasibly applied in broadacre agriculture (Kingwell, 2011); biochar is no

exception. But many studies examining the interaction between biochar and N

fertiliser have not grown crops to maturity, and not assessed crop yield (e.g. Van

Zwieten et al., 2010a; Van Zwieten et al., 2010b). Furthermore, there are not any

studies that assess the best method of application for farmers of the Western

Australian wheatbelt. Within broadacre agro-ecosystems, the most likely application


114

methods are banding or spreading. Banding is the application of the amendment

within the rooting zone of the plant and spreading is the distribution of the

amendment on top of the soil. Previous laboratory and glasshouse studies examining

application method suggest there may be differences in pesticide dynamics (Jones et

al., 2010), but not N leaching (Chapter 3). Both methods contain weaknesses with

their application. Spreading biochar is dusty and prone to loss from wind, while

banding application rates are much lower and biochar is difficult to handle with

current machinery.

It was hypothesised that biochar application would improve N fertiliser use

efficiency, resulting in increased yields. To test these hypotheses, a field experiment

with three biochar and three N rates in full factorial design was designed and

conducted.


5.3.1 Experimental site and design

The field experiment was located on an Acidic Yellow Kandosol (Table 5.1; Isbell,

1996), near Buntine, WA (30°00’S, 116° 21’E). This site had been cropped with

canola (Brassica napus) the previous year and was seeded with wheat (Triticum

aestivum cv. Wyalkatchem) at 75 kg ha-1. This area is located within a mediterranean-

type climate and has an average rainfall of 350 mm, of which 260 mm falls in the

growing season (May - October) (Bureau of Meteorology, 2011). Within the year of


115

this trial, the experimental site received 205 mm of rain (Fig. 5.1). Growing season

mean monthly minimum and maximum temperatures ranged from 13-5 ºC and 26-16

ºC (Fig. 5.1).

The experiment consisted of three biochar treatments and three N fertiliser treatments

in a full factorial design arranged in a randomised block with four replicates. Each

plot was 2 m by 20 m. N fertiliser (urea) treatments were i) control (0 kg N ha-1); ii)

20 kg N ha-1; or iii) 40 kg N ha-1. The three biochar treatments were; 1) the control (0

t ha-1); 2) biochar applied below the surface of the soil at 5-15 cm depth (4 t ha-1;

termed the ‘banded’ treatment); or 3) biochar spread on the surface of the soil (4 t ha-

1; termed the ‘spread’ treatment). Biochar treatments were applied using a cone

seeder one month prior to seeding. The banded treatment was applied by distributing

the biochar through the seeding and fertiliser boots on the tyne of the cone seeder

within a depth range of 50 to 120 mm. For the spread treatment, distribution hoses

were removed from the boots on the tyne, and secured next to the tyne at a height that

ensured biochar was applied to surface. The control treatment was also cultivated in

the same manner, thus ensuring consistent cultivational influences between all three

treatments. All N treatment and basal fertilisers were applied when seeding. All

treatments received 4.5 kg P ha-1, 5 kg K ha-1 and 10 kg Ca ha-1.

The experiment, due to the large ryegrass burden, was sprayed with 3 L ha-1

glyphosate, and re-seeded 3 weeks after initial seeding. No further fertiliser

amendments were applied at this time.


116

Mo

nth

ly r

ain

fall

rece

ived

1

(m

m)

Mea

n m

on

thly

1

1

2

T

emp

erat

ure

(ºC

)

0

10

20

30

40

50

Apr May Jun Jul Aug Sep Oct Nov

0

10

20

30

40

Fig. 5.1: Monthly Rainfall received (mm; histogram) and mean monthly minimum

(triangles) and maximum (circles) temperature (ºC) received over the year of the

trial. Note: January data not available.

The biochar applied was a wheat (Triticum aestivum) chaff biochar pyrolysed at 450

°C for 20 minutes (Pacific Pyrolysis, Somersby, New South Wales). Total C and N

were determined by combustion analysis (Table 5.1). The EC and pH of the biochar

was determined in a 1:5 (w/v) soil-to-water mixture (Table 5.1). Random powder X-

ray diffraction pattern of biochar obtained for the range 0-70° 2θ showed the biochar

contained amorphous carbon, sylvite and quartz. The specific surface area and

micropore volume of the biochar were 214 m2 g-1 and 70 cm3 g-1 respectively,

determined by a surface area analyser using the Brunauer, Emmett and Teller (BET)

method (Brunauer et al., 1938).

Tab

le 5

.1:B

asi

c p

rope

rtie

s of

bio

cha

r a

nd s

oil

bef

ore s

eed

ing

(N

A -

No

t A

ssess

ed).

Va

lues

are

mean

s w

ith s

tand

ar

d err

ors

(n=

4).

So

il D

epth

(cm

) 0

-10

1

0-2

0

20

-30

30

-60

60

-90

B

ioch

ar

pH

6

.24

± 0.

09

NA

N

A

NA

N

A

9.2

3 ±

0.0

2

EC

(μS

cm

-1)

82

.5 ±

13.

6 N

A

NA

N

A

NA

2

980

± 2

70

NH

4+ (m

g N

kg-1

)

1.1

± 0

.1

0.3

± 0

.1

0.3

± 0

.1

0.3

± 0

.1

0.3

± 0

.1

1.4

2 ±

0.2

1

NO

3- (m

g N

kg-1

) 7

.8 ±

0.4

2

.6 ±

0.5

1

.8 ±

0.3

0

.7 ±

0.1

N

D 0

±0

1.

05 ±

0.0

5

Mic

robi

al

Bio

mas

s

(mg

N k

g -1)

7.0

3 ±

1.45

3

.95

± 0.

69

NA

N

A

NA

N

A

Tot

al C

(%

) 1

.05

± 0.

06

0.5

6 ±

0.0

3 0

.23

± 0.

01

0.1

3 ±

0.0

1 0

.11

± 0.

01

60

.9 ±

0.4

4

Tot

al N

(%

) 0

.08

± 0.

004

0.0

4 ±

0.0

05

0.0

2 ±

0.0

02 0

.02

± 0.

00

2 0

.02

± 0.

003

2.3

2 ±

0.0

2


118

5.3.2 Sampling times and analysis

Soils were sampled at the seeding, re-seeding, terminal spikelet, anthesis and harvest

stages of plant growth. Soil was samples were collected at 0-10, 10-20, 20-30, 30-60

and 60-90 cm depth. Soil from the 0-10 and 10-20 cm layers was assessed for MBN

using the fumigation-extraction method (Vance et al., 1987) and the ninhydrin

method (Joergensen & Brookes, 1990) and the flush (fumigated less non-fumigated

values) adjusted by a factor of 6.47 (Sparling et al., 1993). Soil extracts were

analysed colourimetrically for ammonium (NH4+) using the salicylate-nitroprusside

method (Krom, 1980; Searle, 1984) and nitrate (NO3-) concentration using the

hydrazine reduction method (Kamphake et al., 1967; Kempers & Luft, 1988) on an

automated flow injection Skalar Auto-analyser (Skalar San plus).

Plant shoot samples were taken at terminal spikelet, anthesis and harvest. These were

analysed for biomass and N concentration. At harvest, grain yield and grain N

concentration were also determined. All plant N content was assessed by combustion

analysis using an elementar (vario Macro CNS; Elementar, Germany).


Statistical analysis was performed using a series of two-way ANOVAs using

GENSTAT 10th Edition (Lawes Agricultural Trust, 2007). Multiple comparisons

were done using Tukey’s 95% confidence intervals. Significance considered at the P

< 0.05 significance level.


119

5.4 Results

5.4.1 Soil microbial biomass

The size of the MBN was 19.2 μg N g-1 soil at terminal spikelet sampling in the 0-10

cm layer. No difference in MBN, due to either biochar or N addition, was detected at

any sampling time or analysed soil depth (0-10 or 10-20 cm).

5.4.2 Soil ammonium and nitrate

There was no effect of biochar application or interaction between biochar application

and N fertiliser at re-seeding. At re-seeding statistical differences in NH4+ at 0-10 cm

and 10-20 cm and NO3- at 0-10 cm depth were caused by N application, where the

greatest concentrations NH4+ and NO3

- could be found within the soils applied with

the most N (Table 5.2).

At terminal spikelet, an interaction was found between the two factors for NH4+ at 10-

20 cm depth, but this was the only instance in all soil samples in which statistical

interaction between the two factors was present. At 20-30 cm depth, in the terminal

spikelet sampling there was a significantly more NH4+ was wheat 40 kg N ha-1 was

applied than 0 kg N ha-1 (Table 5.2). The only significant differences in soil N with

biochar addition were in NO3- at 20-30 cm depth and in NH4

+ at 60-90 cm depth

during terminal spikelet sampling. There was significantly more nitrate detected in

the spread treatment (1.69 mg N kg-1) than in the control (0.77 mg N kg-1) at 20-30

cm depth and less NH4+ detected in both biochar amended treatments (banded - 0.09


120

mg N kg-1; spread - 0.01 mg N kg-1) than the control (0.20 mg N kg-1) at 60-90 cm

depth. At anthesis and harvest significant differences in soil NH4+ and NO3

- were

only due to the application of N fertiliser (Table 5.2).

5.4.3 Plant biomass and nitrogen content

At terminal spikelet plant biomass was affected by both N application (P<0.001) as

well as biochar application (P=0.027), but there was no interaction between

amendments (Table 5.3). Increasing N application significantly increased plant

biomass. Plant biomass was significantly increased with the spread application of

biochar (from 0.60 to 0.72 t ha-1) but not the banded application (Fig. 5.2a).

Interaction between biochar and N applications was present at anthesis sampling

(P=0.015). At this time point, the greatest plant biomass was in the 40 kg N ha-1 and

banded biochar treatment (5.53 t ha-1) and the least plant biomass was in the banded

biochar and 0 kg N ha-1 treatment (3.83 t ha-1; Fig. 5.2b).

At harvest there was no difference in plant biomass due to biochar, and there was no

interaction. N application at 40 kg ha-1 increased plant biomass from 4.70 t ha-1 to

5.69 t ha-1 (P=0.016; Table 3; Fig. 5.2c).

Tab

le 5

.2:

Soi

l N

O 3- a

nd N

H 4+ v

alu

es

(mg

N k

g-1 s

oil)

fo

r sa

mpl

es

influ

enc

ed

by

N a

pplic

atio

n.

Val

ues

rep

rese

nt

mea

ns w

ithin

ea

ch

bio

cha

r tr

ea

tmen

t ±

sta

nda

rd e

rror

s (n

=4)

, le

tter

s re

pre

sent

sig

nifi

cant

diff

ere

nce

s.

Sa

mp

ling

Tim

e R

e-se

edin

g

Re-

seed

ing

R

e-se

edin

g

Ter

min

al

Sp

ikel

et

An

thes

is

An

thes

is

Har

vest

Dep

th (

cm)

0-1

0

0-1

0

10-

20

2

0-3

0

0-1

0

10-

20

0

-10

N t

ype

N

H 4+

NO

3- N

H4+

NH

4+ N

H4+

NH

4+ N

H4+

P v

alu

e

0.0

32

<0

.00

1 0

.009

0

.029

0

.019

0

.014

0

.016

40

kg N

ha-1

0

.35

± 0.

10 a

5

.54

± 0

.50

a

0.3

7 ±

0.3

5 a

0

.71

± 0

.2

0 b

1

.01

± 0

.14

b

0.7

7 ±

0.2

7 b

1

.11

± 0

.10

b

20

kg N

ha-1

5

.36

±1.7

3 a

b

8.5

8 ±

1.4

2 a

1

.37

± 0

.59

a

0.3

2 ±

0.1

0 a

b

0.8

0 ±

0.1

0 a

b

0.5

2 ±

0.0

6 a

b

1.1

0 ±

0.0

8 b

0 k

g N

ha-1

1

4.59

± 6

.41

b

12.

38 ±

1.6

4 b

5

.24

± 1

.87

b

0.1

3 ±

0.0

4 a

0

.60

± 0

.04

a

0.3

0 ±

0.0

5 a

0

.81

± 0

.08

a


122

Table 5.3: Table of significant differences for plant biomass, plant N and grain yield

and N for all measured time points. P values for significant differences due to

biochar (*), N application (^) or interaction (†) are shown; NS represents not

significant.

Terminal Spikelet Anthesis Harvest

Plant Biomass (t ha-1) 0.027 *; <0.001 ^ 0.015 † 0.016 ^

Plant N (%) <0.001 ^ <0.001 ^ <0.001 ^

Grain Yield (t ha-1) - - 0.042 *

Grain N (%) - - <0.001 ^

At all measured time points, there was no significant effect of biochar on plant N

concentration and there was no interaction between biochar and N application.

Increasing N application always increased plant N concentration (P<0.001 at all time

points).

5.4.4 Grain yield and nitrogen content

Grain yield decreased with banded biochar application (P=0.042). The control plot

yielded 2.1 t ha-1, the spread biochar plots yielded 1.8 t/ha and the banded biochar

plots yielded 1.7 t ha-1 (Fig. 5.2d). Yield was not influenced by N application and

there was no interaction between biochar and N fertiliser.


123

P

lan

t Bio

mas

s (t

ha

-1)

Pla

nt B

iom

ass

(t h

a-1)

Pla

nt B

iom

ass

(t h

a-1)

Gra

in Y

ield

(t h

a-1)

0

0.2

0.4

0.6

0.8

1

(a)

0

1

2

3

4

5

6

(b) a

aaaa

bb

bb

bc

ccd d

0

1

2

3

4

5

6

7

Con

trol

Ban

ded

Spr

ead

Con

trol

Ban

ded

Spr

ead

Con

trol

Ban

ded

Spr

ead

0 kg N ha 20 kg N ha 40 kg N ha

(c)

Time

0

0.5

1

1.5

2

2.5

3

Con

trol

Ban

ded

Spr

ead

Con

trol

Ban

ded

Spr

ead

Con

trol

Ban

ded

Spr

ead

0 kg N ha 20 kg N ha 40 kg N ha

(d)

Fig. 5.2: Dry plant biomass at terminal spikelet (a), anthesis (b) and harvest (c) and

grain yield (d). Error bars represent standard errors (n=4).

The percentage of N in the grain was not affected by biochar application, but

increased with increasing N application quantities (P<0.001). There was no

interaction between N fertiliser and biochar application.


124

5.5 Discussion

5.5.1 Interaction between nitrogen fertiliser and biochar

The interaction between biochar and N fertiliser has been suggested to be an

important reason for agronomic improvement resulting in increased yields (Chan et

al., 2007; Chan et al., 2008; Van Zwieten et al., 2010a). It has also been suggested

that biochar can provide nutrients other than N to plants (Chan et al., 2007; Chan et

al., 2008). Phosphorus and potassium, within the biochar, may have been plant

available in the early part of the growing season and thus contributed to the greater

plant biomass seen at terminal spikelet sampling with the spread application of

biochar (Fig. 5.2a). Although the plant samples were not tested for any nutrients other

than N, the decreased yields seen at harvest suggest this is unlikely.

Increased N fertiliser use efficiency has been seen with large application rates of

biochar; greater than 10 t ha-1 for nutrient rich biochars, and even greater rates for

nutrient poor biochars (Chan et al., 2007; Chan et al., 2008; Van Zwieten et al.,

2010a; Van Zwieten et al., 2010b). Given the low bulk density of biochar (0.2 g cm-3

in this study), application rates above 4 t ha-1 would be logistically difficult,

expensive and time consuming. Therefore it is unlikely that significant improvements

in N fertiliser use efficiency as seen in previous research (Chan et al., 2007; Chan et

al., 2008; Van Zwieten et al., 2010a) are to be achieved in broadacre agricultural

environments.


125

5.5.2 Yield influence of biochar

Previous research has suggested that application of biochar frequently increases plant

and/or crop yield (Glaser et al., 2002; Atkinson et al., 2010; Blackwell et al., 2010),

however in this study biochar application resulted in decreased wheat yields

(significant only where biochar was banded). Biochar may have decreased plant yield

by limiting the rate of N mineralisation (Chapter 2). However using soil collected

from this experiment, organic N mineralisation did not change with biochar addition

(Chapter 4). Both that study and the current study did not assess changes in

nitrification rates or urea hydrolysis rates as a result of biochar addition, but they are

unlikely to have changed upon biochar addition analogous to organic N

mineralisation. Furthermore, there was no significant difference in grain N,

suggesting the yield difference maybe due to another factor, potentially a change in

soil nutritional status (Chan et al., 2008; Blackwell et al., 2010).

One mechanism suggested for this is by enhancing plant water supply by increasing

the WHC of the soil (Atkinson et al., 2010; Blackwell et al., 2010). This is likely to

be especially relevant in dry environments with coarse-textured soils; however the

current study suggests this mechanism is unlikely, especially at field applicable

application rates in this agro-ecosystem. This experiment received 195 mm of rainfall

in between February and October of the year, yet moisture content did not change

with biochar addition at any measured time point (data not shown). If WHC of the

biochar is 400% (determined in a small packed core, data not shown), 4 t ha-1 may be

able to hold a maximum of 1.6 mm of rainfall extra at any time. A water use


126

efficiency of 12.7 kg ha-1 grain mm-1 (French & Shultz, 1984) suggests that this could

lead to an increased 20.3 kg ha-1 assuming that all water is plant available and this

increased WHC is only fully utilised once per growing season. As yield decreased

with biochar application (significant decreases with banded biochar application),

biochar is unlikely to have affected this experiment by increasing WHC.

5.5.3 Biochar application method

Biochar application method did influence the effect of biochar in some instances,

although the overall influence of biochar was limited. This was seen with plant

biomass at the terminal spikelet stage where plant biomass was greatest with spread

application of biochar (Fig. 5.2a) and at harvest, where grain yield was lowest with

banded application of biochar (Fig. 5.2d). In both instances, the other biochar

treatment did not significantly change the assessed parameter. This demonstrates that

varying application methods may alter the crop influence of biochar. These results

suggest that application of biochar to the surface of the soil may be more effective

than subsurface application, but further research is required replicate these effects

establish whether the effect is consistent.

5.5.4 Conclusion

In a semi-arid, broadacre environment the application of biochar did not provide

agronomic benefit in the first season of application. Grain yield was decreased with

biochar application. This decrease was significant only when the biochar was banded,

as such spread application appeared less agronomically detrimental than banded


127

application. Interaction between biochar and N fertiliser was sporadic and minimal.

Furthermore, N grain and shoot N content did not change with biochar addition, as

such this biochar did not improve N fertiliser use efficiency.

128

General Discussion Chapter 6

129

Chapter 6

General Discussion

Biochar has previously been preposed to influence soil fertility via a number of

changes to soil physical, chemical and biological properties. By focusing on the

interaction between biochar and the terrestrial N cycle a number of questions have

been addressed in this thesis. These questions revolved around changes to N cycling

rates that are likely to influence on farm productivity, specifically N mineralisation, N

immobilisation and N leaching (Summarised in Table 6.1). The experiments

investigating these questions used biochar types with potential to be used in the

Western Australian wheatbelt. Biochar either decreased or did not change N

mineralisation or the soil microbial biomass. However biochar did decrease N

leaching. This chapter discusses five specific processes that impact (directly or

indirectly) terrestial N cycling, and consequently farm productivity. The specific

processes are: the sorption of LMW N compounds to biochar, the sorption of high

molecular weight (HMW) organic compounds to biochar, the addition of labile C

contained in biochar, and increased WHC resulting from biochar application.


130

Table 6.1: A summary of the questions addressed within each thesis chapter, and the

corresponding answers found in this thesis.

Chapter & Questions Addressed Answers to questions

Chapter 2:

� When added to a coarse textured soil, does

biochar sorb NH4+, NO3

- or amino acids?

� Does biochar influence net N

mineralisation when added to a coarse

textured soil?

� Does biochar addition to a coarse textured

soil induce microbial immobilisation of N?

� No.

� Yes, Jarrah biochar

decreased net N

mineralisation.

� No.

Chapter 3:

� Does biochar addition to a coarse textured

soil decrease N leaching?

� Is sorption the main mechanism

decreasing leaching when added to a

coarse-textured soil?

� Yes, Jarrah biochar did.

� No, not the main

mechanism, maybe due

to increased WHC.

Chapter 4:

� Does organic N mineralisation change

when biochar is added to a coarse textured

soil?

� Does organic N mineralisation change

over time after biochar addition to a coarse

textured soil?

� No, not with wheat

residue biochar.

� No, not with wheat

residue biochar.


131

Chapter 5:

� When added to a coarse textured soil in a

broadacre Western Australian agro-

ecosystem, does biochar interact with N

fertiliser to increase soil N?

� Does biochar influence plant N uptake

when added to a coarse textured soil in a

broadacre Western Australian agro-

ecosystem?

� Generally no, but

interaction between

biochar and N fertiliser

was found during one of

five sampling times.

� No difference was found

in plant N uptake with

wheat residue biochar

added at 4 t ha-1.

6.1 Sorption of low molecular weight nitrogen to biochar

Biochar can sorb nitrate (NO3-; Mizuta et al., 2004; Chapter 2; Chapter 3),

ammonium (NH4+; Hina et al., 2010; Chapter 2; Chapter 3), ammonia (NH3;

Taghizadeh-Toosi et al., 2012) and amino acids (Chapter 2). Consequently sorption

processes have been hypothesised to decrease nitrification, mineralisation and N

leaching (Lehmann et al., 2003; Laird et al., 2010b). In this thesis evidence is

provided to suggest that sorption of LMW N compounds (NO3-, NH4

+; NH3; and

amino acids) to biochar does not influence nitrification (Chapter 2) or mineralisation

(Chapter 2, Chapter 4) and is not the main cause of decreased N leaching with

biochar addition to soil (Chapter 3). The sorption of LMW N has been used to explain

results in previous research (e.g. Lehmann et al., 2003; Laird et al., 2010b) yet prior

to this thesis experiments examining LMW sorption had not been conducted.

Consequently results from other experiments may be interpreted differently with this


132

new finding, provided two important caveats: similar biochar chemistry and structure;

and similar biochar rates of application.

As stated in Section 1.5.1, the sorption affinity of biochar will vary between types

due to variations in surface functional groups (Boehm, 1994) and surface area

(Bornemann et al., 2007). Despite their different pyrolysis temperatures and

feedstocks, the two biochar types applied to coarse textured soils in this thesis, Jarrah

and wheat residue biochars, had a similar pH (9.3 and 9.2, respectively) and hence

may have contained similar surface functional groups and hydrophobic properties. It

could be valid to conclude that most biochar types will influence other experiments

similarly if biochars with contrasting characteristics (i.e. lower pH and more acidic

surface functional groups) to the biochar types used in this thesis had similar

influence on the sorption of LMW N compounds. Thus, further research is required to

assess whether the findings of this thesis can be validly extrapolated to contrasting

biochar types, especially with varying pH and acidic functional groups.

The application rate of biochar is likely to influence N sorption, due to the number of

functional groups introduced to soil, increasing the likelihood for sorption. Practical

application rates in broadacre agriculture in WA are likely to be less than 5 t ha-1

(Blackwell et al., 2010). Consequently, in the drier field environment, changes in

leaching with lower rates of biochar application (4 t ha-1) to coarse-textured soil may

more closely reflect the negligible changes in soil N movement down the soil profile

in Chapter 5 than the significant differences in inorganic N leaching demonstrated in


133

a well watered lysimeter experiment with 25 t ha-1 biochar addition (Chapter 3). In

other agro-ecosystems, rates of application may exceed 25 t ha-1 (e.g. Jones et al.,

2010; Sohi et al., 2010); the sorption of N to biochar with greater application rates

may increase its effect on N leaching and mineralisation.

6.1.1 Applying findings to other experiments

With these two caveats in mind, it is possible to use this new information to re-

interprete the results from some previous experiments. Lehmann et al., 2003 used an

application rate of 20% w/w biochar/soil or 191 t ha-1. Re-interpreting the results of

Lehmann et al. (2003) with this very large rate of application is unwise because if

20% of the soil was biochar; biochar is much more likely to intercept and sorb

nutrients and other compounds at this rate of application. Unfortunately only one

experiment could be found which applied biochar at similar rates and assessed LMW

N leaching. Ding et al., (2010) when applying bamboo biochar at 0.5% w/w, or

approximately 5 t ha-1 (using the conversion rates from chapter 3) state that the

decreased NH4+ leaching in their experiment was due to the sorption of NH4

+ to

biochar; but the findings in this thesis (that the sorption of LMW N to biochar is

unlikely to influence leaching) suggest otherwise. An alternate explanation that is

consistent with their findings (Ding et al., 2010) is that nitrification may have

increased, which would have also increased NO3- leaching. Although this data was

measured, it was not presented. Nitrification increases have been implied in other

agricultural eco-types (e.g. Van Zwieten et al., 2010a), although this is contrary to the


134

findings of this thesis where biochar decreased nitrification (Chapter 2), it could

explain lower NH4+ leaching.

6.1.2 Implications of the minimal impact of sorption of low molecular weight

nitrogen to biochar

In this thesis sorption of LMW N to biochar added to coarse-textured soil did not

influence nitrification and mineralisation and was not the main mechanism decreasing

N leaching, raising further research questions. Three such topics for further

investigation are: the main mechanisms decreasing inorganic N leaching with biochar

addition to a coarse textured soil; the consistency of this LMW sorption over time;

and the scale of measurement.

More research is required to pinpoint other mechanisms causing decreased inorganic

N leaching with biochar addition (discussed in Chapter 3). Increased WHC may

decrease NH4+ and NO3

- leaching by providing refuges for ions to avoid movement

by mass flow. Increased WHC should also have decreased DON leaching similarly,

but no difference in DON leaching with soil amendment was found. Thus, research

investigating other possible mechanisms is that decrease N leaching is required.

Although aging of biochar did not influence organic N mineralisation in this thesis

(Chapter 4), whether more oxidation could change the results or whether the results

from the tested agro-ecosystem are representative remains unclear. Over time the

pores of biochar may become blocked with SOM (Zimmerman et al., 2011), termed


135

protective sorption (Kaiser & Guggenberger, 2000), potentially lessening the

increased WHC with biochar addition and associated decrease in inorganic N

leaching.

This thesis measured changes in bulk soil properties, but differences may be more

likely to be measured at the soil-biochar interface, termed the ‘charosphere’ (Clough

& Condron, 2010). The concept of localisation of NO3- within rhizosphere biochar,

caused by mass flow of NO3- towards the root and sorption to rhizosphere biochar

upon interception (Prendergast-Miller et al., 2011), concurs with the findings of this

thesis. Prendergast-Miller et al. (2011) also found that biochar sorbed N,

demonstrated by greater N content in rhizosphere biochar compared to biochar in

bulk soil, but this did not translate into increased plant N uptake. Similarly, Chapter 2

demonstrated that biochar sorbed N, but the sorption was not detected in the bulk soil

when biochar was applied at paddock applicable rates. Thus sorption of LMW N to

biochar may influence N cycling processes such as N mineralisation, nitrification, N

leaching, and plant N uptake within the ‘charosphere,’ but the influence of biochar on

N cycling within the bulk soil is likely to be minimal, especially at applications rates

likely to be applied in broadacre agriculture in WA.

6.2 Sorption of high molecular weight organic nitrogen compounds to biochar

The negative priming that occurs with biochar addition to soil (Cross & Sohi, 2011;

Keith et al., 2011; Jones et al., 2011; Zimmerman et al., 2011) may be caused by the

sorption of HMW organic N or organic C to biochar. One apparent trend in


136

mineralisation with biochar addition over longer periods of time (greater than 2

weeks) is that increasing pyrolysis temperature of biochar increases its the negative

priming effect when added to soil (Cross & Sohi, 2011; Zimmerman et al., 2011).

Another trend is that the negative priming effect is more apparent in soils with high

organic matter contents (Cross & Sohi, 2011; Keith et al., 2011; Zimmerman et al.,

2011). Negative priming over the long term is thought to be due to the sorption of

organic matter (Zimmerman et al., 2011), in particular HMW organic compounds.

Biochar has been shown to strongly sorb pesticides such as simazine, atrazine and

trifluralin, decreasing pesticide leaching and efficacy (Bornemann et al., 2007; Jones

et al., 2010; Nag et al., 2011). There are multiple mechanisms that can explain

sorption of xenobiotics to biochar (Zhou et al., 2010) but common differences

between LMW N and xenobiotics are the presence of halides and polarity (Fig. 6.1).

Halides are unlikely to cause the sorption differences, because although fluorine is the

most electronegative element, oxygen is the second most electronegative element

(Allred & Rochow, 1958). Therefore oxygen (in leucine and alanine, Fig. 6.1) forms

stronger electron donor interactions than chlorine (in simazine and atrazine, Fig. 6.1).

Thus, the non-polarity of some xenobiotics caused by symmetry surrounding a

benzene or triazine ring must induce hydrophobic based interactions resulting in

greater sorptive affinity of non-polar hydrophobic molecules than polar hydrophilic

molecules. Scaled particle theory suggests that non-polar molecules aggregate to

reduce solute cavity surface area (Jackson & Sternberg, 1994) and increase molecular

entropy (Kleber & Johnson, 2010). Where Zimmerman et al. (2011) hypothesised


137

that the negative priming effect of biochar was caused by organic matter sorption; I

suggest that it is due to the sorption of HMW non-polar organic matter. Although it is

clear that organic matter sorption varies between organic matter molecules (Kasozi et

al., 2010) and with addition to soil (Cornelissen & Gustafsson, 2004), the relationship

between organic matter sorption to biochar and the molecular weight and polarity of

the molecule sorbed requires further investigation. It is likely that the greater the non-

polarity of the organic matter the greater the sorption affinity for biochar and that

correlates with the molecular size of the organic matter. Further, it is unclear whether

the sorption of the hydrophobic tail of amphiphilic organic matter molecules change

the net effect of biochar on mineralisation. It is also unclear whether similar sorption-

desorption hysteresis seen with pesticides (i.e. Yang et al., 2004), is also present in

amphiphilic or hydrophobic organic matter.

HMW compounds have been shown to sorb more strongly to biochar produced under

high pyrolysis temperatures (Kasozi et al., 2010), explaining the trend of greater

negative priming with increased pyrolysis temperature shown by Cross & Sohi

(2011) and Zimmerman et al. (2011). High pyrolysis temperature can decrease the

oxygen content of the biochar, increasing its hydrophobicity, and the accessible area

for adsorption of hydrophobic pesticides (Yang et al., 2004). Sorption of xenobiotics

may also be related to pore area of the biochar, in particular micropores providing

sites for high energy sorption (Zhou et al., 2010), where adsorption energies for high

energy sorption sites may decrease with increasing pyrolysis temperature (Pikaar et

al., 2006). Similarly, the sorption of hydrophobic organic matter compounds, like


138

catechol, increase with increasing pyrolysis temperatures (Kasozi et al., 2010)

explaining the link between higher peak pyrolysis temperatures and the negative

priming effect demonstrated by Cross & Sohi (2011) and Zimmerman et al. (2011).

Fig. 6.1: Chemistries of selected compounds studied for sorption interactions with

biochar. Simazine, trifluralin and atrazine are hydrophobic; alanine and leucine are

hydrophilic. The degree of hydrophobicity is likely to influence their sorption affinity

to biochar.

6.2.1 Implications of the sorption of high molecular weight compounds to biochar

There are a variety of implications of the sorption of hydrophobic HMW organic

matter to biochar in soil. The most important of these is decreased organic matter

mineralisation. Although decreased organic matter mineralisation will increase SOM

(Cross & Sohi, 2011), it will also decrease nutrient availability for plant N uptake.

L-alanine L-leucine

Simazine Atrazine Trifluralin


139

The rate limiting step of N mineralisation is the decomposition of HMW organic N to

LWM organic N (Jones et al., 2004). As mineralised N constitutes a large portion (up

to 90%) of total crop N uptake (Angus, 2001; Jenkinson, 2001), sorption of

hydrophobic HMW N to biochar could potentially decrease crop yields. Decreased

mineralisation causes decreased NH4+ content, which will decrease nitrification and

leaching of inorganic N. Biochar also has a varied influence on enzymatic activity,

decreasing β-glucosidase activity (Bailey et al., 2011; Lammirato et al., 2011) and

increasing β-N-glucoseaminidase activity (Bailey et al., 2011). Although enzymes

must be hydrophilic for mobility within solution, there may be areas with contrasting

electronegativity, and hence varying hydrophilicity, amphiphilicity and

hydrophobicity between enzymes which is likely to alter their sorption affinity to

biochar.

6.3 The addition of labile carbon contained in biochar

To help predict the impact of biochar on microbial abundance and activity, greater

characterisation and quantification of labile C in biochar must be conducted.

Generally, the addition of labile C in biochar increases microbial biomass (Lehmann

et al., 2011) and activity (Steiner et al., 2008a) and increasing addition of biochar and

labile C induces larger increases in microbial abundance (Kolb et al., 2009).

However, labile C is highly variable and can range from being readily microbially

available, such as amino acids and sugars (e.g. Jones & Murphy, 2007) to microbially

inhibitive, such as benzene (e.g. Girvan et al., 2004). Despite potential diversity of

labile C present on biochar, it is commonly documented as VM or volatile content;


140

ranging in concentration from 16-400 g VM kg-1 biochar (Table 1.1). A more

thorough characterisation of labile C in pine biochar has demonstrated the presence of

VOCs such as acetaldehyde, α-pinene and β-pinene (Clough et al., 2010). α-pinene

can decrease N mineralisation and MBN (Uusitalo et al., 2008). Yet, when the pine

biochar was added to soil, α-pinene did not influence N mineralisation and microbial

biomass (Clough et al., 2010) probably because of the small quantity of the α-pinene

added. Benzene has also been detected in low concentrations on biochar (Chapter 2),

although the detected concentrations were lower than previously researched known

toxic concentrations (e.g. Girvan et al., 2004). Thus, the quantity and types of labile

C present on biochar are likely to be a primary cause of the differences in N

immobilisation demonstrated with different biochar types (e.g. Gundale & DeLuca,

2007). Varying labile C through changing pyrolysis temperatures (Baldock &

Smernick, 2002; Mukherjee et al., 2011) is also likely to be partially responsible for

changes in N immobilisation with varying biochar types. It is unclear how the

abundance of specific labile compounds change with other factors of production.

Thus, greater assessment and quantification of specific labile C compounds is

required to predict the net influence of biochar on N immobilisation, net N

mineralisation and the longevity of such changes induced by biochar amendment.


141

6.4 Increasing the supply of phosphorus and potassium in biochar

Previous research has demonstrated that biochar can interact with N fertiliser,

enhancing N fertiliser use efficiency (Chan et al., 2007; Chan et al., 2008; Van

Zwieten et al., 2010a). This is likely to have been caused by biochar addition

simultaneously increasing the supply of other nutrients such as P and K (Chan et al.,

2007; Chan et al., 2008). Changes in P and K availability will vary with biochar

nutrient content and application rate. Other studies demonstrating increases in N

fertiliser use efficiency added biochar at large application rates (50 or 100 t ha-1) for

nutrient poor biochars (e.g. greenwaste; Chan et al., 2007) or lower rates (10 t ha-1)

for nutrient rich biochar (eg. poultry manure biochar; Chan et al., 2008). The nutrient

poor greenwaste biochar contained 400 mg Colwell P kg-1 (Chan et al., 2007), thus

addition at 50 t ha-1 would apply 20 kg Colwell P ha-1. The two poultry manure

biochars contained 11600 and 1800 mg Colwell P kg-1 (Chan et al., 2008);

application of 10 t ha-1 would apply 116 kg Colwell P ha-1 and 18 kg Colwell P ha-1,

respectively. If biochar is to be applied in WA, the two most likely biomass sources

are wheat chaff or Eucalyptus wood. The Jarrah biochar contained 64 mg total P kg-1

and the wheat residue biochar contained 4150 mg total P kg-1 (data not shown). Jarrah

biochar application at 25 and 5 t ha-1 applied 1.6 and 0.3 kg total P ha-1 and wheat

residue biochar application at 4 t ha-1 applied 16.6 kg total P ha-1, although the

proportion of total P that is bio-available is unknown. In WA broadacre agro-

ecosystems application rates greater than 5 t ha-1 are unlikely to be applied, because

of the low bulk density of biochar and thus higher transport and application costs (as

stated in Chapter 5). It is unlikely that increases in N fertiliser use efficiency will


142

occur in WA agro-ecosystems because not enough biochar can be applied to supply P

and K in large enough quantities to induce such a response. Results may be different

if a nutrient rich manure biochar source becomes available or application rates of

biochar can be increased.

To increase biochar application rates, and hence P and K supply, in broadacre WA

agro-ecosystems the bulk density of biochar must be increased. The low bulk density

of biochar causes logistical problems and great expense. For example, a road train,

which can carry 60 t of wheat (bulk density of 0.8 g cm-3) will carry 15 t of biochar in

the same volume, assuming a bulk density of 0.2 g cm-3 (as in Chapter 5). Thus, at an

application rate of 4 t ha-1, this volume of biochar would be applied to only 3.75 ha.

Hence achieving the high application rates that would increase the supply of P and K,

in the broadacre agricultural environment of WA would be expensive and logistically

challenging.

6.5 Water supply and water holding capacity

Adding biochar to soil has been hypothesised to increase WHC and enhancing crop

water supply in sandy soils and dry environments (Atkinson et al., 2010; Blackwell et

al., 2010). Yet when calculated, the amount of water stored appears unlikely to

greatly enhance crop yield at potential application rates in broadacre agriculture in

WA (Chapter 5). Further, in the dry field environment of the WA broadacre agro-

ecosystem movement of inorganic N through the soil profile did not change with

biochar addition (Chapter 5), but in high rainfall environments, such as Amazonia


143

(Lehmann et al., 2003; Steiner et al., 2007) and glasshouse experiments (Chapter 3),

biochar has been shown to decrease inorganic N leaching. This is likely to be due to

the mass flow of water accentuating any changes in soil properties (e.g. refuges and

pores) created by biochar addition. Higher rainfall environments also allow

dissolution and diffusion of nutrients, such as P and K. Evidence of this may have

been provided at terminal spikelet sampling of the field experiment (Chapter 5),

where spread application of biochar significantly increased plant biomass; this

coincided with the wettest period of the growing season. Thus in drier environments

the impact of biochar may be sporadic and highly influenced by rainfall. This

explains the minimal of effect of biochar seen in studies assessing its agronomic

influence in south west Australia (Blackwell et al., 2010; Solaiman et al., 2010).

6.6 No effect of biochar

Previous glasshouse studies have suggested that biochar addition to soil can both

increase N fertiliser use efficiency and plant N uptake (Chan et al., 2007; Chan et al.,

2008; Van Zweiten et al., 2010a) and decrease plant N uptake and N fertiliser use

efficiency (Lehmann et al., 2003; Deenik et al., 2010). In high rainfall field

experiments plant N uptake can also increase (Steiner et al., 2007; Major et al.,

2010), but in a semi-arid environment with wheat residue biochar application at 4 t

ha-1 no influence was apparent (Chapter 5).

Field studies in WA and South Australia have demonstrated only sporadic increases

in shoot N uptake with biochar addition (Blackwell et al., 2010; Solaiman et al.,


144

2010). Shoot N uptake with Jarrah biochar addition at 1 t ha-1 increased in one of nine

treatments in one field experiment (Blackwell et al., 2010). In a second experiment,

35 year old Jarrah and Wandoo (Eucalyptus wandoo Blakely) biochar added at rates

up to 3.3 t ha-1 did not influence shoot N uptake (Blackwell et al., 2010). On a soil

type of greater clay content (sandy clay loam), also in south-western Australia, Oil

Mallee (Eucalyptus sp.) biochar application (6, 3, 1.5 or 0 t ha-1) combined with three

fertiliser application rates (30 or 55 kg ha-1 mono-ammonium phosphate, or 100 kg

ha-1 slow release composite fertiliser) only increased shoot N uptake in one treatment

(6 t biochar ha-1, 30 kg mono-ammonium phosphate ha-1; Solaiman et al., 2010).

Combined, the results of Blackwell et al. (2010), Solaiman et al. (2010) and Chapter

5 suggest that biochar is unlikely to increase plant N uptake in the drier areas of

broadacre agriculture within WA.

For agricultural research, there is a need for biochar based experiments to grow plants

to maturity to properly assess the impact of biochar on plant yield. Within WA and

South Australia, although some yield increases (Blackwell et al., 2010; Solaiman et

al., 2010) and yield decreases (Chapter 5) have been demonstrated, the majority of

the results from field experiments show no effect of biochar addition (Blackwell et

al., 2010; Solaiman et al., 2010; Yield results from year 2 of the field experiment in

Chapter 5 - data not shown). Thus biochar is unlikely to influence wheat yield in the

semi-arid broadacre field environment of WA.


145

6.7 Conclusion

This thesis tested the hypothesis that biochar addition would influence agronomically

important soil N cycling processes. Limited response in N leaching, N mineralisation,

nitrification, N immobilisation, and plant N uptake is likely to occur with

economically/agronomically viable application rates of biochar to coarse-textured

soils used for broadacre agriculture in WA. This is because sorption of LMW N

compounds to biochar added to soil is unlikely to influence N cycling possibly due to

the low application rates. Low application rates also ensure that the addition of P and

K in biochar is also unlikely to increase plant N uptake or nutrient use efficiency.

Where greater application rates can be applied, a high sorption affinity for

hydrophobic HMW organic matter may retard organic matter mineralisation,

decreasing nutrient, especially N, availability to crop plants. Findings reported within

this thesis suggest that biochar is unlikely to influence N cycling for the improvement

of crop productivity on coarse-textured soils in broadacre agriculture of WA.

146

References

147

References

ABARES. (2011) Agricultural commodity statistics 2011. Australian Bureau of

Agricultural and Resource Economics and Sciences, Commonwealth of

Australia.

Accardi-Dey, A., Gschwend, P. M. (2003) Reinterpreting literature sorption data

considering both absorption into organic carbon and adsorption on to black

carbon. Environmental Science & Technology 37, 99 – 106.

Aciego Pietri, J. C., Brookes, P. C. (2008) Relationships between soil pH and

microbial properties in a UK arable soil. Soil Biology & Biochemistry 40, 1856

– 1861.

Allred, A. L., Rochow, E. G. (1958) A scale of electronegativity based electrostatic

force. Journal of Inorganic & Nuclear Chemistry 5, 264 – 268.

Anderson, C. R., Condron, L. M., Clough, T. J., Fiers, M., Stewart, A., Hill, R. A.,

Sherlock, R. R. (2011) Biochar induced soil microbial community change:

Implications for biogeochemical cycling of carbon, nitrogen and phosphorus.

Pedobiologia 54, 309 – 320.

Anderson, G. C., Fillery, I. R. P., Dunin, F. X., Dolling, P. J., Asseng, S. (1998)

Nitrogen and water flows under pasture-wheat and lupin-wheat rotations in deep

sands in Western Australia 2. Drainage and nitrate leaching. Australian Journal

of Agricultural Research 49, 345 – 361.

Anderson J. P. E. (1982) Soil Respiration. 'Methods of Soil Analysis. Part 2.

Chemical and Microbiological Properties'. In: Page A. L., Miller R. H., Keeney

D. R. (Eds). Agronomy Monograph 9: ASA-SSSA, USA.

Anderson, J. P. E., Domsch, K. (1978) A physiological method for the quantitative

measurement of microbial biomass in soils. Soil Biology & Biochemistry 10,

215 – 221.

Anderson M. J. (2001) Permutation tests for univariate or multivariate analysis of

variance and regression. Canadian Journal of Fisheries and Aquatic Sciences

58, 626 – 639.

References

148

Anderson, W. K., Hoyle, F. C. (1999) Nitrogen efficiency of wheat cultivars in a

Mediterranean environment. Australian Journal of Experimental Agriculture

39, 957 – 965.

Angus, J. F. (2001) Nitrogen supply and demand in Australian agriculture. Australian

Journal of Experimental Agriculture 41, 277 – 288.

Antal, M. J., Grønli, M. (2003) The art, science and technology of charcoal

production. Industrial and Engineering Chemistry Research 42, 1619 – 1640.

ANZECC., ARMCANZ. (2000) Australian and New Zealand guidelines for fresh and

marine water quality: volume 1, the quidelines. Australian and New Zealand

Environment and Conservation Council and Agriculture and Resource

Management Council of Australia and New Zealand, Canberra.

Arbeles, F. B., Morgan, P. W., Saltveit, M. E. (1992) Ethylene in plant biology.

Academic press, London.

Atkinson, C. J., Fitzgerald, J. D., Hipps, N. A. (2010) Potential mechanisms for

achieving agricultural benefits from biochar application to temperate soils: a

review. Plant and Soil 337, 1 – 18.

Bailey, V. L., Fansler, S. J., Smith, J. L., Bolton, H. (2011) Reconciling apparent

variability in effects of biochar amendment on soil enzyme activities by assay

optimization. Soil Biology & Biochemistry 43, 296 – 301.

Baldock, J. A., Smernik, R. J. (2002) Chemical composition and bioavailability of

thermally altered Pinus resinosa (Red pine) wood. Organic Geochemistry 33,

1093 – 1109.

Ball, P. N., MacKenzie, M. D., DeLuca, T. H., Holben, W. E. (2010) Wildfire and

charcoal enhance nitrification and ammonium-oxidizing bacterial abundance in

dry montane forest soils. Journal of Environmental Quality 39, 1243 – 1253.

Bellamy, P. H., Loveland, P. J., Bradley, R. I., Lark, R. M., Kirk, G. J. D. (2005)

Carbon losses from all soils across England and Wales 1978-2003. Nature 437,

245 – 248.

Berglund, L. M., DeLuca, T. H., Zackrisson, O. (2004) Activated carbon amendments

to soil alter nitrification rates in Scots pine forests. Soil Biology & Biochemistry

36, 2067 – 2073.

References

149

Blackwell, P., Krull, E., Butler, G., Herbert, A., Solaiman, Z. (2010) Effect of banded

biochar on dryland wheat production and fertilizer use in south-western

Australia: an agronomic and economic perspective. Australian Journal of Soil

Research 48, 531 – 545.

Blackwood, C. B., Hudleston, D., Zak, D. R., Buyer, J. S. (2007) Interpreting

ecological diversity indices applied to terminal restriction fragment

polymorphism data: insights from simulated microbial communities. Applied

and Environmental Microbiology 73, 5276 – 5283.

Boddy, E., Hill., P. W., Farrar, J., Jones, D. L. (2007) Fast turnover of low molecular

weight components of the dissolved organic carbon pool of temperate grassland

field soils. Soil Biology & Biochemistry 39, 827 – 835.

Boehm, H. P. (1994) Some aspects of the surface chemistry of carbon blacks and

other carbons. Carbon 32, 759 – 769.

Bornemann, L. C., Kookana, R. S., Welp, G. (2007) Different sorption behaviour of

aromatic hydrocarbons on charcoals prepared at different temperatures from

grass and wood. Chemosphere 67, 1033 – 1042.

Bowman, G. M., Hutka, J. (2002) Particle size analysis, in: McKenzie, N. J.,

Coughlan, K. J., Cresswell, H. P. (Eds), Soil physical measurement and

interpretation for land evaluation. CSIRO publishing, Melbourne, Australia, pp.

224 – 239.

Bridle, T. R., Pritchard, D. (2004) Energy and nutrient recovery from sewerage

sludge via pyrolysis. Water Science and Technology 50, 169 – 175.

Brunauer, S., Emmett, P. H., Teller, E. (1938) Adsorption of gases in multimolecular

layers. Journal of the American Chemical Society 60, 309 – 319.

Bruun, S., Jensen, E. S., Jensen, L. S. (2008) Microbial mineralization and

assimilation of black carbon: Dependency on degree of thermal alteration.

Organic Geochemistry 39, 839 – 845.

Bureau of Meteorology. (2011) Climate statistics for Australian locations. Monthly

climate statistics. Available from:

<http://www.bom.gov.au/climate/averages/tables/cw_008039.shtml> Accessed

2nd August 2011.

References

150

Bureau of Meteorology. (2012) Decadal and multi-decadal rainfall. Available from:

<http://www.bom.gov.au/climate/averages/tables/cw_008039.shtml> Accessed

2nd March 2012.

Cabrera, M. L., Beare, M. H. (1993) Alkaline persulfate oxidation for determining

total nitrogen in microbial biomass extracts. Soil Science Society of America

Journal 57, 1007 – 1012.

Cao, X., Ma, L., Gao, B., Harris, W. (2009) Dairy-manure derived biochar effectively

sorbs lead and atrazine. Environmental Science & Technology 43, 3285 – 3291.

Chan, K. Y., Van Zwieten, L., Meszaros, I., Downie, A., Joseph, S. (2007)

Agronomic values of greenwaste biochar as a soil amendment. Australian

Journal of Soil Research 45, 629 – 634.

Chan, K. Y., Van Zwieten, L., Meszaros, I., Downie, A., Joseph, S. (2008) Using

poultry litter biochars as soil amendments. Australian Journal of Soil Research

46, 437 – 444.

Chen, D., Suter, H., Islam, A., Edis, R., Freney, J. R., Walker, C. N. (2008) Prospects

of improving efficiency of fertiliser nitrogen in Australian agriculture: a review

of enhanced efficiency fertilisers. Australian Journal of Soil Research 46, 289 –

301.

Chen, W., Bell, R. W., Brennan, R. F., Bowden, J. W., Dobermann, A., Rengel, Z.,

Porter, W. (2009a) Key crop nutrient management issues in the Western

Australian grains industry: a review. Australian Journal of Soil Research 47, 1

– 18.

Chen, H., Yao, J., Wang, F., Choi, M. M. F., Bramanti, E., Zaray, G. (2009b) Study

on the toxic effects of diphenol compounds on soil microbial activity by a

combination of methods. Journal of Hazardous Materials 167, 846 – 851.

Cheng, C., Lehmann, J., Engelhard, M. H. (2008) Natural oxidation of black carbon

in soils: Changes in molecular form and surface charge along a climosequence.

Geochimica et Cosmochimica Acta 72, 1598 – 1610.

Cheng, C., Lehmann, J., Thies, J. E., Burton, S. D., Engelhard, M. H. (2006)

Oxidation of black carbon by biotic and abiotic processes. Organic

Geochemistry 37, 1477 – 1488.

References

151

Chotte, J. L., Ladd, J. N., Amato, M. (1998) Sites of microbial assimilation, and

turnover of soluble and particulate 14C-labelled substrates decomposing in a

clay soil. Soil Biology & Biochemistry 30, 205 – 218.

Chun, Y., Sheng, G., Chiou, C. T., Xing, B. (2004) Compositions and sorptive

properties of crop residue-derived chars. Environmental Science & Technology

38, 4649 – 4655.

Clarholm, M. (1985) Interactions of bacteria, protozoa and plants leading to

mineralization of soil nitrogen. Soil Biology & Biochemistry 17, 181 – 187.

Clough, T. J., Bertram, J. E., Ray, J. L., Condron, L. M., O’Callaghan, M., Sherlock,

R. R., Wells, N. S. (2010) Unweathered wood biochar impact on nitrous oxide

emissions from a bovine-urine-amended pasture soil. Soil Science Society of

America Journal 74, 852 – 860.

Clough, T. J., Condron, L. M. (2010) Biochar and the nitrogen cycle: Introduction.

Journal of Environmental Quality 39, 1218 – 1223.

Cornelissen, G., Gustafsson, Ö. (2004) Sorption of phenanthrene to environmental

black carbon in sediment with and without organic matter and native sorbates.

Environmental Science & Technology 38, 148 – 155.

Cross, A., Sohi, S. P. (2010) The priming potential of biochar products in relation to

labile carbon contents and soil organic matter status. Soil Biology &

Biochemistry 43, 2127 – 2134.

Dalal, R. C., Chan, K. Y. (2001) Soil organic matter in rainfed cropping systems of

the Australian cereal belt. Australian Journal of Soil Research 39, 435 – 464.

Dalal, R. C., Mayer, R. J. (1986) Long-term trends in fertility of soils under

continuous cultivation and cereal cropping in southern Queensland. II Total

organic carbon and its rate of loss from the soil profile. Australian Journal of

Soil Research 24, 281 – 292.

De Boer, W., Kowalchuk, G. A. (2001) Nitrification in acid soil: micro-organisms

and mechanisms. Soil Biology & Biochemistry 33, 853 – 866.

Deenik, J. L., McClellan, T., Uehara, G., Antal, M. J., Campbell, S. (2010) Charcoal

volatile matter content influence plant growth and soil nitrogen transformations.

Soil Science Society of America Journal 74, 1259 – 1270.

References

152

Degens, B. P., Harris, J. A. (1997) Development of a physiological approach to

measuring the catabolic diversity of soil microbial communities. Soil Biology &


DeLuca, T. H., MacKenzie, M. D., Gundale, M. J., Holben, W. E. (2006) Wildfire-

produced charcoal directly influences nitrogen cycling in Ponderosa pine

forests. Soil Science Society of America Journal 70, 448 – 453.

Ding, Y., Liu, Y. X., Wu, W. X., Shi, D. Z., Yang, M., Zhong, Z. K. (2010)

Evaluation of biochar effects on nitrogen retention and leaching in multi-layered

soil columns. Water, Air, and Soil Pollution 213, 47 – 55.

Dolling, P. J., Porter, W. M. (1994) Acidification rates in the central wheatbelt of

Western Australia. 1. On a deep yellow sand. Australian Journal of

Experimental Agriculture 34, 1155 – 1164.

Dolling, P. J., Porter, W. M., Rowland, I. C. (1994) Acidification rates in the central

wheatbelt of Western Australia. 2. On a sandy duplex soil. Australian Journal

of Experimental Agriculture 34, 1165 – 1172.

Downie, A., Crosky, A., Munroe, P. (2009) Physical properties of biochar. In

‘Biochar for environmental management, science and technology’. (Eds J.

Lehmann, S. Joseph) pp. 13-32. (Earthscan, London, UK)

Dunbar, J., Ticknor, L. O., Kuske, C. R. (2001) Phylogenetic specificity and

reproducibility and new method for analysis of terminal restriction fragment

profiles of 16S rRNA genes from bacterial communities. Applied and

Environmental Microbiology 67, 190 – 197.

Durenkamp, M., Luo, Y., Brookes, P.C. (2010) Impact of black carbon addition to

soil on the determination of soil microbial biomass by fumigation extraction.

Soil Biology & Biochemistry 42, 2026 – 2029.

FAO. (1988) FAO/UNESCO Soil map of the World, revised legend, with corrections

and updates. World soil resources report 60, FAO, Rome.

Flavel, T. C., Murphy, D. V. (2006) Carbon and nitrogen mineralization rates after

application of organic amendments to soil. Journal of Environmental Quality

35, 183 – 193.

References

153

French, R. J., Shultz, J. E. (1984) Water use efficiency of wheat in a Mediterranean

type environment. 1 The relationship between yield, water use and climate.

Australian Journal of Agricultural Research 35, 743 – 764.

Gazey, C., Andrew, J. (2009) Soil pH in northern and southern areas of the WA

wheatbelt. Bulletin 4761. Department of Agriculture and Food, Western

Australia.

Gentile, R., Vanlauwe, B., van Kessel, C., Six, J. (2009) Managing N availability and

losses by combining fertilizer-N with different quality residues in Kenya.

Agriculture, Ecosystems and Environment 131, 308 – 314.

Girvan, M. S., Campbell, C. D., Killham, K., Prosser, J. I., Glover, L. A. (2005)

Bacterial diversity promotes community stability and functional resilience after

perturbation. Environmental Microbiology 7, 301 – 313.

Glaser, B., Lehmann, J., Zech, W. (2002) Amerliorating physical and chemical

properties of highly weathered soils in the tropics with charcoal – a review.

Biology and Fertility of Soils 35, 219 – 230.

Gleeson, D. B., Müller, C., Banerjee, S., Ma, W., Siciliano, S. D., Murphy, D. V.

(2010) Response of ammonia oxidizing archaea and bacteria to changing water

filled pore space. Soil Biology & Biochemistry 42, 1888 – 1891.

Graber, E. R., Harel, Y. M., Kolton, M., Cytryn, E., Silber, A., David, D. R.,

Tsechansky, L., Borenshtein, M., Elad, Y. (2010) Biochar impact on

development and productivity of pepper and tomato grown in fertigated soilless

media. Plant and Soil 337, 481 – 496.

Griffiths, R. I., Whiteley, A. S., O’Donnell, A. G., Bailey, M. J. (2000) Rapid method

for coextraction of DNA and RNA from natural environments for analysis of

ribosomal DNA- and RNA-based microbial community composition. Applied

and Environmental Microbiology 66, 5488 – 5491.

Gundale, M. J., DeLuca, T. H. (2007) Charcoal effects on soil solution chemistry and

growth of Koeleria macrantha in the ponderosa pine/Douglas fir ecosystem.

Biology and Fertility of Soils 43, 303 – 311.

Hall, D. J. M., Jones, H. R., Crabtree, W. L., Daniels, T. L. (2010) Claying and deep

ripping can increase crop yields and profits on water repellent sands with

References

154

marginal fertility in southern Western Australia. Australian Journal of Soil

Research 48, 178 – 187.

Harper, R. J., Gilkes, R. J. (2004) The effects of clay and sand additions on the

strength of sandy topsoils. Australian Journal of Soil Research 42, 39 – 44.

Harper, R. J., McKissock, I., Gilkes, R. J., Carter, D. J., Blackwell, P. S. (2000) A

multivariate framework for interpreting the effects of soil properties, soil

management and landuse on water repellency. Journal of Hydrology 231 – 232,

371 – 383.

Hart, R. D., Gilkes, R. J., Siradz, S., Singh, B. (2002) The nature of soil kaolins from

Indonesia and Western Australia. Clays and Clay Minerals 50, 198 – 207.

Hill P. W. Farrell, M., Roberts, P., Farrar, J., Grant, H., Newsham, K. K., Hopkins, D.

W., Bardgett, R. D., Jones, D. L. (2011) Soil- and enantiomer-specific

metabolism of amino acids and their peptides by Antartic soil microorganisms.


Hina, K., Bishop, P., Camps Arbestain, M., Calvelo-Pereira, R., Maciá-Agulló,

Hindmarsh, J., Hanly, J. A., Macías, F., Hedley, M. J. (2010) Producing

biochars with enhanced surface activity through alkaline pretreatment of

feedstocks. Australian Journal of Soil Research 48, 606 – 617.

Holmes, K. W., Wherrett, A., Keating, A., Murphy, D. V. (2011) Meeting bulk

density sampling requirements efficiently to estimate soil carbon stocks. Soil

Research 49, 680 – 695.

Hoyle, F. C., Baldock, J. A., Murphy, D. V. (2011) Soil organic carbon. Role in

rainfed farming systems. With particular reference to Australian conditions. In

‘Rainfed farming systems’. (Eds P Tow, I Partridge, I Cooper, C Birch) pp. 339

– 364. (Springer Science, London)

Hoyle, F. C., Murphy, D. V. (2011) Influence of organic residues and soil

incorporation on temporal measures of microbial biomass and plant available

nitrogen. Plant and Soil 347, 53 – 64.

Horz H. P., Barbrook A., Field C. B., Bohannan B. J. M. (2004) Ammonia-oxidizing

bacteria respond to multifactorial global change. Proceedings of the National

Academy of Science of the United States of America 101, 15136 – 15141.

References

155

Isbell, R.F. (1996) The Australian Soil Classification. CSIRO Publishing, Melbourne,

Australia.

Jackson, R. B., Schlesinger, W. H. (2004) Curbing the US carbon deficit.

Proceedings of the National Academy of the United States of America 101,

15827 – 15829.

Jackson, R. M., Sternberg, M. J. E. (1994) Application of scaled particle theory to

model the hydrophobic effect: implications for molecular association and

protein stability. Protein Engineering 7, 371 – 383.

Jeffery, S., Verheijen, F. G. A., Van Der Velde, M., Bastos, A. C. (2011) A

quantitative review of the effects of biochar application to soils on crop

productivity using meta-analysis. Agriculture, Ecosystems & Environment 144,

175 – 187.

Jenkinson, D. S. (2001) The impact of humans on the nitrogen cycle, with focus on

temperate arable agriculture. Plant and Soil 228, 3 – 15.

Joergensen, R. G., Brookes, P. C. (1990) Ninhydrin-reactive nitrogen measurements

of microbial biomass in 0.5 M K2SO4 soil extracts. Soil Biology & Biochemistry

22, 1023 – 1027.

Jones, D. L., Edwards-Jones, G., Murphy, D. V. (2010) Biochar mediated alterations

in herbicide breakdown and leaching in soil. Soil Biology & Biochemistry 43,

804 – 813.

Jones, D. L., Healey, J. R., Willett, V. B., Farrar, J. F., Hodge, A. (2005) Dissolved

organic nitrogen uptake by plants - an important N uptake pathway? Soil

Biology & Biochemistry 37, 413 – 423.

Jones, D. L., Murphy, D. V. (2007) Microbial response time to sugar and amino acid

additions to soil. Soil Biology & Biochemistry 39, 2178 – 2182.

Jones, D. L., Murphy, D.V., Khalid, M., Ahmad, W., Edwards-Jones, G., DeLuca, T.

H. (2011) Short-term biochar-induced increase in soil CO2 release is both

biotically and abiotically mediated. Soil Biology & Biochemistry 43, 1723 –

1731.

References

156

Jones, D. L., Owen, A. G., Farrar, J. F. (2002) Simple method to enable the high

resolution of total free amino acids in soil solutions and soil extracts. Soil


Jones, D. L., Shannon, D., Murphy, D. V., Farrar, J. (2004) Role of dissolved organic

nitrogen (DON) in soil N cycling in grassland soils. Soil Biology &


Kaiser, K., Guggenberger, G. (2000) The role of DOM sorption to mineral surfaces in

the preservation of organic matter in soils. Organic Geochemistry 31, 711 –

725.

Kamphake, L.J., Hannah, S.A., Cohen, J.M. (1967) Automated analysis for nitrate by

hydrazine reduction. Water Research 1, 205 – 216.

Kasozi, G. N., Zimmerman, A. R., Nkedi-Kizza, P., Goa, B. (2010) Catechol and

humic acid sorption onto a range of laboratory-produced black carbons

(biochars). Environmental Science & Technology 44, 6189 – 6195.

Kastner, J. R., Miller, J., Das, K. C., (2009) Pyrolysis conditions and ozone oxidation

effects on ammonia adsorption in biomass generated chars. Journal of

Hazardous Materials 164, 1420 – 1427.

Keating, B. A., Carberry, P. S. (2010) Emerging opportunities and challenges for

Australian broadacre agriculture. Crop & Pasture Science 61, 269 – 278.

Keiluweit, M., Kleber, M. (2009) Molecular-level interactions in soils and sediments:

The role of aromatic π-systems. Environmental Science & Technology 43, 3421

– 3429.

Keith, A., Singh, B., Singh, B. P. (2011) Interactive priming of biochar and labile

organic matter mineralization in a smectite-rich soil. Environmental Science &

Technology 45, 9611 – 9618.

Kempers, A.J., Luft, A.G. (1988) Re-examination of the determination of

environmental nitrate as nitrite by reduction with hydrazine. The Analyst 113,

1117 – 1120.

Khodadad, C. L. M., Zimmerman, A. R., Green, S. J., Uthandi, S., Foster, J. S. (2011)

Taxa-specific changes in soil microbial community composition induced by

pyrogenic carbon amendments. Soil Biology & Biochemistry 43, 385 – 392.

References

157

Killham, K. (2006) Soil Ecology. Cambridge University Press, Cambridge.

Kingwell, R. (2011) Managing complexity in modern farming. The Australian

Journal of Agricultural and Resource Economics 55, 12 – 34.

Kingwell, R., Pannell, D. (2005) Economic trends and drivers affecting the wheatbelt

of Western Australia to 2030. Australian Journal of Agricultural Research 56,

553 – 561.

Kleber, M., Johnson, M. G. (2010) Advances in understanding the molecular

structure of soil organic matter: implications for interactions in the environment.

Advances in Agronomy 106, 77 – 142.

Kolb, S. E., Fermanich, K. J., Dornbush, M. E. (2009) Effect of charcoal quantity on

microbial biomass and activity in temperate soils. Soil Science Society of


Krom, M. D., (1980) Spectrophotometric determination of ammonia; a study of a

modified Berthelot reaction using salicylate and dichloroisocyanurate. The

Analyst 105, 305 – 316.

Kuzyakov, Y., Subbotina, I., Chen, H.Q., Bogomolova, I., Xu, X.L. (2009) Black

Carbon decomposition and incorporation into the microbial biomass estimated

by 14C labeling. Soil Biology & Biochemistry 41, 210 – 219.

Ladha, J. K., Pathak, H., Krupnik, T. J., Six, J., Van Kessel, C. (2005) Efficiency of

fertilizer nitrogen in cereal production: retrospects and prospects. Advances in

Agronomy 87, 85 – 156.

Laird, D. A. (2008) The charcoal vision: A win-win-win scenario for simultaneously

producing bioenergy, permanently sequestering carbon, while improving soil

and water quality. Agronomy Journal 100, 178 – 181.

Laird, D., Fleming, P., Davis, D. D., Horton, R., Wang, B., Karlen, D. (2010a) Impact

of biochar amendments on the quality of a typical Midwestern agricultural soil.

Geoderma 158, 443 – 449.

Laird, D., Fleming, P., Wang, B., Horton, R., Karlen, D. (2010b) Biochar impact on

leaching from a Midwestern agricultural soil. Geoderma 158, 436 – 442.

References

158

Lalor, B. M., Cookson, W. R., Murphy, D.V. (2007) Comparison of two methods that

assess soil community level physioligical profiles in a forest ecosystem. Soil


Lammirato, C., Miltner, A., Kaestner, M. (2011) Effects of wood char and activated

carbon on the hydrolysis of cellobiose by β-glucosidase from Aspergillus niger.


Lawes Agricultural Trust. 2007. Genstat 10th Edition. VSN International Ltd, Hemel

Hempstead, U.K.

Lehmann, J. (2007) Bioenergy in the black. Frontiers in Ecology and the

Environment 5, 381 – 387.

Lehmann, J., da Silva., J. P., Steiner, C., Nehls, T., Zech, W., Glaser, B. (2003)

Nutrient availability and leaching in an archaeological Anthrosol and a Ferralsol

of the Central Amazon basin: fertilizer, manure and charcoal amendments.

Plant and Soil 249, 343 – 357.

Lehmann, J., Gaunt, J., Rondon, M. (2006) Bio-char sequestration in terrestrial

ecosystems – a review. Mitigation and Adaptation Strategies for Global Change

11, 395 – 419.

Lehmann, J., Joseph, S. (2009) Biochar for environmental management: An

introduction. In ‘Biochar for environmental management, science and

technology’. (Eds J Lehmann, S Joseph) pp. 1-12. (Earthscan, London, UK)

Lehmann, J., Rillig, M., Thies, J., Masiello, C. A., Hockaday, W. C., Crowley, D.

(2011) Biochar effects on soil biota - a review. Soil Biology & Biochemistry 43,

1812 – 1836.

Liang, B., Lehmann, J., Sohi, S. P., Thies, J. E., O’Neill, B., Trujillo, L., Gaunt, J.,

Solomon, D., Grossman, J., Neves, E. G., Luizão, F. J. (2010) Black carbon

affects the cycling of non-black carbon in soil. Organic Geochemistry 41, 206 –

213.

Liang, B., Lehmann, J., Solomon, D., Kinyangi, J., Grossman, J., O’Neill, B.,

Skjemstad, J. O., Thies, J., Luizão, F. J., Peterson, J., Neves, E. G. (2006) Black

carbon increases cation exchange capacity in soils. Soil Science Society of


References

159

Ma, Y., Rate, A. W. (2007) Metals adsorbed to charcoal are not identifiable by

sequential extraction. Environmental Chemistry 4, 26 – 34.

Major, J., Rondon, M., Molina, D., Riha, S. J., Lehmann, J. (2010) Maize yield and

nutrition during 4 years after biochar application to a Colombian savanna

oxisol. Plant and Soil 333, 117 – 128.

Manzoni, S., Jackson, R. B., Trofymow, J. A., Porporato, A. (2008) The global

stoichiometry of litter nitrogen mineralisation. Science 321, 684 – 686.

Mason, M. G., Rowley, A. M., Quayle, D. J. (1972) The fate of urea applied at

various intervals after the sowing of a wheat crop on sandy soil in Western

Australia. Australian Journal of Experimental Agriculture and Animal

Husbandry 12, 171 – 175.

McDonald, G. K. (1989) The contribution of nitrogen fertiliser to the nitrogen

nutrition of rainfed wheat crops in Australia: a review. Australian Journal of

Experimental Agriculture 29, 455 – 481.

McKissock, I., Gilkes, R. J., Walker, E. L. (2002) The reduction of water repellency

by added clay is influenced by clay and soil properties. Applied Clay Science

20, 225 – 241.

McKissock, I., Walker, E. L., Gilkes, R. J., Carter, D. J. (2000) The influence of clay

type on reduction of water repellency by applied clays: a review of some West

Australian work. Journal of Hydrology 231-232, 323 – 332.

McNeill, A. M., Sparling, G. P., Murphy, D. V., Braunberger, P., Fillery, I. R. P.

(1998) Changes in extractable and microbial C, N, and P in a Western

Australian wheatbelt soil following simulated summer rainfall. Australian

Journal of Soil Research, 36, 841 – 854.

Miranda, K. M., Epsey, M. G., Wink, D. A. (2001) A rapid, simple

spectrophotometric method for simultaneous detection of nitrate and nitrite.

Nitric Oxide: Biology and Chemistry 5, 62 – 71.

Mizuta, K., Matsumoto, T., Hatate, Y., Nishihara, K., Nakanishi, T. (2004) Removal

of nitrate-nitrogen from drinking water using bamboo powder charcoal.

Bioresource Technology 95, 255 – 257.

References

160

Moore, G. (2004) ‘Soil guide - A handbook for understanding and managing

agricultural soils’. National Landcare and Department of Agriculture Western

Australia Bulletin No. 4343 (compiled and edited), Perth, Australia

Mukherjee, A., Zimmerman, A. R., Harris, W. (2011) Surface chemistry variations

among a series of laboratory-produced biochars. Geoderma 163, 247 – 255.

Mulvaney, R. L. (1996) Nitrogen - inorganic forms, in: Sparks, D. L. (Ed), Methods

of soil analysis, Part 3, Chemical Methods. SSSA, Madison, USA, pp. 1123 –

1184.

Murphy, D. V., Cookson, W. R., Braimbridge, M., Marschner, P., Jones, D. L.,

Stockdale, E. A., Abbott, L. K. (2011) Relationships between soil organic

matter and the soil microbial biomass (size, functional diversity, and

community structure) in crop and pasture systems in a semi-arid environment.

Soil Research 49, 582 – 594.

Murphy, D. V., MacDonald, A. J., Stockdale, E. A., Goulding, K. W. T., Fortune, S.,

Gaunt, J. L., Poulton, P. R., Wakefield, J. A., Webster, C. P., Wilmer, W. S.

(2000) Soluble organic nitrogen in agricultural soils. Biology and Fertility of

Soils 30, 374 – 387.

Murphy, D.V., Recous, S., Stockdale, E. A., Fillery, I. R. P., Jensen, L. S., Hatch, D.

J., Goulding, K. W. T. (2003) Gross nitrogen fluxes in soil: theory,

measurement and application of 15N pool dilution techniques. Advances in

Agronomy 79, 69 – 119.

Muñoz-Leoz, B., Ruiz-Romera, E., Antigüedad, I., Garbisu, C. (2011) Tebuconazole

application decreases soil microbial biomass and activity. Soil Biology &


Nag, S. K., Kookana, R., Smith, L., Krull, E., Macdonald, L. M., Gill, G. (2011) Poor

efficacy of herbicides in biochar-amended soils as affected by their chemistru

and mode of action. Chemosphere 84, 1572 – 1577.

Ogawa, M., Okimori, Y., Takahashi, F. (2006) Carbon sequestration by carbonisation

of biomass and forestation: three case studies. Mitigation and Adaptation

Strategies for Global Change 11, 421 – 436.

References

161

Paavolainen, L., Kitunen, V., Smolander, A. (1998) Inhibition of nitrification in

forest soils by monoterpenes. Plant and Soil 205, 147 – 154.

Pietikäinen, J., Kiikkilä, O., Fritze, H. (2000) Charcoal as a habitat for microbes and

its effect on the microbial community of the underlying humus. Oikos 89, 231 –

242.

Pikaar, I., Koelmans, A. A., van Noort, P. C. M. (2006) Sorption of organic

compounds to activated carbons. Evaluation of isotherm models. Chemosphere

65, 2343 – 2351.

Pitman, A. J., Narisma, G. T., Pielke, R. A., Holbrook, N. J. (2004) Impact of land

cover change on the climate of southwest Western Australia. Journal of

Geophysical Research - Atmospheres 109, D18109, doi:

10.1029/2003JD004347.

Plymouth Marine Laboratory. 2007. Primer 6 and Permanova+β18. Primer E Ltd,

Roborough, Plymouth, UK.

Porter, L. K. (1992) Ethylene inhibition of ammonium oxidation in soil. Soil Science

Society of America Journal 56, 102 – 105.

Power, J. F., Wiese, R.., Flowerday, D. (2001) Managing farming systems for nitrate

control: a research review from management systems evacuation areas. Journal

of Environmental Quality 30, 1866 – 1880.

Prado, A. G. S., Airoldi, C. (2001) The effect of the herbicide diuron on soil

microbial activity. Pest Management Science 57, 640 – 644.

Prendergast-Miller, M. T., Duvall, M., Sohi, S. P. (2011) Localisation of nitrate in the

rhizosphere of biochar-amended soils. Soil Biology & Biochemistry 43, 2243 –

2246.

Preston, C. M., Schmidt, M. W. I. (2006) Black (pyrogenic) carbon: a synthesis of

current knowledge and uncertainties with special consideration of boreal

regions. Biogeosciences 3, 397 – 420.

Quiroga-Garza, H. M., Picchioni, G. A., Remmenga, M. D. (2001) Bermudagrass

fertilized with slow-release nitrogen sources. 1. Nitrogen uptake and potential

leaching losses. Journal of Environmental Quality 30, 440 – 448.

References

162

Raun, W. R., Johnson, G. V. (1999) Improving nitrogen use efficiency for cereal

production. Agronomy Journal, 91, 357 – 363.

Rayment, G. E., Higginson, F. R. (1992) Australian laboratory handbook of soil and

water chemical methods. Inkata Press, Melbourne, Victoria, Australia.

Rayment, G. E., Lyons, D. J. (2011) Alkaline earth carbonates. In ‘Soil chemical

methods - Australasia’. (Eds G. E. Rayment, Lyons, D. J.) pp 415 - 425.

(CSIRO Publishing: Melbourne)

R Core Development Team (2007) R: A language and environment for statistical

computing. R Foundation for Statistical Computing, Vienna

Ridley, A. M., White, R. E., Helyar, K. R., Morrison, G. R., Heng, L. K., Fisher, R.

(2001) Nitrate leaching loss under annual and perennial pastures with and

without lime on a duplex (texture contrast) soil in humid southeastern Australia.

European Journal of Soil Science 52, 237 – 252.

Rosswall, T. (1982) Microbial regulation of the biogeochemical nitrogen cycle. Plant

and Soil 67, 15 – 34.

Rotthauwe, J. H., Witzel, K. P., Liesack, W. (1997) The ammonia monooxygenase

structural gene amoA as a functional marker: molecular fine scale analysis of

natural ammonia oxidising populations. Applied and Environmental

Microbiology 63, 4704 – 4712.

Rovira, A. D. (1992) Dryland Mediterranean farming systems in Australia. Australian

Journal of Experimental Agriculture 32, 801 – 809.

Saggar, S., Parshotam, A., Hedley, C., Salt, G. (1999) 14C-labelled glucose turnover

in New Zealand soils. Soil Biology & Biochemistry 31, 2025 – 2037.

Scallan, Ú., Liliensiek, A., Clipson, N., Connolly, J., 2008. RiboSort: a program for

automated data preparation and exploratory analysis of microbial community

fingerprints. Molecular Ecology Notes 8, 95 – 98.

Schmidt, M. W. I., Noack, A. G. (2000) Black carbon in soils and sediments:

Analysis, distribution, implications, and current challenges. Global

Biogeochemical Cycles 14, 777 – 793.

References

163

Schoknecht, N. (2002) Soil groups of Western Australia: A simple guide to the main

soils of Western Australia. Resource Management Technical Report 246.

Department of Agriculture, Western Australia.

Searle, P.L. (1984) The Berthelot or indophenol reaction and its use in the analytical

chemistry of nitrogen – a review. The Analyst 109, 549 – 568.

Seredych, M., Bandosz, T. J. (2007) Mechanism of ammonia retention on graphite

oxides: Role of surface chemistry and structure. Journal of Physical Chemistry

111, 15596 – 15604.

Singh, B., Singh, B. P., Cowie, A. L. (2010) Characterisation and evaluation of

biochars for their application as a soil amendment. Australian Journal of Soil

Research 48, 516 – 525.

Smith, J. L., Collins, H. P., Bailey, V. L. (2010) The effect of young biochar on soil

respiration. Soil Biology & Biochemistry 42, 2345 – 2347.

Smolders, A. J. P., Lucassen, E. C. H. E. T., Bobbink, R., Poelofs, J. G. M., Lamers,

L. P. M. (2010) How nitrate leaching from agricultural lands provokes

phosphate eutrophication in groundwater fed wetlands: the sulphur bridge.

Biogeochemistry 98, 1 – 7.

Sohi, S. P., Krull, E., Lopez-Capel, E., Bol, R. (2010) A review of biochar and its use

and function in soil. Advances in Agronomy 105, 47 – 82.

Solaiman, Z. M., Blackwell, P., Abbott, L. K., Storer, P. (2010) Direct and residual

effect of biochar application on mycorrhizal root colonisation, growth and

nutrition of wheat. Australian Journal of Soil Research 48, 546 – 554.

Sparling, G. P., Gupta, V. V. S. R., Zhu, C. (1993) Release of ninhydrin-reactive

compounds during fumigation of soil to estimate microbial C and N. Soil


Spokas, K. A., Baker, J. M., Reicosky, D. C. (2010) Ethylene: potential key for

biochar amendment impacts. Plant and Soil 333, 443 – 452.

Spokas, K. A., Novak, J. M., Ventera, R. T. (2012) Biochar’s role as an alternative N-

fertilizer: ammonia capture. Plant and Soil 350, 35 – 42.

References

164

Steinbeiss, S., Gleixner, G., Antonietti, M. (2009) Effect of biochar amendment on

soil carbon balance and soil microbial activity. Soil Biology & Biochemistry 41,

1301 – 1310.

Steiner, C., Das, K. C., Garcia, M., Förster, B., Zech, W. (2008a) Charcoal and

smoke extract stimulate the soil microbial community in a highly weathered

xanthic Ferralsol. Pedobiologia 51, 359 – 366.

Steiner, C., Glaser, B., Teixeira, W. G., Lehmann, J., Blum, W. E. H., Zech, W.

(2008b) Nitrogen retention and plant uptake on a highly weathered central

Amazonian Ferralsol amended with compost and charcoal. Journal of Plant

Nutrition and Soil Science 171, 893 – 899.

Steiner, C., Teixeira, W. G., Lehmann, J., Nehls, T., Vasconcelos de Macêdo, J. L.,

Blum, W. E. H. (2007) Long term effects of manure, charcoal and mineral

fertilization of crop production and fertiliser on a highly weathered Central

Amazonian upland soil. Plant and Soil 291, 275 – 290.

Stevenson, B.A., Sparling, G. P., Schipper, L. A., Degens, B. P., Duncan, L. C.

(2004) Pasture and forest soil microbial communities show distinct patterns in

their catabolic respiration responses at a landscape scale. Soil Biology &


Strong, D. T., Sale, P. W. G., Helyar, K. R. (1998) The influence of the soil matrix on

nitrogen mineralisation and nitrification. II. The pore system as a framework for

mapping the organisation of the soil matrix. Australian Journal of Soil Research

36, 855 – 872.

Sverdrup, L. E. Ekelund, F., Krogh, P. H., Nielsen, T., Johnsen, K. (2002) Soil

microbial toxicity of eight polycyclic aromatic hydrocarbons: effects on

nitrification, the genetic diversity of bacteria, and the total number of

protozoans. Environmental Toxicology and Chemistry 21, 1644 – 1650.

Taghizadeh-Toosi, A., Clough, T. J., Sherlock, R. R., Condron, L. M. (2012) Biochar

adsorbed ammonia is bioavailable. Plant and Soil 350, 57 – 69.

Tennant, D., Scholz, G., Dixon, J., Purdie, B. (1992) Physical and chemical

characteristics of duplex soils and their distribution in the south-west of

References

165

Western Australia. Australian Journal of Experimental Agriculture 32, 827 –

843.

Tryon E. H. (1948) Effects of charcoal on certain physical, chemical and biological

properties of forest soils. Ecological Monographs 18, 81 – 115.

Uusitalo, M., Kitunen, V., Smolander, A. (2008) Response of C and N

transformations in birch soil to coniferous resin volatiles. Soil Biology &


Vance, E. D., Brookes, P. C., Jenkinson, D. S. (1987) An extraction method for

measuring soil microbial biomass C. Soil Biology & Biochemistry 19, 703 –

707.

Van Hees, P. A. W., Jones, D. L., Finlay, R., Godbold, D. L., Lundström, U. S.

(2005) The carbon we do not see - the impact of low molecular weight

compounds on carbon dynamics and respiration in forest soil: a review. Soil


Van Zwieten, L., Kimber, S., Downie, A., Morris, S., Petty, S., Rust, J., Chan, K. Y.

(2010a) A glasshouse study on the interaction of low mineral ash biochar with

nitrogen in a sandy soil. Australian Journal of Soil Research 48, 569 – 576.

Van Zwieten, L., Kimber, S., Morris, S., Chan, K. Y., Downie, A., Rust, J., Joseph,

S., Cowie, A. (2010b) Efects of biochar from slow pyrolysis of papermill waste

on agronomic performance and soil fertility. Plant and Soil 32, 327 – 246.

Van Zwieten, L., Kimber, S., Morris, S., Downie, A., Berger, E., Rust, J., Scheer, C.

(2010c) Influence of biochars on flux of N2O and CO2 from Ferrosol.

Australian Journal of Soil Research 48, 555 – 568.

Van Zwieten, L., Singh, B. P., Joseph, S., Kimber, S., Cowie, A., Chan, K. Y. (2009)

Biochar and emissions of non-CO2 greenhouse gases from soil. In ‘Biochar for

environmental management, science and technology’. (Eds J. Lehmann, S.

Joseph) pp.227 – 249. (Earthscan, London, UK)

Vlahos, S., Summers, K. J., Bell, D. T., Gilkes, R. J. (1989) Reducing phosphorus

leaching from sandy soils with red mud bauxite processing residues. Australian

Journal of Soil Research 27, 651 – 662.

References

166

Wardle, D. A., Nilsson, M–C., Zackrisson, O. (2008) Fire-derived charcoal causes

loss of forest humus. Science 320, 629.

Wardle, D. A., Zackrisson, O., Nilsson, M–C. (1998) The charcoal effect in Boreal

forests: mechanisms and ecological consequences. Oecologia 115, 419 – 426.

White, C. S. (1988) Nitrification inhibition by monoterpenoids: Theoretical mode of

action based on molecular structures. Ecology 69, 1631 – 1633.

White, C. S. (1991) The role of monoterpenes in soil nitrogen cycling processes in

panderosa pine. Biogeochemistry 12, 43 – 68.

White, C. S. (1994) Monoterpenes: Their effects on ecosystem nutrient cycling.

Journal of Chemical Ecology 20, 1381 – 1406.

Wu, J., Joergensen, R. G., Pommerening, B., Chaussod, R., Brookes, P. C. (1990)

Measurement of soil microbial biomass C by fumigation-extraction — an

automated procedure. Soil Biology & Biochemistry 22, 1167 – 1169.

Yang, Y., Chun, Y., Sheng, G., Huang, M. (2004) pH-dependence of pesticide

adsorption by wheat-residue-derived black carbon. Langmuir 20, 6736 – 6741.

Yu, X. Y., Ying, G. G., Kookana, R. S. (2006) Sorption and desorpton behaviours of

diuron in soils amended with charcoal. Journal of Agricultural and Food

Chemistry 54, 8545 – 8550.

Zackrisson, O., Nilsson, M. C., Wardle D. A. (1996) Key ecological function of

charcoal from wildfire in the Boreal forest. Oikos 77, 10 – 19.

Zhou, Z., Shi, D., Qiu, Y., Sheng, G. D. (2010) Sorptive domains of pine shars as

probed by benzene and nitrobenzene. Environmental Pollution 158, 201 – 206.

Zhu, D., Pignatello, J. J., (2005) Characterisation of aromatic compound sorptive

interactions with black carbon (charcoal) assisted by graphite as a model.

Environmental Science & Technology 39, 2033 – 2041.

Zimmerman, A. R. (2010) Abiotic and microbial oxidation of laboratory-produced

black carbon (biochar). Environmental Science & Technology 44, 1295 – 1301.

Zimmerman, A. R., Gao, B., Ahn, M. (2011) Positive and negative carbon

mineralization priming effects among a variety of biochar-amended soils. Soil


References

167

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Biochar and the Soil Nitrogen Cycle: Unravelling the ... › files › ...Biochar and the Soil Nitrogen Cycle: Unravelling the Interactions Daniel Norman Dempster BSc (Agric, Hons);