the role of temperature, nutrient availability and organo ... · decomposition while organo-mineral...

The Role of Temperature, Nutrient Availability and Organo-

Mineral Interactions in Altering Soil Organic Matter

Composition

by

Olivia Oi Ying Lun

A thesis submitted in conformity with the requirements

for the degree of Master of Science

Graduate Department of Chemistry

University of Toronto

© Copyright by Olivia Oi Ying Lun 2016

ii

The Role of Temperature, Nutrient Availability and Organo-Mineral

Interactions in Altering Soil Organic Matter Composition

Olivia Oi Ying Lun

Master of Science

Graduate Department of Chemistry

University of Toronto

2016

Abstract

Increases in temperature and nutrient availability may accelerate soil organic matter (SOM)

decomposition while organo-mineral interactions may potentially stabilize SOM components.

After 32 years of warming, nitrogen and phosphorus (N+P) fertilization, and the combined

treatment, SOM degradation was not significantly enhanced in Arctic soils. Considerable

variability in the biomarker data may be attributable to spatial heterogeneity among the blocks.

In the second project, biomarker investigations indicated that mineral interactions likely protect

lignin from extraction. Suberin and cutin protection of lignin was not observed until after clay

mineral dissolution by hydrofluoric acid (HF). This suggests that clay mineral interactions may

play a more dominant role in the protection of lignin than interactions with suberin and cutin.

Overall, this thesis demonstrates that biomarker methods can show how the distribution of SOM

may be reflective of topographical patterns and can provide further evidence that mineral

protection mechanisms help stabilize SOM components.

iii

Acknowledgments

Firstly, I would like to express my deepest gratitude to Prof. Myrna Simpson for her

guidance and support, particularly during the compilation process of this thesis. Her

encouragement along with her patience and understanding are greatly appreciated. Prof. André

Simpson and Prof. Jennifer Murphy are thanked for agreeing to serve on my supervisory

committee. Dr. Megan Machmuller and Dr. Eldor Paul, our collaborators from Colorado State

University, are thanked for collecting samples from Toolik Lake, Alaska.

I would also like to thank all the past and present members of the M. Simpson and A.

Simpson Research Groups. In particular, I would like to thank Dr. Oliva Pisani and Perry

Mitchell for helpful discussions on organic geochemistry. Thank you for guiding me through the

extraction methods and data analysis procedures. Thank you to Kalyani Sabanayagam for always

keeping a smile. More importantly, thank you for updating the blackboard outside our lab with

your inspirational quotes and creative drawings. They never fail to make my day. Thank you to

Lisa Lin for your mentorship. Thank you to Lori vandenEnden for all your laughs and smiles.

Thank you to Justin Wang and Duke (Zhangliu) Du for the interesting lunchtime discussions.

Thank you to Vera Kovacevic for teaching me how to speak Serbian and Croatian. Thank you to

Paris Ning for your help with NFSS, and offering lots of food for thought in general. Thank you

to Vivek Dani for giving me your stapled booklet(s) of victory chits from Seven Wonders. They

will serve as a source of motivation for me to work towards my own victory chits. Thank you to

Hussain Masoom for inviting me to join GAPS. I definitely learned many transferrable skills

during my involvement. Roji Seevachandran, thank you for making undergrad life so memorable

#DSL #FEPmentorship.

iv

Thank you to Edward Nagato for coming out to our girls’ nights out. They wouldn’t

have been possible without you! Don’t ever lose that sassiness. Thank you for experiencing the

Bismack Biyombo and Jose Bautista eras with me #WeTheNorth #BautistaBatFlip #TDot. Let’s

hope there will be many more to come. Yes, we will make it to Tokyu hands and many other

places! And may the force of Miku always be with you.

Rebecca, Eunice, Joyce and Chun: From the very beginning, you girls have accompanied

me every step of the way, through sunny and rainy days. I am truly blessed to have wonderful

people like you.

Words cannot merely describe the gratitude I feel towards my family. To my mother and

my late father, your unconditional love and support have shaped me into the individual I am

today. Never have you doubted my ambitions, and this has allowed me to dream of achieving

what I wish to achieve in life. Thank you for believing in me and for all the sacrifices you have

made. To my brother, thank you for putting up with me and for all the scrumptious comfort food

you have made for me during my times of stress. Lastly, I would like to extend my deepest

appreciation to my aunts, uncles and cousins, for providing their support in times of need.

v

Table of Contents

Acknowledgments.......................................................................................................................... iii

Table of Contents .............................................................................................................................v

List of Tables ............................................................................................................................... viii

List of Figures ................................................................................................................................ ix

List of Appendices ......................................................................................................................... xi

Chapter 1: Introduction ....................................................................................................................1

1.1 The Arctic: from a net sink to a net source of atmospheric carbon ......................................1

1.2 Soil Organic Matter (SOM) ..................................................................................................5

1.3 Factors controlling SOM decomposition ..............................................................................6

1.3.1 Temperature ................................................................................................................6

1.3.2 Nitrogen and phosphorus amendments .......................................................................7

1.4 Major recalcitrant biomolecules in SOM: cutin, suberin and lignin ...................................10

1.5 Organo-mineral interactions................................................................................................13

1.6 Biomarker analysis of SOM ................................................................................................14

1.7 Characterization of SOM by nuclear magnetic resonance (NMR) spectroscopy ...............17

1.7.1 Solid-state 13

C cross polarization magic angle spinning (CPMAS) NMR ...............17

1.8 Research objectives .............................................................................................................18

1.9 References ...........................................................................................................................21

Chapter 2: Molecular-level characterization of Arctic soils after 32 years of in situ warming

and nitrogen + phosphorus fertilization ....................................................................................31

2.1 Abstract ...............................................................................................................................31

2.2 Introduction .........................................................................................................................32

2.3 Materials and methods ........................................................................................................35

2.3.1 Site description..........................................................................................................35

vi

2.3.2 Experimental design and sample collection ..............................................................35

2.3.3 Carbon (C) and nitrogen (N) analysis .......................................................................36

2.3.4 Biomarker extractions and gas chromatography-mass spectrometry (GC-MS)

analysis ...................................................................................................................36


C cross polarization magic angle spinning (CPMAS) NMR ...............38

2.3.6 Statistical analyses ....................................................................................................39

2.4 Results .................................................................................................................................40

2.4.1 Total carbon (C) and nitrogen (N) content ...............................................................40

2.4.2 Sources of biomarkers in Arctic soils .......................................................................42

2.4.3 SOM composition of warmed soils...........................................................................47

2.4.4 SOM composition of N+P fertilized soils .................................................................54

2.4.5 SOM composition of warmed + N+P fertilized soils ...............................................56


C NMR of Arctic soils .........................................................................58

2.5 Discussion ...........................................................................................................................61

2.5.1 Spatial heterogeneity and vertical mixing.................................................................61

2.5.2 Labile SOM components ..........................................................................................64

2.5.3 Recalcitrant SOM components .................................................................................66

2.5.4 Lignin-derived components ......................................................................................68

2.5.5 Implications on warming and N+P fertilizer addition on SOM degradation ............71

2.6 Conclusions .........................................................................................................................73

2.7 References ...........................................................................................................................75

Chapter 3: Evaluation of clay mineral and suberin and cutin protection of lignin in temperate

soils from surface horizons .......................................................................................................82

3.1 Abstract ...............................................................................................................................82

3.2 Introduction .........................................................................................................................83

3.3 Materials and methods ........................................................................................................85

vii

3.3.1 Description of soil samples and sampling sites ........................................................85

3.3.2 Determination of carbon (C) content ........................................................................88

3.3.3 Biomarker extractions and HF demineralization ......................................................88

3.3.4 Derivatization and gas chromatography-mass spectrometry (GC-MS) ....................91

3.3.5 Lignin-derived phenol analysis and calculation of % mineral-protected lignin

and % suberin- and cutin- protected lignin ............................................................91

3.4 Results and discussion ........................................................................................................93

3.4.1 Carbon (C) content ....................................................................................................93

3.4.2 Extraction yields of lignin-derived phenols and mineral protection of lignin ..........93

3.4.3 Suberin- and cutin-protected lignin ..........................................................................99

3.4.4 Implications for multilayer arrangement of organo-mineral interactions ...............102

3.5 Conclusions .......................................................................................................................104

3.6 References .........................................................................................................................106

Chapter 4: Conclusions and Future Directions ............................................................................112

4.1 Summary ...........................................................................................................................112

4.1.1 Molecular-level characterization of Arctic soils (Chapter 2) .................................112

4.1.2 Clay mineral, suberin and cutin protection of lignin (Chapter 3) ...........................114

4.2 Limitations and future work ..............................................................................................115

4.3 Research implications .......................................................................................................118

4.4 References .........................................................................................................................120

Appendices ...................................................................................................................................122

viii

List of Tables

Table 2-1: Total carbon and nitrogen content (%) and carbon: nitrogen ratios of the upper and

lower horizon soils of the control, warming, N+P fertilization and warming + N+P fertilization

treatments from all four blocks. .....................................................................................................41

Table 2-2: Solid-state 13

C CPMAS-NMR integration results with relative contribution (%) of the

four main carbon structures and calculated alkyl/O-alkyl ratios for the Control, Warming, N+P

Fertilization and Warming + N+P Fertilization treatments of the upper and lower horizon soils of

Block 1. ..........................................................................................................................................59

Table 3-1: Selected properties of four soils used in this study. ....................................................86

Table 3-2: Concentrations in μg/g soil of eight main lignin-derived phenols released after CuO

oxidation by comparing residues 1 (pre-HF) and 3 (post-HF). Values were determined from

triplicate samples (n = 3), unless otherwise indicated, followed by standard error. ......................94

Table 3-3: Percentages (%) of mineral protection of eight main lignin-derived phenols released

after CuO oxidationb, calculated by comparing the average yield of triplicate samples (n = 3), in

mg/g soil from residues 1 (pre-HF) and 3 (post-HF), followed by standard error. .......................97

ix

List of Figures

Figure 1-1: A soil profile for an Arctic soil core underlain by permafrost. ....................................2

Figure 1-2: Rising temperatures will result in permafrost thawing, which will cause carbon that

was previously frozen within permafrost to become susceptible to decomposition. The

accelerated degradation of carbon will increase carbon dioxide emissions into the atmosphere

and result in the permafrost carbon feedback. (Schaefer et al., 2014, reproduced with permission

from IOP Publishing Limited). ........................................................................................................4

Figure 1-3: Structures of the cutin (A) and suberin (B) biopolymers (Kögel-Knabner, 2002,

reproduced with permission from Elsevier). ..................................................................................11

Figure 1-4: Structural model of spruce lignin (Kögel-Knabner, 2002, reproduced with

permission from Elsevier). .............................................................................................................12

Figure 2-1: Concentrations (μg g-1

soil) of major SOM components from the upper horizon soil

samples after 32 years of warming, N+P fertilization and combined treatments. SOM

components include aliphatic and cyclic lipids, simple carbohydrates (galactose, glucose,

mannose), cutin- and suberin-derived lipids and lignin-derived phenols. All values are reported

as mean ± standard error (n = 2). Asterisks denote statistical significance from the control

treatment (P < 0.05). ......................................................................................................................43


soil) of major SOM components from the lower horizon soil

samples after 32 years of warming, N+P fertilization and combined treatments. SOM


mannose), cutin- and suberin-derived lipids and lignin-derived phenols. All values are reported

as mean ± standard error (n = 2). Asterisks denote statistical significance from the control

treatment (P < 0.05). ......................................................................................................................44


soil) of lignin-derived phenols (vanillyl, syringyl and

cinnamyl monomers) released from CuO oxidation of the upper horizon soil samples after 32

years of warming, N+P fertilization and combined treatments. All values are reported as mean ±

standard error (n = 2). Asterisks denote statistical significance from the control treatment (P <

0.05). ..............................................................................................................................................46



cinnamyl monomers) released from CuO oxidation of the lower horizon soil samples after 32

years of warming, N+P fertilization and combined treatments. All values are reported as mean ±

standard error (n = 2). Asterisks denote statistical significance from the control treatment (P <

0.05). ..............................................................................................................................................47

Figure 2-5: Plots of the acid to aldehyde ratios for syringyl (Ad/Al)s and vanillyl (Ad/Al)v

monomers of the upper horizon soils in each block. (Ad/Al)s = syringic acid/syringaldehyde;

(Ad/Al)v = vanillic acid/vanillin. Asterisks denote statistical significance from the control

treatment (P < 0.05). ......................................................................................................................50

x


monomers of the lower horizon soils in each block. (Ad/Al)s = syringic acid/syringaldehyde;

(Ad/Al)v = vanillic acid/vanillin. ...................................................................................................51

Figure 2-7: Plots of the syringyl/vanillyl monomers (S/V) and cinnamyl/vanillyl monomers

(C/V) ratios of the upper horizon soils in each block. Asterisks denote statistical significance

from the control treatment (P < 0.05). ...........................................................................................52


(C/V) ratios of the lower horizon soils in each block. Asterisks denote statistical significance

from the control treatment (P < 0.05). ...........................................................................................53

Figure 2-9: Solid-state 13

C CPMAS-NMR spectra of the Block 1 upper (a) and lower (b) horizon

soils of the Control, Warming, N+P Fertilization and Warming + N+P Fertilization treatments

with the four major spectral regions: alkyl (0-50 ppm), O-alkyl (50-110 ppm), aromatic and

phenolic (110-165 ppm) and carboxylic and carbonyl carbon (165- 215 ppm). ...........................60

Figure 3-1: Flowchart of the extraction sequence used to isolate the extracts and residues. Whole

soils were subject to solvent extraction to remove free lipids. Extract 1 was isolated from solvent

extraction and CuO oxidation. Extract 3 was isolated from solvent extraction, HF

demineralization and CuO oxidation. Extracts 2 and 4 were isolated in a similar fashion as

extracts 1 and 3 respectively, except with the addition of the base hydrolysis (BH) procedure. ..89

Figure 3-2: Percentage (%) of mineral protected-lignin in each VSC class from triplicate

samples (n = 3) of all four soils after HF treatment (comparison of extracts 2 and 4). NG,

Northern grassland soil; SG, Southern grassland soil; AGR, Agricultural soil; FOR, Forest soil.

Total vanillyls = vanillin, acetovanillone, vanillic acid; total syringyls = syringaldehyde,

acetosyringone, syringic acid; total cinnamyls = p-coumaric acid and ferulic acid. Error bars

indicate standard error....................................................................................................................95

Figure 3-3: Changes in average concentrations of lignin monomers from triplicate samples (n =

3) in the four soils: (a) Northern grassland; (b) Southern grassland; (c) Agricultural; (d) Forest,

suggesting suberin and cutin protection of lignin with mineral interference (comparison of

extracts 1 and 2) and without mineral interference (comparison of extracts 3 and 4). Error bars

indicate standard error..................................................................................................................101

xi

List of Appendices

Table A1: Concentrations (μg g-1

soil) of n-alkanes identified from the total solvent extracts of

the upper and lower horizon soils of the control, warming, N+P fertilization and warming +N+P

fertilization treatments. All values are reported as mean ± standard error (n = 2). .....................122


soil) of n-alkanols identified from the total solvent extracts of

the upper and lower horizon soils of the control, warming, N+P fertilization and warming +N+P

fertilization treatments. All values are reported as mean ± standard error (n = 2). .....................124


soil) of n-alkanoic acids and total aliphatic compounds

identified from the total solvent extracts of the upper and lower horizon soils of the control,

warming, N+P fertilization and warming +N+P fertilization treatments. All values are reported

as mean ± standard error (n = 2). .................................................................................................126


soil) of major compound classes identified in the total solvent

extracts (excluding aliphatic compounds) of the upper and lower horizon soils of the control,

warming, N+P fertilization and warming +N+P fertilization treatments. All values are reported

as mean ± standard error (n = 2). Numbers in bold denote statistical significance from the control

treatment (P < 0.05). ....................................................................................................................129


soil) of major SOM components released from the base

hydrolysis of the upper and lower horizon soils of the control, warming, N+P fertilization and

warming +N+P fertilization treatments. All values are reported as mean ± standard error (n = 2).

Numbers in bold denote statistical significance from the control treatment (P < 0.05). .............132

Table A6: Solid-state 13

C CPMAS-NMR integration results with relative contribution (%) of the

four main carbon structures and calculated alkyl/O-alkyl ratios for the Control treatments of the

upper horizon soils of each block. These soils were not treated with hydrofluoric acid. ............137

Figure A1: Solid-state 13

C CPMAS-NMR spectra of the upper horizon soils of the Control

treatments of each block with the four major spectral regions: alkyl (0-50 ppm), O-alkyl (50-110

ppm), aromatic and phenolic (110-165 ppm) and carboxylic and carbonyl carbon (165- 215

ppm). These soils were not treated by hydrofluoric acid. ............................................................138

1

Chapter 1: Introduction

1.1 The Arctic: from a net sink to a net source of atmospheric carbon

Permafrost is defined as soil which maintains sub-zero temperatures for at least two

consecutive years (Davidson et al., 2000; Schaefer et al., 2011). The Arctic has long been

regarded as a net sink of atmospheric carbon (C), storing approximately 1672 gigatons of C,

most of which is contained within permafrost (Tarnocai et al., 2009; Schaefer et al., 2011).

Permafrost can extend to several hundred meters below ground, but nearly 50% of

permafrost-derived C is situated in the upper 3 meters of the soil (Tarnocai et al., 2009;

Schaefer et al., 2011). Considerable amounts of frozen organic matter (OM) are contained in

permafrost (Tarnocai, 1997; Ping et al., 2008; Schaefer et al., 2011), which is believed to

have been buried by sedimentation processes since the last ice age (Schuur et al., 2008;

Schaefer et al., 2011). Vertical mixing between soil horizons caused by freezing and thawing

cycles, otherwise known as cryoturbation, may also have facilitated the burying of OM in

deep soil layers (Bockheim and Tarnocai, 1998; Davidson and Janssens, 2006). This may

explain why permafrost soils contain large stocks of global C (Davidson and Janssens, 2006).

A soil profile of an Arctic soil core is shown in Figure 1-1, which demonstrates that

permafrost soils are buried deep underneath the surface ground layer. The soil layer that

thaws annually during summer and re-freezes in winter is referred to as the active layer

(Schuur et al., 2008). Rising temperatures are expected to trigger active layer thickening as

the depth of permafrost layers becomes reduced by thawing. As a result, it is likely that

greater amounts of soil will be exposed to above-freezing seasonal temperatures (Schuur et

al., 2008).

2

Inc

rea

sin

g d

ep

th

Upper

Transition

Lower

Active layer

Permafrost

Figure 1-1: A soil profile for an Arctic soil core underlain by permafrost.

3

Greenhouse gases absorb solar radiation and re-emit it back to the Earth’s surface,

which prevents heat from escaping the atmosphere (IPCC, 2013). Elevated concentrations of

greenhouse gases are expected to increase the absorption and re-emission of radiation, which

will promote the trapping of heat within the Earth’s atmosphere (IPCC, 2013). Increasing

carbon dioxide (CO2) emissions from anthropogenic activities such as fossil fuel burning and

deforestation are projected to elevate Arctic temperatures by 7- 8°C by the end of the 21st

century (Schuur et al., 2013). Models also predict that during the same time period, the

current areal extent of near-surface (top 2- 3 m) permafrost will be reduced by 53%- 66% as

a result of rising temperatures (Schuur et al., 2013). As such, there is accruing evidence that

the Arctic is becoming a net source of atmospheric C (Qian et al., 2010; Hayes et al., 2011).

Warming-induced permafrost thawing will cause C that was previously frozen within the

permafrost to become susceptible to microbial decomposition (Osterkamp and Romanovsky,

1999; Serreze et al., 2000; Davidson and Janssens, 2006). The microbial breakdown of

permafrost-derived C will facilitate the release of CO2 (Schuur et al., 2013), which will in

turn exacerbate atmospheric warming and subsequently expedite permafrost thawing (Schuur

et al., 2008). The further release of CO2 from the decomposition of permafrost-derived C will

ultimately contribute to a positive climate feedback as outlined in Figure 1-2 (Hobbie et al.,

2002; Schaefer et al., 2011).

4

Figure 1-2: Rising temperatures will result in permafrost thawing, which will cause carbon

that was previously frozen within permafrost to become susceptible to decomposition. The

accelerated degradation of carbon will increase carbon dioxide emissions into the atmosphere

and result in the permafrost carbon feedback (Schaefer et al., 2014, reproduced with

permission from IOP Publishing Limited).

5

1.2 Soil Organic Matter (SOM)

Soil organic matter (SOM) is a heterogeneous mixture of organic materials derived

from plant, microbial and animal residues, at varying stages of decomposition (Feng and

Simpson, 2011). SOM retains essential nutrients and water in the soil, which are necessary

for supporting living organisms (Davidson and Janssens, 2006; Simpson and Simpson, 2012).

SOM also plays an important role in the C cycle (Sulzman et al., 2005), particularly in the

sequestration of soil C (Schlesinger, 1991; Batjes, 1996; Trumbore and Czimczik, 2008).

SOM consists of nearly two-thirds of the terrestrial C in the world (Schlesinger, 1991; Batjes,

1996), which is more C than in global vegetation and the atmosphere combined (Lehmann

and Kleber, 2015). It is therefore crucial to establish the fundamental factors that govern the

fate of SOM, such as chemical recalcitrance, physical aggregation and mineral protection

mechanisms (Amelung et al., 2009; Kögel-Knabner and Amelung, 2014). This will largely

benefit our understanding on how to maximize C sequestration in soils and to inhibit the

release of elevated C emissions from SOM decomposition (Lehmann and Kleber, 2015).

Information about molecular constituents in SOM will aid in understanding the fate of SOM

(Simpson and Simpson, 2012). Molecular-level data can be used to draw linkages to

macroscopic- and ecosystem-level responses, which will greatly facilitate the predictions of

potential ecosystem shifts (Simpson and Simpson, 2012). More specifically, this type of

research will provide information on the role of temperature and nutrient availability in SOM

decomposition processes and microbial community structures in Arctic tundra ecosystems.

This will be pertinent for elucidating the widespread shifts in terrestrial ecosystem functions

and biogeochemical cycles in response to rapidly rising temperatures (von Lützow and

Kögel-Knabner, 2009). Altogether, this knowledge will be instrumental in the development

of mitigation strategies for adaptation to a changing climate.

6

1.3 Factors controlling SOM decomposition

1.3.1 Temperature

The mean annual temperatures of mid- to high-latitude regions are projected to rise

by 3-5°C, with the greatest temperature increases expected in the high-latitude and Arctic

regions (Christensen et al., 2007). Understanding the temperature dependence of SOM

decomposition will ultimately help to determine the strength of the climate change feedback

loop as a function of atmospheric CO2 concentration (Ågren and Wetterstedt, 2007). Using a

simple model, Ågren and Wetterstedt (2007) verified the temperature dependence on the

breakdown of SOM, which ultimately contributes to the rate of C release. Enzyme activity

and interactions between OM and mineral complexes, which contribute to SOM degradation

processes, are governed by temperature (Ågren and Wetterstedt, 2007). With the onset of

global climate change, it is still unclear how shifts in the microbial community composition

will contribute to SOM decomposition patterns (Biasi et al., 2005; Frey et al., 2008; Feng and

Simpson, 2011). Rising temperatures may stimulate microbial activity which will expedite

SOM decomposition (Davidson and Janssens, 2006; Lehmann and Kleber, 2015).

The temperature sensitivity of decomposition varies between the labile and

recalcitrant SOM pools, where the decomposition of the labile SOM pool has been believed

to be more susceptible to temperature changes than the recalcitrant SOM pool (Schlesinger

and Andrews, 2000; Pautler et al., 2010). More specifically, labile OM compounds such as

carbohydrates and proteins are thought to be more sensitive to degradation at elevated

temperatures than recalcitrant OM compounds such as cutin-, suberin- and lignin-derived

compounds. Hence, the breakdown of these labile OM compounds may be a major source for

elevated CO2 emissions to the atmosphere (Schlesinger and Andrews, 2000; Pautler et al.,

7

2010). However, recent evidence suggests that SOM pools with slower turnover rates are

more sensitive to changes in temperature than those with faster turnover rates (Davidson and

Janssens, 2006; Conant et al., 2008; Craine et al., 2010; Lehmann and Kleber, 2015). For

example, results from a long-term field warming experiment of forest soils revealed that once

labile substrates have been depleted, microbes begin to decompose more recalcitrant

substrates, which leads to an enhancement in the overall decay of more stable SOM

compounds over long-term temperature changes (Frey et al., 2013). Short-term field warming

experiments (< 10 years) may produce bias towards temperature responses of labile SOM

pools compared to those from recalcitrant SOM pools (Davidson et al., 2000; Ågren and

Bosatta, 2002; Leifeld and Fuhrer, 2005; von Lützow and Kögel-Knabner, 2009).

Consequently, long-term ecosystem-level warming experiments are necessary to facilitate a

more mechanistic understanding of how recalcitrant and labile SOM pools respond to

climate-induced changes over broader time scales (Van Wijk et al., 2004).

1.3.2 Nitrogen and phosphorus amendments

The response of SOM decomposition to changes in nutrient availability is important

for determining net ecosystem C balance in a changing climate (Mack et al., 2004). Previous

modelling studies have suggested that elevated temperatures may stimulate SOM

decomposition and increase nutrient availability in soils (Shaver, 1992; Hobbie et al., 2002;

Mack et al., 2004). Nitrogen (N) and phosphorus (P) are essential nutrients for plant growth

(Vitousek et al., 2002; Shaver et al., 2014; Pisani et al., 2015). The cycling of N and P in

soils is strongly correlated to the C cycle (Kögel-Knabner and Amelung, 2014), but their

interactions with the C cycle have yet to be established (Shaver et al., 2006). N and P are of

current interest particularly in high latitude ecosystems because the availability of these

nutrients is believed to strongly constrain ecosystem gain of C (Hobbie et al., 2002). The

8

thawing of permafrost in tundra ecosystems is expected to expedite the release of N and P in

soils, which will likely enhance plant productivity (Hobbie et al., 2002). However, it is still

unclear how SOM biogeochemistry in Arctic ecosystems will be altered in response to

increased N and P availability in soils.

In the Arctic, the main sources of N inputs are bacterial fixation and rainfall and snow

deposition while the main losses of N are by leaching and denitrification (Shaver et al.,

2014). The mean residence time of N in soils is about 50 years (Schlesinger, 1991; Kögel-

Knabner and Amelung, 2014), which indicates that N is mostly conserved in soils. Numerous

experiments on N fertilization of soils have been conducted (Vitousek, 1982; Berg and

Matzner, 1997; Hobbie et al., 2000; Neff et al., 2002; Mack et al., 2004), where positive

(Waldrop et al., 2004; Bradford et al., 2008) and negative (Mack et al., 2004; Waldrop et al.,

2004; Bradford et al., 2008) correlations between N fertilization and SOM decomposition

have been observed. This may be due to the differences in responses between low- and high-

latitude ecosystems (Mack et al., 2004) and in soils from various environments (Bradford et

al., 2008). For example, N fertilization in agricultural soils was found to inhibit SOM

decomposition (Gregorich et al., 1996; McLauchlan, 2006; Bradford et al., 2008) but this was

not observed in grassland soils (Bradford et al., 2008). In boreal forests of Sweden and

Finland, long-term N fertilization resulted in the accumulation of soil C due to a decline in

SOM decomposition from a reduction in heterotrophic respiration (Hyvönen et al., 2008).

After 20 years of N fertilization in temperate forest soils, SOM decomposition was observed

to be suppressed by inhibited microbial activity (Frey et al., 2014). However, in a different

northern temperate forest, N fertilization caused a loss of soil C, which was likely attributable

to enhanced microbial activity (Waldrop et al., 2004). The contrasting responses to N

addition may also be ascribed to variations in the decomposition of various SOM pools

9

(Lavoie et al., 2011). N fertilization has been observed to stimulate the decomposition of

labile C but may suppress the decomposition of recalcitrant compounds (Berg and Matzner,

1997; Lavoie et al., 2011). Previous N addition experiments conducted in the Arctic tundra

have demonstrated that plant productivity and biomass accumulation in this ecosystem are

strongly limited by N availability (Shaver et al., 2001). However, it is still unclear how SOM

biogeochemistry in Arctic ecosystems will change in response to greater N availability.

Previous field experiments conducted in Arctic ecosystems have also observed that in

addition to N limitation, the plant productivity of tussock tundra vegetation may also be

limited by P availability (Shaver and Chapin, 1980; Shaver et al., 2001). The mean residence

time of organic P in soils is estimated to be between 350 to 2000 years (Paul and Clark,

1996; Kögel-Knabner and Amelung, 2014). From a 400-day long incubation study of

Alaskan tundra soils, Shaver et al. (2006) found that although P content was higher in

fertilized soils, C losses were predominantly due to N fertilization and not P fertilization.

This suggests that N may play a more vital role in SOM degradation processes and C cycling

than P in high-latitude environments (Shaver et al., 2006).

Mack et al. (2004) reported that N and P fertilization caused a net ecosystem loss of C

despite a twofold increase in litter over 20 years in an Arctic ecosystem. However, when the

same soils were analyzed in an incubation study, the net loss of C was considerably less

(Schimel and Weintraub, 2003; Shaver et al., 2006). The presence of live vegetation and

inputs of fresh litter may have played an important role in microbial growth and uptake,

which may have increased C mineralization rates in the field experiment (Shaver et al.,

2006). Although a general consensus has yet to be reached regarding the general roles of N

and P in SOM degradation processes and C cycling, it is important to acknowledge that these

10

roles may differ among ecosystems. N and P fertilization may expedite SOM decomposition

processes especially in nutrient-limited environments (Shaver et al., 2014).

1.4 Major recalcitrant biomolecules in SOM: cutin, suberin and lignin

With rising global temperatures (Schaefer et al., 2011), much research has been

dedicated towards examining how elevated amounts of C released into the atmosphere from

SOM degradation processes will contribute to global warming (Mack et al., 2004). As a

result, there is also a pressing need to better understand how SOM can be stabilized to

maximize C sequestration in soils (Christensen, 2001; Six et al., 2002; Clemente and

Simpson, 2013). The stability of OM may be governed by a resistance to degradation due to

the structural properties of the OM compounds, which is also known as inherent chemical

recalcitrance (Six et al., 2002; Lorenz et al., 2007; Clemente et al., 2011). For example,

suberin and cutin, components of SOM, experience long residence times in soil, possibly due

to the nature of their molecular structures such as alkyl C chains in lipids and aromatic

structures which are more difficult to break down (Mikutta et al., 2006; Lorenz et al., 2007;

Clemente et al., 2011). Cutin forms the macromolecular framework of the plant cuticle,

which protects plant surfaces against aridity (Kögel-Knabner, 2002). Cutin is produced in the

epidermis of leaves of vascular plants (Holloway, 1982; Otto and Simpson, 2006b). The cutin

polymer (Figure 1-3a) is believed to be composed of di- and tri-hydroxy and epoxy fatty

acids with C16 and C18 chain lengths (Kögel-Knabner, 2002), which are linked by ester bonds

(Kolattukudy, 1981; Kögel-Knabner, 2002). Suberin is a cell wall component of cork cells

which forms the periderm layer of subterranean parts of woody plants (Kögel-Knabner,

2002). Suberin is found in the periderm of roots and barks of vascular plants and consists of

long-chain (C20-C32) aliphatic lipids, diacids and ω-hydroxy acids and some phenolic

11

moieties (Kolattukudy and Espelie, 1989; Bernards, 2002; Otto and Simpson, 2006b). The

structure of the suberin polymer is shown in Figure 1-3b.

Figure 1-3: Structures of the cutin (A) and suberin (B) biopolymers (Kögel-Knabner, 2002,

reproduced with permission from Elsevier).

Lignin is the second most abundant biomolecule in vascular plants after

polysaccharides (Derenne and Largeau, 2001), and provides strength and rigidity to plant

structures (Brown, 1961; Kirk and Farrell, 1987; Argyropoulos and Menachem, 1997;

Higuchi, 2006). Lignin is a biopolymer composed of three types of phenylpropanoid units:

vanillyl, syringyl and cinnamyl (Adler, 1977; Derenne and Largeau, 2001). The structure of

lignin is illustrated in Figure 1-4. In soils, the concentration of compounds from each of these

classes is not only characteristic of its vegetation source, but is also indicative of the degree

of degradation in organic and mineral horizons (Hedges and Mann, 1979). Lignin was

previously thought to be biochemically recalcitrant (Berg and Staaf, 1980; Lehmann and

Kleber, 2015) due to its aromaticity (Feng and Simpson, 2011). In comparison to

macromolecules such as cellulose and hemicellulose, lignin has been thought to be more

resistant to microbial degradation because only white-rot and brown-rot fungi are able to

12

completely decompose lignin to CO2 (Kögel-Knabner, 2002). However, recent evidence

suggests that as long as lignin is easily accessible to microbial attack, lignin can be degraded

(Klotzbücher et al., 2011; Lehmann and Kleber, 2015). In comparison to suberin and cutin,

lignin is believed to be less resistant to degradation (Mikutta et al., 2006; Lorenz et al., 2007;

Clemente et al., 2011).

Figure 1-4: Structural model of spruce lignin (Kögel-Knabner, 2002, reproduced with

permission from Elsevier).

13

1.5 Organo-mineral interactions

In addition to chemical recalcitrance, SOM can also be stabilized through association

with clay mineral surfaces, which likely protects SOM from microbial degradation (Baldock

and Skjemstad, 2000; Eusterhues et al., 2003; Kaiser and Guggenberger, 2003; Mikutta et al.,

2006). Sorptive interactions between OM and clay mineral surfaces are governed by clay

mineralogy and the composition of the OM sorbate (Asselman and Garnier, 2000; Chi and

Amy, 2004; Feng et al., 2005; Mikutta et al., 2007; Ghosh et al., 2009; Clemente and

Simpson, 2013). From previous sorption studies, polymethylene structures have been

observed to preferentially sorb to kaolinite and montmorillonite (Feng et al., 2005; Simpson

et al., 2006; Ghosh et al., 2009; Clemente et al., 2011), which suggests that aliphatic

compounds may be selectively preserved through sorption (Clemente et al., 2011). In

addition, proteins have also been observed to sorb onto montmorillonite (Feng et al., 2005;

Ghosh et al., 2009; Clemente et al., 2011) while carboxyl groups from OM have been

observed to sorb to goethite (Ghosh et al., 2009; Clemente et al., 2011). In addition, the

ability of minerals to preserve OM is regulated by the number of sites on the mineral surface

which are available for sorption (Kaiser and Guggenberger, 2003). Based on the findings

from previous studies, recalcitrant biomacromolecules in soil such as cutin, suberin and

lignin, are likely to be associated with clay mineral surfaces by sorptive interactions (Bahri et

al., 2006; Mikutta et al., 2006; Rumpel et al., 2006; Heim and Schmidt, 2007; Hernes et al.,

2013; Lin and Simpson, 2016). Lignin is of particular interest because they may represent an

important part of stabilized OM when associated with clay minerals (Thevenot et al., 2010;

Clemente and Simpson, 2013).

Various chemical extraction methods involving desorbing, hydrolyzing and oxidizing

reagents have been employed to extract OM bound to clays and Fe and Al oxides (von

14

Lützow et al., 2007). Some methods include acid hydrolysis with hydrochloric acid,

oxidative degradation with sodium hypochlorite or disodium peroxodisulfate (von Lützow et

al., 2007). In particular, mineral dissolution with hydrofluoric (HF) acid has been commonly

used to release mineral-bound OM (Schmidt et al., 1997; Eusterhues et al., 2003; Mikutta et

al., 2006; Rumpel et al., 2006). HF reacts with silicates and oxides to form soluble silicate

minerals but OM is presumed to remain intact (Eusterhues et al., 2007). The dissolution of

minerals is achieved based on the breakdown of Si-O bonds (Rumpel et al., 2006). HF

treatment has also been used extensively for the removal of paramagnetic substances in SOM

before conducting solid-state 13

C nuclear magnetic resonance (NMR) spectroscopy

(Zegouagh et al., 2004; Rumpel et al., 2006).

1.6 Biomarker analysis of SOM

Biomarkers are defined as tracers of biosynthesized organic molecules and are

analogous to OM fingerprints (Simpson and Simpson, 2012). Their unique C skeleton

information remains intact during abiotic or biotic degradation and can be traced to a specific

plant or microbial source (Amelung et al., 2009; Simpson and Simpson, 2012) and a marine

or terrestrial environment (Hedges et al., 2000; Kögel-Knabner, 2002; Amelung et al., 2009;

Feng and Simpson, 2011). Biomarker analyses can provide information on the degree of

SOM degradation which can be used to understand how SOM may be altered in response to

global environmental changes such as elevated temperatures (Feng and Simpson, 2011).

Sequential biomarker extractions can be used to isolate SOM components which are

indicative of SOM sources (gymnosperm and angiosperm plants) and their degradation stage

in soil (Otto and Simpson, 2007; Feng and Simpson, 2011). Extraction with organic solvents

can remove unbound lipids including n-alkanes, n-alkanols, n-alkanoic acids, steroids and

15

terpenoids, along with carbohydrates (Otto and Simpson, 2007). Short-chain n-alkanes, n-

alkanols and n-alkanoic acids (< C20) typically originate from fungi and bacteria (Dinel et al.,

1990; Collister et al., 1994; Bourbonniere et al., 1997; Amelung et al., 2009) while their

long-chain counterparts (> C20) originate from the cuticle waxes of terrestrial plants (Collister

et al., 1994; Amelung et al., 2009). Subsequent to solvent extraction, base hydrolysis can be

used to release ester-bound monomers that originate from suberin and cutin biopolymers

(Otto and Simpson, 2006b). The proportion of bound lipids in soil that are derived from

leaves vs. roots can be estimated using a ratio based on the amount of extractable cutin and

suberin monomers (Kögel-Knabner et al., 1989; Otto and Simpson, 2006b). Following base

hydrolysis, copper (II) oxide (CuO) oxidation is used to cleave ether bonds which releases

lignin-derived phenols that can be divided into three structural classes: vanillyl, syringyl and

cinnamyl phenols (Otto and Simpson, 2006a). Specific types of plants are known to produce

certain types of lignin-derived phenols (Hedges and Mann, 1979). For example,

gymnosperms produce vanillyl phenols, angiosperms produce both vanillyl and syringyl

phenols while nonwoody vascular plants produce cinnamyl phenols (Hedges and Mann,

1979). Based on the yields of each lignin-derived phenol class from the CuO oxidation

extracts, the ratios of syringyl/vanillyl phenols (S/V) and cinnamyl/vanillyl phenols (C/V)

can be used to differentiate lignin inputs from gymnosperm vs. angiosperms sources and

from woody vs. nonwoody vascular plants (Hedges and Mann, 1979; Hedges and Ertel,

1982; Otto and Simpson, 2006a). Acid to aldehyde (Ad/Al) ratios of vanillyl and syringyl

phenols, which reflect the level of oxidation of lignin side chains (Derenne and Largeau,

2001), increase with progressive lignin oxidation (Ertel and Hedges, 1985; Hedges et al.,

1988; Opsahl and Benner, 1995; Otto and Simpson, 2006a).

16

Gas chromatography-mass spectrometry (GC-MS) is commonly used for biomarker

analyses (Simoneit, 2005). The gas chromatograph (GC) is the compound-separation

instrument while the mass spectrometer (MS) analyzes compounds based on their mass-to-

charge ratios (Simoneit, 2005). After the sample is volatilized in the GC, analytes in the

sample are partitioned between a gaseous mobile phase and a column stationary phase such

as poly(dimethylsiloxane). Helium is often used as a carrier gas to transport analytes through

the column. Separation is achieved based on the analyte’s affinity for the mobile and

stationary phases. Analytes that interact more strongly with the mobile phase will travel

faster through the column (Skoog et al., 2007). Extracts are derivatized to convert polar

functional groups into species that are amenable for GC-MS analysis (Horning et al., 1969).

A common derivatization method is the conversion of reactive hydrogen atoms to their

trimethylsilyl derivatives using N,O-bis-(trimethylsilyl) trifluoroacetamide (BSTFA) and

pyridine (Horning et al., 1969; Otto et al., 2005). After separation in the chromatography

column, molecules are ionized before they are introduced into the MS detector. The most

common ionization method for biomarkers is by electron impact ionization, where vaporized

molecular compounds are ionized by the bombardment of energetic electrons (Skoog et al.,

2007). Ionized molecules are then detected by quadrupole mass analyzers (Skoog et al.,

2007). Quadrupole mass analyzers consist of a set of four electrodes and resolve ions based

on their mass-to-charge ratios. Only ions within a specific mass region (i.e. 50 – 650 Da)

pass through the mass analyzer (Miller and Denton, 1986). Based on unique fragmentation

patterns of each compound, the mass spectra can be used for compound identification.

17

1.7 Characterization of SOM by nuclear magnetic resonance (NMR)

spectroscopy

NMR spectroscopy is a nondestructive spectroscopic technique which can provide a

compositional overview of complex samples such as SOM (Simpson et al., 2011). NMR can

yield information about the abundance of functional groups (Otto and Simpson, 2007).

Biomarker methods only analyze the extractable portion of SOM, whereas NMR techniques

can offer a structural overview of SOM composition. Therefore, biomarkers and NMR serve

as complementary methods to present an overall picture of SOM biogeochemistry (Feng and

Simpson, 2011).


C cross polarization magic angle spinning (CPMAS) NMR

Solid-state 13

C NMR is the most commonly used NMR technique in soil analyses

because it requires minimal amounts of sample and provides general information regarding

the distribution of C structures in the sample (Kögel-Knabner, 2000; Feng et al., 2010; Feng

and Simpson, 2011). Pre-treatment of soils with HF removes paramagnetic materials and

dissolves minerals to concentrate SOM (Simpson et al., 2011). Since 13

C only represents

1.13% of total C found in nature, 13

C NMR may not detect subtle changes in SOM

composition (Feng and Simpson, 2011). Spinning at the magic angle (54.74° with respect to

the applied magnetic field) along with the suppression of 1H-

13C dipolar interactions by high

power decoupling collectively reduces chemical shift anisotropy and line broadening

(Andrew et al., 1958; Simpson et al., 2011). Cross polarization magic angle spinning

(CPMAS) is considered a semi-quantitative technique because it relies on the magnetization

transfer between protons and the C that are directly attached. Hence, C which are remote

from protons may not be detected as well by CPMAS (Conte et al., 2004). A 13

C NMR

18

spectrum of SOM can be divided into four major spectral regions: aliphatic, 0 -50 ppm; O-

alkyl, 50-110 ppm; aromatic and phenolic, 110-165 ppm and carboxyl and carbonyl C, 165-

220 ppm (Baldock et al., 1992; Simpson et al., 2008). Based on the abundances of the alkyl

and O-alkyl C from the 13

C CPMAS NMR spectrum, the alkyl/O-alkyl ratio has been

observed to increase with progressive SOM degradation (Baldock and Preston, 1995;

Sjögersten et al., 2003; Simpson et al., 2008).

1.8 Research objectives

With the onset of global environmental change, there is a growing concern over the

extent of CO2 emissions released from the thawing of permafrost soils (Schaefer et al., 2011).

The amount of C stored in soils is governed by the stabilization mechanisms which are

controlled by the chemical, physical and biological interactions within the soil matrix

(Kögel-Knabner et al., 2008). Therefore, it is important to establish the mechanisms which

stabilize and destabilize SOM, and to recognize their roles in C sequestration in soils. SOM

could be stabilized by interactions with mineral surfaces (Baldock and Skjemstad, 2000) but

could be degraded with environmental changes such as elevated temperatures (Feng et al.,

2008) and increased nutrient availability (Mack et al., 2004). Since the chemical composition

of SOM is a useful predictor of C turnover (Ågren and Bosatta, 1996; Shaver et al., 2006), a

molecular-level investigation will deepen our understanding of how SOM biogeochemical

shifts can be used to predict changes in C cycling in response to global environmental

change. The objectives of this research were to characterize SOM:

1. In whole soils from an Arctic ecosystem which were subjected to 32 years of

manipulated soil warming and N and P fertilization. It is hypothesized that N and P

fertilization will be a stronger contributing factor to SOM decomposition than soil

19

warming. However, it is predicted that the combined treatments of soil warming and

N and P fertilization will expedite SOM decomposition processes the most.

2. In grassland, agricultural and forest soils from temperate ecosystems which are rich in

clay minerals and Fe and Al oxides. This will verify the role of mineral protection and

identify possible OM-OM interactions between major recalcitrant biopolymers (i.e.

suberin and cutin protection of lignin). We hypothesize that mineral protection will

play a more major role in stabilizing SOM, compared to OM-OM interactions.

To determine the source and degradation state of SOM in whole soils from an Arctic

ecosystem after 32 years of soil warming and nutrient addition, sequential biomarker

extractions were employed followed by GC-MS analysis. Samples were also investigated by

solid-state 13

C NMR to present an overall picture of the SOM biogeochemistry in these

Arctic soils. SOM in the organic and mineral horizon soils from Toolik Lake, Alaska were

extracted by solvent extraction, base hydrolysis and CuO oxidation. Biomarkers were

identified and quantified by GC-MS. Solid-state 13

C NMR was employed to characterize the

structural components of SOM in these samples. Results are presented in Chapter 2.

SOM in grassland, agricultural and forest soils from Alberta, Ontario and British

Columbia, Canada, respectively were characterized using similar molecular-level techniques.

These soils were subjected to HF demineralization to investigate organo-mineral interactions.

Biomarkers were identified and quantified after base hydrolysis and CuO oxidation

extraction procedures, followed by GC-MS analysis. Results are presented in Chapter 3.

Overall, the aim of this thesis is to determine the possible factors which govern SOM

stabilization and degradation mechanisms and how they contribute to ecosystem storage of

soil C. In Arctic tundra soils, rising temperatures and nutrient addition may accelerate SOM

20

degradation processes, triggering elevated levels of CO2 emissions to the atmosphere and

potentially contributing to the positive feedback of global warming. In temperate soils, clay

mineral interactions may be able to protect SOM from degradation, thus contributing to the

overall stabilization of SOM. Our goal is to employ biomarkers and NMR methods to predict

possible SOM biogeochemical shifts with global warming and how SOM stabilization

mechanisms could potentially contribute to our understanding of C sequestration in soils.

21

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Chapter 2: Molecular-level characterization of Arctic soils

after 32 years of in situ warming and nitrogen + phosphorus

fertilization

2.1 Abstract

Recent studies have revealed that rising global temperatures will cause permafrost to

thaw. This is expected to accelerate the decomposition of soil organic matter (SOM) that was

once frozen within the permafrost and will cause elevated amounts of carbon dioxide (CO2)

to be released into the atmosphere. Nutrients such as nitrogen (N) and phosphorus (P) may

also be released from the thawing of permafrost soils. However, it is unclear how elevated

global temperatures and the subsequent release of N and P from thawed permafrost will alter

SOM biogeochemistry in Arctic ecosystems. In this study, we employed biomarker and

nuclear magnetic resonance (NMR) techniques to investigate the degradation of SOM in the

upper and lower horizons of soils from a Long-Term Ecological Research site located in

Toolik Lake, Alaska, USA, after 32 years of warming, nitrogen + phosphorus (N+P)

fertilization and warming + N+P fertilization. Biomarker analyses revealed variable

responses among the replicate blocks of each treatment, which suggests there was

considerable landscape heterogeneity. Warming, N+P fertilization and their combined

treatment facilitated the accumulation of cyclic lipids in the upper horizon soils. Cutin-

derived biomarkers in the upper horizon slightly increased with each treatment but NMR

analysis did not reveal any enrichment in alkyl C compounds. The degradation of cutin was

not enhanced in these Arctic soils, likely due to its chemical recalcitrance. The abundances of

suberin-derived components decreased in the lower horizon in response to elevated

temperatures and N+P fertilization, but these were not statistically significant changes.

32

Particularly in the lower horizons of all the treatments, low abundances of simple sugars

(glucose, galactose and mannose) coupled with high intensities of the O-alkyl C signals from

the solid-state 13

C NMR spectra suggest ample amounts of cellulose inputs. Lignin oxidation

was not promoted by any of the treatments except in the upper horizon of block 4, where

lignin was most oxidized in the N+P fertilization treatment and least oxidized in the warming

+N+P fertilization treatment. Overall, our results demonstrate that after 32 years of warming

and increased nutrient availability at Toolik Lake, Alaska, SOM composition was altered in

some plots but not all. This study shows the importance of landscape properties which

control plant-soil interactions and their subsequent responses to climate change.

2.2 Introduction

The Arctic has long been regarded as a net sink of atmospheric carbon dioxide (CO2),

storing approximately 1672 gigatons of carbon (C; Tarnocai et al., 2009; Schaefer et al.,

2011). However, recent studies have revealed that the Arctic is transitioning from a net sink

to a net source of atmospheric C (Qian et al., 2010; Hayes et al., 2011). Projected rising

temperatures are expected to accelerate permafrost thawing and active layer deepening

(Schaefer et al., 2011). Higher temperatures may further stimulate microbial activity

(Schaefer et al., 2011) causing labile C that was once frozen within permafrost to become

more susceptible to decomposition (Osterkamp and Romanovsky, 1999; Serreze et al., 2000;

Davidson and Janssens, 2006). As a result, elevated fluxes of CO2 from Arctic soils will be

released into the atmosphere (Mack et al., 2004; Koven et al., 2011; MacDougall et al.,

2012). Elevated CO2 emissions will absorb more infrared radiation (Hobbie, 1996) and

amplify atmospheric warming and permafrost degradation, which will ultimately result in a

positive climate feedback (Schlesinger and Andrews, 2000; Davidson and Janssens, 2006). In

addition, permafrost soils of northern ecosystems contain large amounts of organic nitrogen

33

(N) and phosphorus (P) that are currently unavailable to plants (Shaver et al., 2006). The

thawing of permafrost and stimulation of microbial activity with warming may promote net

mineralization of N and P in soils (Chapin, 1983; Nadelhoffer et al., 1992; Aerts, 2006;

Deslippe et al., 2011). These augmented rates of conversion from organic to inorganic forms

of N and P, which are more readily usable by plants, could enhance plant productivity

(Shaver et al., 2006). This may alter N and P cycling in Arctic ecosystems and induce shifts

in plant community composition such that the growth of woody shrubs will be enhanced

(Hobbie et al., 2002). However, it is still unclear how global environmental changes such as

rising temperatures and enhanced nutrient availability may alter soil organic matter (SOM)

composition in Arctic ecosystems.

Toolik Lake, Alaska is located in the northern foothills of the Brooks Range and has

been studied intensively as part of the Long-Term Ecological Research Network (Hobbie,

1995; Shaver, 1996; Van Wijk et al., 2004). Various long-term ecosystem-scale experiments

have been conducted at Toolik Lake, mostly to examine the responses in aboveground plant

biomass to experimental manipulations such as warming, shading and nitrogen + phosphorus

(N+P) fertilization (Shaver and Jonasson, 1999; Van Wijk et al., 2004; Shaver et al., 2006;

Deslippe et al., 2011). Aboveground plant biomass has been observed to be more responsive

to N+P fertilization than to soil warming, suggesting that the Arctic ecosystem at Toolik

Lake is limited by N+P availability (Shaver et al., 2014). For example, 15 years of N+P

addition stimulated an accumulation of aboveground biomass of Arctic vegetation. Soil

warming for the same duration also promoted increases in aboveground biomass, but to a

lesser extent than N+P fertilization (Shaver and Jonasson, 1999). Hobbie et al. (1996)

conducted a field warming study at Toolik Lake and found that a soil temperature elevation

of 5°C enhanced soil C loss. Results from a 20-year N+P fertilization experiment also

34

conducted at Toolik Lake showed a net ecosystem loss of C, despite enhanced plant

productivity (Mack et al., 2004). These findings suggest that elevated temperatures and

fertilization may enhance SOM decomposition.

The primary objective of this study was to investigate the extent of SOM degradation

in soils from Toolik Lake after long-term warming and nutrient addition. Based on earlier

studies (Hobbie, 1996; Mack et al., 2004), elevated temperatures and N+P fertilization were

hypothesized to enhance SOM decomposition. However, due to the N and P-limited

conditions at Toolik Lake (Shaver et al., 2014), N+P fertilization was hypothesized to

expedite SOM decomposition to a greater extent than soil warming alone. Since elevated

temperatures and increased nutrient availability both enhance SOM decomposition processes

according to previous observations (Hobbie, 1996; Mack et al., 2004), it was hypothesized

that SOM from the warming and N+P fertilization treatment would be the most degraded. To

test this hypothesis, soil samples were collected from the Toolik Lake Long-Term Ecological

Research site in Alaska, USA after 32 years of warming, N+P fertilization and warming

+N+P fertilization. We employed three biomarker extraction techniques (solvent extraction,

base hydrolysis and CuO oxidation) followed by gas chromatography-mass spectrometry

(GC-MS) analysis to isolate and quantify SOM biomarkers, which provide insight into the

sources and stage of SOM decomposition (Otto and Simpson, 2005; 2006a; 2006b). These

extraction techniques yield free compounds, cutin- and suberin-derived compounds and

lignin-derived phenols, respectively (Otto and Simpson, 2007). GC-MS was used to identify

and quantify biomarkers, while solid-state 13

C nuclear magnetic resonance (NMR)

spectroscopy was used to examine changes in overall SOM composition (Simpson and

Simpson, 2012) in response to soil warming and N+P fertilization. The complementary

35

information obtained from biomarker and NMR methods can offer insight into SOM

responses to global climate change (Feng and Simpson, 2011).

2.3 Materials and methods

2.3.1 Site description

Samples were collected from the Long-Term Ecological Research site near Toolik

Lake, Alaska, USA (68°38’N, 149°34’W, elevation 780 m), in the northern foothills of the

Brooks Range. This moist acidic tundra site is situated on a gentle north-facing slope

(Clemmensen et al., 2006). The dominant vegetative species are the moist acidic tussock-

forming sedge Eriophorum vaginatum and the deciduous dwarf shrub Betula nana, which

make up approximately 20% of the aboveground vascular plant biomass (Clemmensen et al.,

2006). These tundra soils are classified as histic pergelic cryaquepts (Mack et al., 2004),

which are characterized by a thick upper horizon 10-20 cm in depth, with a pH of 3.7 (Shaver

et al., 2014). The lower horizon, around 20 cm in depth, has a silty texture (Mack et al.,

2004) with a pH of 4.6 and is underlain by continuous permafrost 200 m in depth (Shaver et

al., 2014). The mean air temperature during the June-August growing season is 9.3°C (Mack

et al., 2004) and the mean annual precipitation is approximately 350 mm (Clemmensen et al.,

2006). Soil surface temperatures range from 10 to 20°C in the summer, but the bottom of the

thawed active layer is consistently at 0°C.

2.3.2 Experimental design and sample collection

Established in 1982, the experiment is composed of four replicate blocks that have

been randomly assigned one of four treatments: control, warming, N+P fertilization and

warming + N+P fertilization. Each block contained four 5 m x 20 m plots and five 20 cm x

20 cm quadrats were harvested from each plot. Fertilizer treatments, which consist of 10 g

36

m-2

additions of N as NH4NO3 and 5 g m-2

additions of P as P2O5, were applied as pellets

after snowmelt annually. Warming was accomplished in greenhouses by elevating the

average air temperature by 2°C and average soil temperature was increased by 1°C (up to 40

cm in depth).

Soil samples were collected in September 2014. Fresh litter was removed and organic

monoliths (8 x 8 cm) were collected and separated into upper horizon (0-5 cm), upper-lower

transition horizon (5 cm to top of the lower horizon) and lower horizon soils (sampled to the

permafrost). The transition horizon soils were not analyzed in this study because not all the

treatment plots within each block consisted of a transition layer between the upper and lower

horizons (Table 2-1). Also, the lower horizon soil of the warming +N+P fertilization

treatment of block 3 was not analyzed because this sample was unavailable. All samples

were frozen immediately after sampling to prevent any degradation and were transported to

the laboratory to be freeze-dried. Prior to analysis, the soil samples were passed through a 2

mm sieve and were then ground using a mortar and pestle.

2.3.3 Carbon (C) and nitrogen (N) analysis

C and N analysis of the samples was performed at the Natural Resource Ecology

Laboratory at Colorado State University in Fort Collins, Colorado, USA. Soil samples were

dried at 60°C and then ground to a fine powder using a mortar and pestle under liquid N2.

Total C and total N were measured using the Carlo Erba NA 1500 elemental analyzer.

2.3.4 Biomarker extractions and gas chromatography-mass spectrometry (GC-

MS) analysis

Sequential biomarker extractions (solvent extraction, base hydrolysis and CuO

oxidation) were performed in duplicate on the soil samples to isolate free lipids, cutin- and

37

suberin-derived ester-bound lipids and lignin-derived phenols respectively (Hedges and Ertel,

1982; Otto and Simpson, 2005; 2006a; 2006b). 0.5 g of upper horizon soils and 1.0 g of

lower horizon soils were weighed out and sonicated in DCM, DCM: MeOH (1:1 v/v) and

MeOH sequentially using 180 ml of each solvent. Solutions were filtered through glass fibre

filters (Whatman GF/A and GF/F), concentrated by rotary evaporation and dried under N2.

Air-dried residues (0.1 – 0.4 g) were heated at 100°C for 3 h in Teflon-lined bombs with 20

ml of 1M methanolic KOH. After acidification to pH 1 with 6M HCl, hydrolysable lipids

were recovered by liquid-liquid extraction with diethyl ether. The extracts were dried with

anhydrous Na2SO4 to remove remaining water, concentrated by rotary evaporation and dried

under N2. Air-dried residues from base hydrolysis were added to 1 g of copper (II) oxide

(CuO), 100 mg of ammonium (II) iron sulfate hexahydrate [Fe(NH4)2(SO4)•6H2O] and 15 ml

of 2 M NaOH and heated at 170°C for 2.5 h. Acidification to pH 1 was accomplished with 6

M HCl and extracts were then stored for 1 h at room temperature in the dark to prevent

reactions of cinnamic acids. After centrifugation, solid phase extraction (Oasis HLB

Cartridges 60 mg, 3 ml capacity; Waters, Milford, MA) was used was used to isolate lignin-

derived phenols from extracts (Kaiser and Benner, 2012). The cartridges were pre-

conditioned twice with 2 ml of methanol and 2ml of deionized water under gravity flow

before extracts were loaded through the cartridges at a flow rate of 1 ml/min under vacuum

(Supelco Preppy™

12-port vacuum manifold, St. Louis, MO). Cartridges were rinsed three

times with 0.5 ml of water/ MeOH 70/30 v/v%. The cartridges were then dried under vacuum

for 10 min before elution with three rinses of 0.5 ml of dichloromethane/methyl

acetate/pyridine 70/25/5 v/v/v% followed by two rinses of 0.5 ml of methanol. The eluates

were then combined, put through Pasteur pipet columns packed with anhydrous Na2SO4 to

remove any remaining water and then dried under N2. The solvent and CuO extracts were

converted to trimethylsilyl (TMS) derivatives by reaction with 100 μl of N,O-

38

Bis(trimethylsilyl) trifluoroacetamide (BSTFA) and 10 μl of pyridine at 70°C for 1.5 h. The

extracts from base hydrolysis were first derivatized with N,N-dimethylformamide dimethyl

acetal and then with BSTFA and pyridine. After cooling, hexane was added to dilute the

extracts prior to GC-MS analysis. Samples were analyzed using an Agilent model 6890 N

gas chromatograph (GC) coupled to an Agilent model 5973 quadrupole mass selective

detector with an Agilent model 7683 autosampler. The GC was fitted with a HP5-MS fused

silica capillary column (30 m x 0.25 mm i.d., 0.25 µm film thickness). The GC oven

temperature was held at 65°C for 2 min, increased to 300°C at a rate of 6°C per min and held

at 300°C for 20 min. The injector temperature was set at 280°C. Helium was used as the

carrier gas. The mass spectrometer was operated in electron impact ionization mode with

ionization energy of 70 eV and scanned from 50 to 650 Da. Identification of individual

compounds was achieved by comparison of mass spectra to those of authentic standards

along with NIST and Wiley 275 MS library data. Quantification was performed with the

following external standards: tetracosane, behenyl alcohol (TMS ester), methyl tricosanoate

and ergosterol (TMS ester) for the solvent-extractable lipids; methyl tricosanoate for the

hydrolysable lipids; syringic acid and syringaldehyde (TMS esters) for the lignin-derived

phenols.


C cross polarization magic angle spinning (CPMAS) NMR

Preliminary NMR experiments were conducted on the control soils of each block. For

analysis by solid-state 13

C cross polarization magic angle spinning (CPMAS) NMR, 100 mg

of each soil were packed into 4 mm zirconium rotors with Kel-F caps. The spectra were

acquired on a 500 MHz Bruker BioSpin Avance III spectrometer (Bruker BioSpin,

Rheinstetten, Germany) equipped with a 4 mm H-X MAS probe, using a ramp-CP pulse

program (Conte et al., 2004; Pisani et al., 2015) with a spinning rate of 11 KHz and a ramp-

39

CP contact time of 1 ms. Preliminary data were first collected using a recycle delay of 1 s,

but the spectra exhibited low signal-to-noise ratios so a recycle delay of 3 s was used. Since

the signal-to-noise ratios of the spectra from the preliminary experiment did not substantially

improve even with a longer recycle delay, soils were treated with 10% hydrofluoric (HF)

acid as discussed below before NMR analyses. Since the preliminary experiments showed

that the spectra of the control treatments of each block were dissimilar, NMR analyses were

performed only on the soil samples from block 1. Whole soil samples from each treatment in

block 1 (20 g of upper horizon soils and 40 g of lower horizon soils) were repeatedly treated

with HF to concentrate the organic matter and to remove paramagnetic minerals before NMR

analysis (Rumpel et al., 2006; Simpson et al., 2012; Pisani et al., 2015). After HF treatment,

samples were rinsed with deionized water to remove excess salts and then freeze-dried. NMR

spectra were acquired for these treated soils with a 1 s recycle delay. All spectra were

processed using a zero filling factor of 2 and line broadening of 50 Hz. The following

spectral regions were integrated: alkyl (0-50 ppm), O-alkyl (50-110 ppm), aromatic and

phenolic (110-165 ppm), and carboxyl and carbonyl C (165-215 ppm). Chemical shift

assignments were normalized to 100% for relative comparisons of each type of C. Alkyl/O-

alkyl ratios were determined by dividing the areas of the alkyl and O-alkyl regions of the

spectra (Baldock et al., 1992; Simpson et al., 2008).

2.3.6 Statistical analyses

A one way analysis of variance (ANOVA) using the Tukey Honestly Significant

Difference (HSD) test was employed to compare the concentration of SOM components

between the control and the treatment plots (warming, N+P fertilization and warming + N+P

fertilization) of the upper and lower horizons in each of the four replicate blocks (duplicate

40

biomarker analysis). A difference was considered significant at the level of P < 0.05.

Statistical analyses were performed using OriginPro (v8.0932).

2.4 Results

2.4.1 Total carbon (C) and nitrogen (N) content

In the upper horizon soils, the total C content in the control treatments varied from

13.7% to 48.9% while in the warming treatments, it varied from 36.9% to 44.9% (Table 2-1).

In the N+P fertilization and the warming +N+P fertilization treatments, the total C content of

the upper horizon soils ranged from 35.8% to 54.9% and from 11.7% to 52.3%, respectively.

In the lower horizon soils, the total C content in the control treatments ranged from 2.5% to

14.2% while in the warming treatments, it differed from 2.7% to 25.7% (Table 2-1). In the

N+P fertilization treatments, the total C content ranged from 2.6% to 16.9% and in the

warming +N+P fertilization treatments it ranged from 4.0% to 6.1%. In the upper horizon,

the total N content varied from 0.5% to 4.9% among the treatments, while in the lower

horizon, the total N content varied from 0.1% to 1.5% (Table 2-1). The total N content

increased as a result of warming and N+P fertilization in the upper horizon, except in block 1

where N+P fertilization decreased the total N content (Table 2-1). The lower horizon soils

elevated the total N content in response to warming but did not exhibit any overall

differences in the total N content with N+P fertilization. The total N content did not differ

overall with warming +N+P fertilization in either horizon (Table 2-1).

41

Table 2-1: Total carbon and nitrogen content (%) and carbon: nitrogen ratios of the upper

and lower horizon soils of the control, warming, N+P fertilization and warming + N+P

fertilization treatments from all four blocks.

NA = not analyzed (the lower horizon soil sample for the warming +N+P fertilization treatment of block 3 was

unavailable)

Block Treatment Depth (cm) % C % N C:N

1 Control

0-5 34.7 3.1 11.3

5-20 6.3 0.3 20.3

Warming 0-12 44.9 4.0 11.3

12-27 25.7 2.7 11.3

N+P

Fertilization

0-10 48.2 1.8 26.5

18-25 16.9 1.5 11.4

Warming +

N+P

Fertilization

0-12 48.2 2.0 24.0

12-27 6.1 0.5 11.4

2 Control 0-5 33.0 1.0 32.9

12-27 2.5 0.1 20.5

Warming 0-5 37.5 1.1 33.9

5-20 7.0 0.3 22.6

N+P

Fertilization

0-5 54.9 4.9 11.3

15-22 2.6 0.1 19.0

Warming +

N+P

Fertilization

0-5 31.4 1.4 23.3

16-18 4.0 0.4 11.5

3 Control

0-6 13.7 0.6 22.5

6-12 4.9 0.2 25.9

Warming 0-15 41.1 1.9 22.0

15-25 2.7 0.1 20.7

N+P

Fertilization

0-8 47.7 1.6 29.9

15-20 3.4 0.2 22.1

Warming +

N+P

Fertilization

0-10 52.3 2.4 21.6

NA NA NA NA

4 Control 0-12 48.9 0.9 54.3

12-25 14.2 0.4 32.7

Warming 0-10 36.9 1.1 34.2

10-27 25.6 1.3 20.2

N+P

Fertilization

0-10 35.8 1.9 19.2

10-24 2.7 0.1 19.8

Warming +

N+P

Fertilization

0-10 11.7 0.5 22.7

10-20 4.1 0.4 11.5

42

2.4.2 Sources of biomarkers in Arctic soils

The distribution of the solvent extract compound classes (normalized to g of soil) in

the four replicate blocks is shown in Figures 2-1 (upper horizon) and 2-2 (lower horizon).

The majority of compounds identified from solvent extraction included: aliphatic lipids (n-

alkanes, n-alkanols, and n-alkanoic acids), cyclic lipids (steroids and triterpenoids) and

simple carbohydrates (glucose, galactose and mannose). In both horizons of each treatment,

the n-alkanes ranged from C21 to C33, with an odd preference, the n-alkanols ranged from C16

to C32 with an even preference and the n-alkanoic acids ranged from C14 to C32 with an even

preference (Tables A1 to A3). In both horizons, long-chain homologues (≥ C20) of n-alkanols

and n-alkanoic acids were considerably more abundant than their short-chain counterparts

(Tables A2 to A3). Long-chain aliphatic lipids likely originate from epicuticular waxes of

higher plants (Simoneit, 2005; Pisani et al., 2015) while short-chain compounds may

originate from soil microbes including fungi and bacteria (Lichtfouse et al., 1995; Otto and

Simpson, 2005; Pisani et al., 2015). Simple carbohydrates which are likely to originate from

animals, plants and microbes (Simoneit et al., 2004; Pisani et al., 2015), were found to

decrease from the upper horizon to the lower horizon in all treatments (Figures 2-1 and 2-2;

Table A4). Steroids including β-sitosterol, stigmasterol and campesterol were detected (Table

A4) and are commonly found in higher plants (Baker, 1982; Harwood and Russel, 1984;

Bianchi, 1995; Simpson et al., 2008; Pautler et al., 2010). Sitosterone and stigmasta-3,5-dien-

7one, degradation products of β-sitosterol and stigmasterol respectively (Mackenzie et al.,

1982; Simpson et al., 2008; Pautler et al., 2010), were also observed (Table A4). Ergosterol,

a fungal biomarker, was not detected in either horizon of these Arctic soils (Table A4).

Cholesterol was found in both horizons of all treatments (Table A4) but is not a source-

specific biomarker because it can originate from animals, fungi or plants (Simpson et al.,

43

2008). Triterpenoids such as oleanolic acid and ursolic acid, which are characteristic

biomarkers for angiosperms (Tulloch, 1976; Baker, 1982; Bianchi, 1995; Otto and Simpson,

2005; Simpson et al., 2008), were detected as well (Table A4).


soil) of major SOM components from the upper horizon

soil samples after 32 years of warming, N+P fertilization and combined treatments. SOM


mannose), cutin- and suberin-derived lipids and lignin-derived phenols. All values are

reported as mean ± standard error (n = 2). Asterisks denote statistical significance from the

control treatment (P < 0.05).

44


soil) of major SOM components from the lower horizon

soil samples after 32 years of warming, N+P fertilization and combined treatments. SOM


mannose), cutin- and suberin-derived lipids and lignin-derived phenols. All values are

reported as mean ± standard error (n = 2). Asterisks denote statistical significance from the

control treatment (P < 0.05).

From the base hydrolysis extracts of the upper and lower horizon soils, a series of

aliphatic lipids (n-alkanols, n-alkanoic acids, α,ω-alkane dioic acids, ω-hydroxyalkanoic

acids, α-hydroxyalkanoic acids, mid-chain substituted hydroxyl and epoxy acids), benzyls,

phenols (vanillin, acetovanillone, vanillic acid, syringaldehyde, syringic acid, p-coumaric

acid and ferulic acid) and sterols such as β-sitosterol were identified (Table A5). These

compounds are likely attributed to suberin, cutin (Kolattukudy, 1980; Otto and Simpson,

2006b; Pisani et al., 2015), plant surface waxes and ligno-cellulose complexes (Otto and

Simpson, 2006b). The long-chain n-alkanols and n-alkanoic acids identified in the base

hydrolysis extracts may either originate from the plant surface waxes of leaves or from roots

45

(Kolattukudy and Espelie, 1989; Bernards, 2002; Otto and Simpson, 2006b). The C16-C22

α,ω-alkane dioic acids and ω-hydroxyalkanoic acids (Table A5) likely originate from suberin

(Kolattukudy and Espelie, 1989; Bernards, 2002; Otto and Simpson, 2005; 2006b). The mid-

chain-substituted hydroxyl C15 acids and the C16 mono- and dihydroxy acids and diacids may

be derived from cutin (Holloway, 1982; Kolattukudy and Espelie, 1989; Bernards, 2002;

Otto and Simpson, 2006b). β-sitosterol, identified in the upper and lower horizon soils (Table

A5) is likely derived from leaf waxes (Otto and Simpson, 2006b). The benzenes are

nonspecific biomarkers because they are degradation products of proteins and tannins (Goñi

et al., 2000; Otto and Simpson, 2006b). The α-hydroxyalkanoic acids are also nonspecific

biomarkers because they can be found in membranes of animals, fungi and leaf waxes (Otto

and Simpson, 2006b).

CuO oxidation cleaves the aryl ether bonds of the lignin macromolecule and releases

eight major lignin-derived phenols of the vanillyl (vanillin, acetovanillone, vanillic acid),

syringyl (syringaldehyde, acetosyringone, syringic acid) and cinnamyl (p-coumaric acid,

ferulic acid) monomers (Hedges and Mann, 1979; Otto and Simpson, 2006a). In both

horizons, the composition of lignin-derived phenols is dominated by vanillyl and syringyl

monomers (Figures 2-3 and 2-4).

46



cinnamyl monomers) released from CuO oxidation of the upper horizon soil samples after 32

years of warming, N+P fertilization and combined treatments. All values are reported as

mean ± standard error (n = 2). Asterisks denote statistical significance from the control

treatment (P < 0.05).

47



cinnamyl monomers) released from CuO oxidation of the lower horizon soil samples after 32

years of warming, N+P fertilization and combined treatments. All values are reported as

mean ± standard error (n = 2). Asterisks denote statistical significance from the control


2.4.3 SOM composition of warmed soils

In the upper horizon, the concentration of total aliphatic lipids did not vary overall

with soil warming (Figure 2-1; Table A4). The total cyclic lipids concentration increased

with soil warming in the upper horizon but only significantly in blocks 2 and 3 (P < 0.05).

Aliphatic compounds have been found to preferentially degrade compared to cyclic

compounds (Otto and Simpson, 2005). As a result, the aliphatic to cyclic compound

(aliphatic/cyclic) ratio can be used to assess the degradation stage of SOM, where the ratio

decreases with enhanced SOM degradation (Otto and Simpson, 2005; Pisani et al., 2013).

48

Warming decreased the aliphatic/cyclic ratios (P < 0.05) but did not change the

concentrations of simple carbohydrates in the upper horizon soils (Figure 2-1; Table A4). In

the upper horizon of block 4, total aliphatic lipids and total cyclic lipids, along with simple

carbohydrates, all decreased in abundance with warming (Figure 2-1; Table A4; P < 0.05). In

the lower horizon, both total aliphatic lipids and total cyclic lipids were found to accumulate

with warming except in block 3 where the concentrations of both aliphatic and cyclic lipids

decreased (Figure 2-2; Table A4; P < 0.05). Warming did not alter the aliphatic/cyclic ratios

but caused enrichment of simple carbohydrates in the lower horizon soils (Figure 2-2; Table

A4; P < 0.05).

While the concentrations of suberin-derived compounds (∑S) did not change with

warming in the upper horizon, higher concentrations of cutin-derived compounds (∑C) were

observed, but only significantly in block 3 (Figure 2-1; Table A5; P < 0.05). The

suberin/cutin ratio was calculated to determine SOM inputs from roots and leaves (Otto and

Simpson, 2006b). The suberin/cutin ratio was lowered by warming in the upper horizon of

block 3 ( P < 0.05) but did not vary in the upper horizons of the other blocks (Table A5).

Biomarkers derived from suberin or cutin (∑SvC) include C16 and C18 ω-hydroxy acids, α,ω-

alkanedioic, di- and tri-hydroxy C18 acids and the 9,10-epoxy-ω-OH C18 acid (Otto and

Simpson, 2006b). ∑SvC was increased by warming in the upper horizon, but only

significantly in block 3 (Table A5; P < 0.05). The ratio of mid-chain-substituted acids to the

total suberin and cutin acids (∑Mid/∑SC) can be used to evaluate the extent of cutin and

suberin degradation, where decreasing values indicate enhanced degradation (Otto and

Simpson, 2006b). In the upper horizon, the ∑Mid/∑SC ratios did not differ overall with

warming (Table A5). The degradation of cutin-derived compounds can be estimated by

determining the changes in the relative abundances of the ω-C16 and ω-C18 hydroxyalkanoic

49

acids. Expressed as ω-C16 /∑C16 and ω-C18 /∑C18 respectively, these ratios have been shown

to increase with progressive cutin degradation in soil (Otto and Simpson, 2006b). Warming

did not vary the ω-C16/∑C16 and ω-C18/∑C18 cutin degradation ratios in the upper horizon

(Table A5). In the lower horizon, ∑S declined with warming but only significantly in block 3

(P < 0.05), while warming did not elicit any changes in ∑C (Figure 2-2; Table A5). ∑SvC

decreased with warming in the lower horizon, but only significantly in blocks 2 and 3 (P <

0.05). In the lower horizon of block 4, warming elevated both ∑S and ∑SvC (Figure 2-2;

Table A5; P < 0.05). The lower horizons of all the blocks exhibited lowered ω-C16/∑C16

ratios as a result of warming, although only the decrease in block 4 was statistically

significant (Table A5; P < 0.05). No major differences were observed in the ω-C18/∑C18 and

the ∑Mid/∑SC ratios with warming in the lower horizon (Table A5). The suberin/cutin ratios

decreased in all the blocks as a result of warming in the lower horizon although not

significantly (Table A5).

In the upper horizons, total extractable lignin phenol concentrations decreased with

warming but only significantly in block 4 (Figure 2-1; P < 0.05). In the upper horizon of

block 2, these concentrations were elevated by warming (Figure 2-1). Vanillyl monomers

decreased with warming in block 4 (P < 0.05) but did not vary in the other blocks (Figure 2-

3). Decreases in syringyl and cinnamyl monomers due to warming were observed, which

were more pronounced in blocks 3 and 4 (P < 0.05). In the upper horizon of block 2,

warming promoted enrichment of syringyl and cinnamyl monomers but only the cinnamyl

monomers increased significantly (Figure 2-3; P < 0.05). Ratios of lignin-derived phenolic

acids and their corresponding aldehydes (Ad/Al) for both syringyl and vanillyl monomers

can provide information on lignin degradation where increasing Ad/Al ratios suggest

progressive lignin oxidation (Hedges et al., 1988; Otto and Simpson, 2006a). The plots of

50

Ad/Al ratios of the upper and lower horizons soils of each treatment are shown in Figures 2-5

and 2-6 respectively. The Ad/Al ratios did not vary with warming in the upper horizons

except in block 4 where higher Ad/Al ratios were observed (Figure 2-5).


monomers of the upper horizon soils in each block. (Ad/Al)s = syringic acid/syringaldehyde;

(Ad/Al)v = vanillic acid/vanillin. Asterisks denote statistical significance from the control


51


monomers of the lower horizon soils in each block. (Ad/Al)s = syringic acid/syringaldehyde;

(Ad/Al)v = vanillic acid/vanillin.

52

The syringyl/vanillyl (S/V) and cinnamyl/vanillyl (C/V) ratios are used to determine

the specific botanical sources of lignin (Ertel and Hedges, 1984; Prahl et al., 1994; Goñi et

al., 2000; Otto and Simpson, 2006a). The plots of the S/V and C/V ratios of the upper and

lower horizon soils are illustrated in Figures 2-7 and 2-8 respectively.


(C/V) ratios of the upper horizon soils in each block. Asterisks denote statistical significance

from the control treatment (P < 0.05).

53


(C/V) ratios of the lower horizon soils in each block. Asterisks denote statistical significance

from the control treatment (P < 0.05).

The S/V and C/V ratios of the upper horizon soils declined from warming,

particularly in blocks 3 and 4 (Figure 2-7; P < 0.05). In the upper horizon of block 2, the S/V

ratio remained unchanged whereas the C/V ratio increased (Figure 2-7). In the lower horizon

soils, higher extractable lignin phenol concentrations were observed with warming, notably

in blocks 1 and 4 (Figure 2-2; P < 0.05). Warming augmented the abundance of vanillyl

monomers in the lower horizon (P < 0.05), except in block 3 where this increase was not

statistically significant (Figure 2-4). Warming also enriched the lower horizon in syringyl

monomers (P < 0.05), except in block 2 where the increase was not significant (Figure 2-4).

Higher concentrations of cinnamyl monomers were detected in blocks 1 and 4 (P < 0.05) but

54

no changes were observed in blocks 2 and 3 with warming in the lower horizon (Figure 2-4).

The Ad/Al ratios of the lower horizon of each block did not show a uniform response under

warmed conditions (Figure 2-6) which did not present any evidence of accelerated lignin

oxidation. The S/V and C/V ratios of the lower horizon of the warming treatment also

decreased particularly in block 1 (Figure 2-8; P < 0.05). In the lower horizon of block 2, the

S/V ratio was elevated by warming (Figure 2-8).

2.4.4 SOM composition of N+P fertilized soils

With N+P fertilization, the concentration of total aliphatic lipids did not vary in the

upper horizons while the concentration of total cyclic lipids increased in blocks 1 and 2 (P <

0.05) but not significantly in block 3 (Figure 2-1; Table A4). In the upper horizon of block 4,

the abundance of total aliphatic lipids did not change but a pronounced decline in total cyclic

lipids was observed (Figure 2-1; Table A4; P < 0.05). N+P fertilization decreased the

aliphatic/cyclic ratios of the upper horizon soils (P < 0.05) but did not trigger any overall

differences in the concentration of simple carbohydrates (Figure 2-1; Table A4). In the lower

horizon, the abundances of total aliphatic lipids and total cyclic lipids were not altered by

N+P fertilization, which did not lead to any differences in the aliphatic/cyclic ratios (Figure

2-2; Table A4). The concentrations of simple carbohydrates also did not alter with N+P

fertilization in the lower horizon (Figure 2-2; Table A4).

N+P fertilization did not elicit any changes in ∑S and ∑SvC in the upper horizon

soils (Figure 2-1; Table A5). ∑C increased with N+P fertilization in the upper horizon,

notably in block 1 (P < 0.05) but decreased in the upper horizon of block 4 (Figure 2-1; Table

A5). N+P fertilization did not trigger any overall variation in the suberin/cutin ratios of the

upper horizons of the remaining blocks (Table A5). In the lower horizon, N+P fertilization

55

reduced ∑S, ∑C and ∑SvC but these decreases were only statistically significant in blocks 2

and 3 (Figure 2-2; Table A5; P < 0.05). The suberin/cutin ratios were not altered by N+P

fertilization in the lower horizon (Table A5). No overall differences were observed in the ω-

C16/∑C16 and ω-C18/∑C18 cutin degradation ratios and the ∑Mid/∑SC ratios in the upper and

lower horizons of the remaining blocks with N+P fertilization (Table A5).

The total extractable lignin phenol concentrations in the upper horizon did not vary

with N+P fertilization, except for the decrease observed in block 4 (Figure 2-1; P < 0.05).

N+P fertilization augmented the abundance of vanillyl monomers, while the abundance of

syringyl monomers did not vary in the upper horizon (Figure 2-3). Cinnamyl monomers

increased with N+P fertilization in the upper horizon, although only significantly in blocks 2

and 3 (Figure 2-3; P < 0.05). In the upper horizon of block 4, all the lignin monomers were

reduced by N+P fertilization (Figure 2-3; P < 0.05). The Ad/Al ratios did not vary, except in

block 4 where these ratios increased (Figure 2-5). In response to N+P fertilization, the S/V

ratios of the upper horizon soils decreased, except in block 4 where no changes were

observed. The C/V ratios decreased in blocks 1 and 4 but were elevated in blocks 2 and 3 of

the upper horizon (Figure 2-7). In the lower horizon of block 1, the total extractable lignin

phenol concentrations were augmented by N+P fertilization, caused by higher abundances of

vanillyl, syringyl and cinnamyl monomers (Figures 2-2 and 2-4). In the lower horizons of the

other blocks, total extractable lignin declined in response to N+P fertilization, although only

the decreases in blocks 2 and 4 were significant (Figure 2-2; P < 0.05). All lignin monomer

concentrations were reduced in the lower horizon, but only the decreases in vanillyl and

cinnamyl monomers of block 2 were statistically significant (Figure 2-4; P < 0.05). Similar to

our observations in the upper horizon, the Ad/Al ratios did not differ in response to N+P

fertilization in the lower horizon (Figure 2-6). N+P fertilization triggered decreases in the

56

S/V and C/V ratios of the lower horizons of blocks 1 and 4 (Figure 2-8). Higher S/V ratios

were exhibited in blocks 2 and 3 while the C/V ratios decreased in block 2 and increased in

block 3 with N+P fertilization in the lower horizon (Figure 2-8).

2.4.5 SOM composition of warmed + N+P fertilized soils

In the upper horizons, warming +N+P fertilization elevated the concentrations of total

aliphatic lipids significantly in blocks 1 and 3 (P < 0.05) but not significantly in block 2

(Figure 2-1; Table A4). Enrichment in total cyclic lipids was also detected (Figure 2-1; Table

A4; P < 0.05). Although the concentrations of aliphatic and cyclic lipids both increased, the

aliphatic/cyclic ratios decreased with warming +N+P fertilization in the upper horizon (Table

A4; P < 0.05). In the upper horizon of block 4, total aliphatic lipids and total cyclic lipids

were lowered by warming +N+P fertilization (P < 0.05), causing no variation in the

aliphatic/cyclic ratio (Figure 2-1; Table A4). No major differences were detected in the

concentration of simple carbohydrates from the upper horizon of the warming +N+P

fertilization treatment (Figure 2-1; Table A4). In the lower horizon, warming +N+P

fertilization led to enrichment in total aliphatic lipids which was significant in blocks 1 and 2

but not in block 4 (Figure 2-2; Table A4; P < 0.05). Warming +N+P fertilization did not alter

the abundances of total cyclic lipids and simple carbohydrates in the lower horizons (Figure

2-2; Table A4). The aliphatic/cyclic ratios notably increased in the lower horizons of blocks

2 and 4 (P < 0.05) but not significantly in the lower horizon of block 1 in response to

warming +N+P fertilization (Table A4).

∑S decreased as a result of warming +N+P fertilization in the upper horizons,

particularly in block 4 (P < 0.05) but increased in block 3 (Figure 2-1; Table A5). Warming

+N+P fertilization caused enrichment in ∑C in the upper horizon, except in block 4 where

57

∑C decreased (Figure 2-1; Table A5). ∑SvC in the upper horizon soils did not vary across all

the blocks (Table A5). Warming +N+P fertilization augmented the ω-C18/∑C18 and the

∑Mid/∑SC ratios of the upper horizon soils but only significantly in block 2 (Table A5; P <

0.05). In the upper horizon of block 1, these ratios did not change (Table A5). No major

differences were observed with the ω-C16/∑C16 ratios of the upper horizon (Table A5). In the

lower horizon, both ∑S and ∑SvC were reduced by warming +N+P fertilization notably in

block 2 (Figure 2-2; Table A5; P < 0.05). ∑C decreased in block 2 (P < 0.05) but did not alter

in the lower horizons of the other blocks (Figure 2-2; Table A5). The ω-C16/∑C16, ω-

C18/∑C18 and ∑Mid/∑SC ratios did not differ with warming +N+P fertilization in the lower

horizon (Table A5). The suberin/cutin ratios of both horizons declined as a result of warming

+N+P fertilization (Table A5).

Warming +N+P fertilization reduced the abundances of total extractable lignin

phenols in the upper horizon but only the decrease in block 4 was significant (Figure 2-1; P <

0.05). Vanillyl monomers did not vary overall while syringyl and cinnamyl monomers in the

upper horizon declined in response to warming +N+P fertilization (Figure 2-3). The upper

horizon of block 2 elicited higher concentrations of total extractable lignin phenols (Figure 2-

1; P < 0.05), caused by enrichment in syringyl and cinnamyl monomers but only the increase

in syringyl monomers was significant (Figure 2-3; P < 0.05). In the upper horizon, warming

+N+P fertilization did not vary the Ad/Al ratios of blocks 1 and 3 (Figure 2-5) but lowered

that of block 2 (Figure 2-5; P < 0.05). On the contrary, warming +N+P fertilization enhanced

lignin oxidation in the upper horizon of block 4, as demonstrated by the elevated Ad/Al ratios

(Figure 2-5). The decline in S/V and C/V ratios with warming +N+P fertilization in the upper

horizon soils were only significant in block 3 (Figure 2-7; P < 0.05). The upper horizon of

block 4 showed higher S/V and C/V ratios in response to warming +N+P fertilization (Figure

58

2-7). In the lower horizon soils, warming +N+P fertilization did not promote any overall

differences in the total extractable lignin phenol concentrations (Figure 2-2) or in the

abundances of each lignin monomer (Figure 2-4). The Ad/Al ratios did not vary (Figure 2-6)

but the S/V and C/V ratios declined in response to warming +N+P fertilization in the lower

horizon (Figure 2-8).


C NMR of Arctic soils

Solid-state 13

C NMR can provide basic structural information of SOM components

(Simpson et al., 2008; Simpson and Simpson, 2012). The results of the solid-state 13

C NMR

analysis of the block 1 upper and lower horizons of the four treatments are summarized in

Table 2-2 and their corresponding NMR spectra are illustrated in Figure 2-9. The four main

spectral regions identified and their corresponding chemical shifts are as follows: alkyl C (0-

50 ppm); O-alkyl C (50-110 ppm); aromatic and phenolic C (110-165 ppm) and carboxylic

and carbonyl C (165-215 ppm; Baldock et al., 1992; Simpson et al., 2008). The solid-state

13C NMR spectra revealed that both the upper and lower horizon soils are dominated by alkyl

and O-alkyl C (Figure 2-9; Table 2-2). Alkyl C originates from cutin, suberin, aliphatic side-

chains and lipids whereas O-alkyl C originates from the substituted aliphatic constituents of

carbohydrates, peptides and methoxyl C found in lignin (Baldock et al., 1992; Simpson et al.,

2008). Aromatic and phenolic C along with carboxylic and carbonyl C exhibited relatively

less intense signals compared to the alkyl and O-alkyl C regions in the 13

C NMR spectra of

both horizons (Figure 2-9; Table 2-2). Aromatic and phenolic signals arise from lignin and

aromatic amino acids while carboxylic and carbonyl signals are from fatty acids and peptides

(Baldock et al., 1992; Simpson et al., 2008).

59

Table 2-2: Solid-state 13

C CPMAS-NMR integration results with relative contribution (%) of the four main carbon structures and calculated

alkyl/O-alkyl ratios for the Control, Warming, N+P Fertilization and Warming + N+P Fertilization treatments of the upper and lower horizon

soils of Block 1.

Horizon Treatment Alkyl C

(0-50 ppm)

O-Alkyl C

(50-110 ppm)

Aromatic and Phenolic C

(110-165 ppm)

Carboxylic and Carbonyl C

(165-215 ppm)

Alkyl/O-Alkyl

Upper Control 33 35 20 12 0.94

Warming 31 40 18 11 0.78

N+P Fertilization 34 36 20 10 0.94

Warming + N+P

Fertilization 25 36 28 11 0.69

Lower Control 46 25 19 10 1.84

Warming 30 37 21 12 0.81

N+P Fertilization 37 30 21 12 1.23

Warming + N+P

Fertilization 31 39 18 12 0.79

60

Figure 2-9: Solid-state 13

C CPMAS-NMR spectra of the Block 1 upper (a) and lower (b) horizon soils of the Control, Warming, N+P

Fertilization and Warming + N+P Fertilization treatments with the four major spectral regions: alkyl (0-50 ppm), O-alkyl (50-110 ppm),

aromatic and phenolic (110-165 ppm) and carboxylic and carbonyl carbon (165- 215 ppm).

(A) (B)

61

The alkyl C signal was neither altered by warming nor by N+P fertilization, but was

lowered by warming +N+P fertilization in the upper horizon soils (Figure 2-9; Table 2-2).

Warming elevated the O-alkyl C signal, but no other changes were observed with the other

treatments in the upper horizon. In the lower horizon, warming, N+P fertilization and their

combined treatment facilitated reductions in the alkyl C signals and enhancements in the O-

alkyl C signals relative to the control (Figure 2-9b; Table 2-2). In the upper horizon, the

aromatic and phenolic C signal was slightly reduced by warming, elevated by warming

+N+P fertilization, but remained unchanged with N+P fertilization (Figure 2-9a; Table 2-2).

In the lower horizon, the aromatic and phenolic C signal slightly increased in response to

warming and N+P fertilization, but did not alter as a result of the combined treatment (Figure

2-9b; Table 2-2). The relative percent changes of the carboxylic and carbonyl C regions in

the solid-state 13

C NMR spectra of both horizons did not differ with each treatment (Figure

2-9; Table 2-2). Since O-alkyl compounds are more easily degraded than alkyl compounds,

the ratio of alkyl/O-alkyl C offers information on the relative stage of SOM degradation. This

ratio increases with progressive degradation (Baldock et al., 1992; Simpson et al., 2008). In

both horizons, the alkyl/O-alkyl ratios were lowered by warming and warming + N+P

fertilization but did not alter in response to N+P fertilization (Table 2-2).

2.5 Discussion

2.5.1 Spatial heterogeneity and vertical mixing

The variation in total C content may be due to the differences in depths of the upper

and lower horizons of each replicate block within each treatment (Table 2-1). Although the

collection of distinct layers was attempted, the total C content still differed among the blocks

in both horizons. Hence, samples from the replicate blocks belonging to the same treatment

62

were not composited and were analyzed separately. The biomarker data showed less

variability in response to each treatment, more so when it was normalized to the mass of soil

(Figures 2-1 to 2-4; Tables A1 to A5) than when it was normalized to the C content. The

differences in total C content among the upper and lower horizons of the replicate blocks

within each treatment (Table 2-1) may be attributable to spatial heterogeneity across the

landscape (Burke et al., 1999; Hook and Burke, 2000; Schöning et al., 2006; Spielvogel et

al., 2016). The results from our preliminary solid-state 13

C NMR experiment showed notable

differences in the NMR resonance intensities of SOM components between the control

samples from each block (Table A6; Figure A1), which offer further evidence that each block

may have responded differently to each treatment due to spatial variability. Landscape

heterogeneity will alter soil moisture content, nutrient availability, soil development and

plant growth (Burke et al., 1999; Schöning et al., 2006; Mishra and Riley, 2015). These

factors will be discussed more in depth in section 2.5.5. A recent study employed biomarker

methods and NMR spectroscopy along with multivariate geostatistical approaches to

examine the variation in the distribution of SOM compounds in an ecosystem setting

(Spielvogel et al., 2016). Geostatistical analyses of biomarker and solid-state 13

C NMR

spectroscopy data showed small scale heterogeneity was correlated to topography, where the

distribution of SOM was associated with the topographical pattern of the site. For example,

higher amounts of O-alkyl C components including simple sugars such as glucose, galactose

and mannose were found in the central depression of the field site compared to locations with

steeper terrain (Spielvogel et al., 2016). The authors also observed that the distribution of

lignin and aromatic C in the soil corresponded to the location of specific vegetative inputs

surrounding the sampling sites. Although these observations could be ecosystem-specific,

Spielvogel et al. (2016) demonstrated that the distribution of SOM components likely reflects

63

the spatial heterogeneity of the field site and biomarker and NMR analyses are sensitive

enough to detect localized differences within the landscape.

In addition to spatial heterogeneity among the blocks, the organic and mineral

horizon soils in some blocks from Toolik Lake could be not distinguished based on C content

(Table 2-1). For example, the total C contents of the lower horizons of the warming

treatments in blocks 1 and 4 were greater than 17% (Table 2-1) and therefore cannot be

considered true mineral horizons (Gregorich et al., 2001). In the upper horizon of the

warming +N+P fertilization treatment of block 4, the total C content was less than 17%

(Table 2-1) and therefore cannot be considered a true organic horizon (Gregorich et al.,

2001). The lack of true distinction between the organic and mineral horizons may likely be

caused by cryoturbation, where vertical mixing occurs between surface organic matter and

mineral soils (Bockheim and Tarnocai, 1998). Surface organic matter is churned and moves

downward to the permafrost table surface (Tarnocai and Smith, 1992; Ping et al., 1998;

2008) causing soils in deeper horizons to become exposed at the surface (Bockheim and

Tarnocai, 1998). This vertical mixing likely redistributed C between the soil horizons

(Michaelson et al., 1996) which may explain why the C content was as low as 11.7% in the

upper horizon soils and as high as 25.7% in the lower horizon soils (Table 2-1).

Cryoturbation is triggered by soil freeze-thaw cycles and soil hydrothermal gradients that

result in differential frost heaving (Peterson, 2003; Ping et al., 2008). Frost heaving occurs

when a growing ice layer within the soil horizon thrusts soil upward and fractures overlying

rock (Williams and Smith, 1989; Bockheim and Tarnocai, 1998). The variation in C content

among the blocks from each treatment (Table 2-1) may also be caused by differences in thaw

depths among the blocks. The thaw depth of the active layer may differ depending on the

location and year, where maximum thaw depths range from 0.3 m to more than 1.0 m (Sturm

64

et al., 2005). The thaw depth of the active layer plays an important role in determining the

total amount of C that cycles within the soil. Differences in thaw depths is a reflection of

complex interactions among topography, soil moisture and thermal properties of the upper

and lower horizon soils (Shaver et al., 2014).

2.5.2 Labile SOM components

A greater abundance of long-chain aliphatic lipids over short-chain aliphatic lipids

was observed in these Arctic soils (Tables A1 to A3), which suggests that microbial inputs

are less prevalent than those from vascular plants (Otto and Simpson, 2005). The preferential

accumulation of cyclic compounds in the upper horizon of the warming treatment (Table A4)

is consistent with previous observations where cyclic lipids have been found to preferentially

accumulate over aliphatic lipids, likely because cyclic lipids possess more complex structures

which are less prone to degradation (Otto and Simpson, 2005). Warming did not alter the

abundance of simple carbohydrates in the upper horizon but elicited enrichment in the lower

horizon (Table A4; Figures 2-1 and 2-2). The elevated concentrations of simple

carbohydrates in the lower horizon may have been caused by the breakdown of cellulose

(Amelung et al., 2009), the most abundant plant biopolymer (Kögel-Knabner, 2002). The

enhancement in the O-alkyl C signal in the solid-state 13

C NMR spectrum of the lower

horizon with warming (Figure 2-9b; Table 2-2) is most likely to arise from cellulose (Hatcher

et al., 1983; Salloum et al., 2002). In the upper horizon, the greater intensity of the O-alkyl C

signal with warming (Figure 2-9a; Table 2-2) coupled with no changes in the concentration

of simple carbohydrates (Figure 2-2; Table A4) further indicate that abundant amounts of

cellulose are present in these Arctic soils.

65

Similar to our observations from the warming treatment, N+P fertilization promoted

the preferential accumulation of cyclic compounds in the upper horizon (Table A4), which

may be attributed to the complex structures of cyclic compounds that cause them to be less

susceptible to degradation (Otto and Simpson, 2005). The abundance of simple

carbohydrates was not altered by N+P fertilization in either horizon (Figures 2-1 and 2-2;

Table A4), which corresponded to no observed difference in the O-alkyl C signal from the

solid-state 13

C NMR spectrum of the upper horizon (Figure 2-9a; Table 2-2). However, in the

lower horizon, N+P fertilization did not alter the concentration of simple carbohydrates, yet

the O-alkyl C signal increased in intensity (Figure 2-9b; Table 2-2), which suggests these

soils were enriched in cellulose (Hatcher et al., 1983; Salloum et al., 2002).

Warming +N+P fertilization promoted the preferential accumulation of cyclic

compounds in the upper horizon and the preferential accumulation of aliphatic lipids in the

lower horizon (Figures 2-1 and 2-2; Table A4). The mixing between the upper and lower

horizon soils caused by cryoturbation (Ping et al., 2008) may have facilitated the exchange of

aliphatic and cyclic lipids between the horizons. Aliphatic lipids may also have accumulated

in the lower horizon by the translocation of dissolved organic C from the upper horizon

(Currie et al., 1996; Pisani et al., 2015). The concentration of simple carbohydrates was not

found to change in the upper horizon from warming +N+P fertilization (Figure 2-2), which

corresponds to no observed change in the O-alkyl C signal intensity in the solid-state 13

C

NMR spectrum (Figure 2-9a; Table 2-2). In the lower horizon, warming +N+P fertilization

enhanced the O-alkyl C signal (Figure 2-9b; Table 2-2) but did not change the concentration

of simple carbohydrates, which suggests large amounts of cellulose are contained in these

Arctic soils (Hatcher et al., 1983; Salloum et al., 2002).

66

2.5.3 Recalcitrant SOM components

Warming elevated the concentrations of cutin-derived components in the upper

horizon but not significantly (Figure 2-1; Table A5). However, the abundance of long-chain

n-alkanes (Table A1) derived from epicuticular waxes of plants (Simoneit, 2005; Pisani et al.,

2015) did not vary, which indicates there was a slight accumulation of aboveground inputs in

the upper horizon under elevated temperatures. The abundance of suberin-derived

compounds did not change in response to warming in the upper horizon (Figure 2-1; Table

A5). In the lower horizon, warming enhanced the abundance of long-chain n-alkanes derived

from epicuticular waxes (Table A1) but did not alter the amount of cutin-derived compounds

(Figure 2-2; Table A5). The decreases in suberin-derived compounds in the lower horizon

with warming were not statistically significant (Figure 2-2; Table A5), which signifies

elevated temperatures may not have triggered substantial reductions in belowground inputs.

These observations collectively indicate that warming may not have markedly altered the

amount of cutin- and suberin-derived inputs in the upper and lower horizons.

N+P fertilization enriched the upper horizon in cutin-derived compounds (Figure 2-1;

Table A5) but lowered the abundance of long-chain n-alkanes derived from epicuticular

waxes (Table A1), which suggests increased nutrient availability may not have promoted the

accumulation of aboveground inputs in the upper horizon. The amount of suberin-derived

compounds did not alter in response to N+P fertilization (Figure 2-1; Table A5). The lack of

changes in aboveground and belowground inputs in the upper horizon correspond to no

differences in the alkyl C signals of the solid-state 13

C NMR spectrum of the upper horizon

(Figure 2-9a; Table 2-2). In the lower horizon, N+P fertilization reduced the abundance of

suberin- and cutin-derived biomarkers (Figure 2-2; Table A5), which may have collectively

contributed to the decreased signal of the alkyl C region (Figure 2-9b; Table 2-2). However,

67

N+P fertilization did not alter the abundance of long-chain n-alkanes from epicuticular waxes

(Table A1). This suggests that despite the slight reductions in suberin- and cutin-derived

compounds, the amount of aboveground and belowground inputs may not have been altered

by N+P fertilization in the lower horizon.

Warming +N+P fertilization increased the amount of cutin-derived compounds in the

upper horizon (Figure 2-1; Table A5) yet the abundance of long-chain n-alkanes from

epicuticular waxes did not change (Table A1), which indicates there was a minor

accumulation of aboveground inputs in the upper horizon. Suberin-derived inputs decreased

with warming +N+P fertilization in the upper horizon, but not significantly (Figure 2-1;

Table A5). In the lower horizon, higher abundances of long-chain n-alkanes were observed

(Table A1) but the concentration of the cutin-derived components did not change (Figure 2-

2; Table A5). This suggests elevated temperatures and nutrient availability did not promote

enrichment of aboveground inputs in the lower horizon. Suberin-derived components were

decreased by warming +N+P fertilization in the lower horizon (Figure 2-2; Table A5), but

this was not statistically significant. This indicates belowground inputs in the lower horizon

may not have been substantially altered by warming +N+P fertilization.

The cutin degradation ratios did not present any evidence of enhanced cutin

degradation in either horizon with warming, N+P fertilization or warming +N+P fertilization

(Table A5). The suppression of cutin degradation may be due to the chemical recalcitrance of

this alkyl structure (Riederer et al., 1993; Lorenz et al., 2007). Furthermore, the lack of

microbial inputs, as indicated by the low abundances of short-chain aliphatic lipids (Tables

A2 and A3), suggests low microbial activity. This may also explain why cutin-derived

components did not degrade in response to these treatments (Table A5).

68

2.5.4 Lignin-derived components

The extractability of lignin-derived phenols was considerably lower compared to the

extractability of labile and recalcitrant SOM components observed from the biomarker data

(Figures 2-1 to 2-4). The relatively lower concentrations of extractable lignin are further

supported by the solid-state 13

C NMR spectra of the upper and lower horizons of all the

treatments (Figure 2-9; Table 2-2). The aromatic and phenolic C signals, derived from lignin

as well as the aromatic components of suberin and amino acids (Baldock et al., 1992;

Simpson et al., 2008), were less intense compared to the O-alkyl and alkyl C signals, which

arise from labile and recalcitrant SOM components respectively (Baldock et al., 1992;

Simpson et al., 2008).

Although the warming treatment exhibited decreased lignin phenol concentrations in

the upper horizon and increased concentrations in the lower horizon, these were not

statistically significant changes (Figures 2-1 and 2-2). This is reflected by a reduction and an

enhancement in the aromatic and phenolic C signals in the solid-state 13

C NMR spectra of the

upper and lower horizons respectively in response to warming (Figure 2-9; Table 2-2).

Lignin was likely introduced into the lower horizon via the transport of dissolved organic C

down the soil profile (Currie et al., 1996; Pisani et al., 2015). N+P fertilization did not cause

any changes in the abundance of lignin phenols in the upper horizon (Figure 2-1), which is

consistent with no alteration in the aromatic and phenolic C signal of the solid-state 13

C

NMR spectra in the upper horizon (Figure 2-9a; Table 2-2). Despite decreased lignin phenol

concentrations in the lower horizon as a result of N+P fertilization (Figure 2-2), the solid-

state 13

C NMR spectrum illustrated a slight enhancement in the aromatic and phenolic C

signal of the lower horizon (Figure 2-9b; Table 2-2). A similar observation was made for the

upper horizon of the warming +N+P fertilization treatment where lignin-derived phenols

69

declined (Figure 2-2) but the aromatic and phenolic C signal intensified (Figure 2-9a; Table

2-2). The elevated intensity in the aromatic and phenolic C signal likely did not arise from

suberin because the abundance of suberin-derived components decreased with N+P

fertilization in the lower horizon and with warming +N+P fertilization in the upper horizon

(Figure 2-2; Table A5). The alkyl C signals of their corresponding solid-state 13

C NMR

spectra also decreased (Figure 2-9b; Table 2-2). Warming +N+P fertilization did not alter

lignin phenol concentrations in the lower horizon (Figure 2-2) which corresponds to the lack

of difference in the aromatic and phenolic C signal in the solid-state 13

C NMR spectrum

(Figure 2-9b; Table 2-2).

The Ad/Al ratios also showed no evidence of accelerated lignin oxidation with any of

the treatments, except in the upper horizon of block 4 (Figures 2-5 and 2-6). This will be

discussed in detail below. Fungi are the primary decomposers involved in lignin

depolymerisation and degradation (Hedges et al., 1988). The absence of ergosterol, a fungal

biomarker, from the solvent extracts of all the treatments from both horizons (Table A4),

which indicates low fungal activity (Otto and Simpson, 2005), may support our observation

of suppressed lignin oxidation. Low fungal activity may be attributed to low temperatures in

Arctic environments (Requejo et al., 1991; Pautler et al., 2010). In the upper horizon of block

4, the Ad/Al ratios progressively increased with each treatment (Figure 2-5), which signifies

lignin in the N+P fertilization treatment was the most oxidized while lignin in the warming

+N+P fertilization treatment was the least oxidized. Block 4 behaved differently than the

other blocks, which is evidenced by the distinct decrease in C content with warming only

observed in block 4 (Table 2-1). This enhancement in lignin oxidation in block 4 is not likely

due to fungal activity because ergosterol was not detected in the solvent extracts of the upper

horizon of block 4 (Table A4). Abiotic mechanisms such as photooxidation from the

70

exposure of sunlight may have facilitated lignin oxidation in block 4 (Opsahl and Benner,

1998; Bertilsson et al., 1999; Pautler et al., 2010) but it is unclear why block 4 showed more

lignin oxidation than the other blocks. Localized environmental controls, such as moisture

content, may have also played a role in lignin oxidation.

The CuO oxidation extracts of these Arctic soils were mostly dominated by vanillyl

and syringyl monomers (Figures 2-3 and 2-4), which suggests that the SOM likely originated

from angiosperm sources (Hedges and Mann, 1979). These angiosperm sources are likely to

be the overlying vegetation predominantly comprised of woody shrubs such as B. nana and

tussock-forming sedges such as E. vaginatum (Ping et al., 1998). The S/V and C/V ratios

declined in the upper and lower horizons with all the treatments (Figures 2-7 and 2-8), which

indicates that the lignin may have been derived from woody angiosperms (Hedges and Mann,

1979). Warming, N+P fertilization and their combined treatment may have promoted the

growth of woody angiosperms such as B. nana (Michaelson et al., 1996), and the

decomposed litter from these woody angiosperms may have contributed lignin inputs into the

soil. The S/V and C/V ratios of both horizons of the warming treatment from block 2

(Figures 2-7 and 2-8) signified that lignin may have been derived from nonwoody

angiosperms such as E. vaginatum (Hedges and Mann, 1979). These nonwoody angiosperms

were likely more abundant before warming conditions were established (Hobbie, 1996). The

extent of vertical mixing between horizons (Ping et al., 1998) likely varied across the

landscape. In block 2, vertical mixing may have been more pronounced, causing newer lignin

from woody angiosperms to be embedded in lower horizons and older lignin from nonwoody

angiosperms to be exposed in the upper horizons.

71

2.5.5 Implications on warming and N+P fertilizer addition on SOM degradation

It is important to acknowledge that spatial heterogeneity among the blocks was likely

an important factor behind the variable responses from each treatment. Spatial heterogeneity

in SOM can be ascribed to the topography of the site (Burke et al., 1999; Hook and Burke,

2000; Schöning et al., 2006; Spielvogel et al., 2016). SOM composition may be largely

reflective of topographical controls such as water distribution in soils, vegetative inputs and

soil horizon development (Hook and Burke, 2000). For example, the topography of the

landscape may alter soil moisture content, such that areas at lower elevation tend to promote

the accumulation of water caused by poor drainage (Mishra and Riley, 2015). Areas at higher

elevation will have drier conditions from better drainage (Mishra and Riley, 2015). Plant

inputs into the soil may also contribute to spatial variation in SOM composition since

different types of vegetation produce litter of varying quality (Kristensen et al., 2015). Since

water distribution and plant inputs may regulate soil horizon development (Mishra and Riley,

2015), these factors may have altered the amount of C stocks among the blocks, thus

contributing to the spatial heterogeneity of the site. Furthermore, cryoturbation caused by

repeated freezing and thawing processes mixes soil material, which results in broken and

discontinuous horizons (Ping et al., 1998). Cryoturbation may have promoted vertical mixing

in some blocks more than others, particularly in the warming treatments of blocks 1 and 4

and the warming +N+P fertilization treatment in block 4 (Table 2-1). These cryogenic

processes can also lead to the formation of patterned ground features such as ice wedge

polygons along with sorted and non-sorted polygons (Washburn, 1980; French, 1996; Ping et

al., 1998). As a result, these periglacial features in the landscape can ultimately vary

vegetation growth (Billings et al., 1982), drainage and the depth of the active layer, leading

to distinct differences in soil characteristics within small areas (Tarnocai, 1994; Ping et al.,

72

1998). The biomarker methods and solid-state 13

C NMR techniques employed in this study

were sensitive enough to detect differences in the alteration patterns of SOM composition

among the blocks.

After 32 years of warming, N+P fertilization and warming +N+P fertilization,

although cutin-derived compounds in the upper horizon soils slightly increased (Figure 2-1;

Table A5), the abundance of long-chain n-alkanes derived from epicuticular waxes

(Simoneit, 2005) did not vary (Table A1), which did not indicate a major enrichment of

aboveground inputs in the upper horizon. In the lower horizon soils, we detected a slight

reduction in suberin-derived compounds with each treatment (Figure 2-1; Table A5). Since

this was not a statistically significant change, this observation suggests belowground inputs

in the lower horizon were not altered by each treatment.

The cutin degradation ratios revealed that warming, N+P fertilization and warming

+N+P fertilization did not enhance cutin degradation in these Arctic soils. This is consistent

with literature findings where cuticle-derived compounds are believed to be chemically

recalcitrant structures (Riederer et al., 1993; Lorenz et al., 2007). Low temperatures and

seasonal ice cover in Arctic environments (Requejo et al., 1991; Pautler et al., 2010) may

have further contributed to the accumulation of these recalcitrant compounds. In the upper

horizon, the alkyl C signal from the solid-state 13

C NMR spectra (Figure 2-9a; Table 2-2) did

not vary as a result of warming and N+P fertilization, which further supports our hypothesis

that cutin degradation was not likely promoted. Despite the decrease in the alkyl C signal in

the solid-state 13

C NMR spectrum of the upper horizon of the warming +N+P fertilization

treatment (Figure 2-9a; Table 2-2), the cutin degradation ratios (Table A5) along with an

increase in cutin-derived compounds (Figure 2-1; Table A5) did not offer evidence for

enhanced cutin degradation. The decline in the alkyl C signals of the solid-state 13

C NMR

73

spectra of the lower horizon soils exhibited by warming and warming +N+P fertilization

(Figure 2-9b; Table 2-2) likely did not arise from accelerated cutin degradation because no

differences were detected in the cutin degradation ratios (Table A5) and cutin-derived inputs

(Figure 2-2; Table A5) under these treatments. N+P fertilization elicited decreases in cutin-

and suberin-derived compounds in the lower horizon (Figure 2-2; Table A5) which

corresponded to a reduction in the alkyl C signal of the solid-state 13

C NMR spectrum

(Figure 2-9b; Table 2-2), although the cutin degradation ratios did not indicate any

enhancement in cutin degradation. In addition, the Ad/Al ratios did not vary overall in

response to warming, N+P fertilization and warming +N+P fertilization (Figures 2-5 and 2-6)

which did not suggest any acceleration in lignin oxidation with each treatment, may be

attributable to low fungal activity. This is further supported by the lack of enhancement in the

aromatic and phenolic C signal of the solid-state 13

C NMR spectra with each treatment

(Figure 2-9; Table 2-2). The relatively low abundances of lignin-derived phenols from the

CuO oxidation extracts (Figures 2-1 to 2-4) were consistent with the less intense aromatic

and phenolic C signals from the solid-state 13

C NMR spectra (Figure 2-9; Table 2-2), which

collectively provide little evidence of lignin cycling in these Arctic soils.

2.6 Conclusions

Molecular-level techniques including biomarker analyses and solid-state 13

C NMR

spectroscopy were employed to investigate the extent of SOM degradation in Arctic soils

after 32 years of warming, N+P fertilization and warming +N+P fertilization. The blocks

showed variable responses to each treatment, likely caused by spatial heterogeneity. These

results suggest that in a complex landscape impacted by cryoturbation, alterations in SOM

composition may vary because of differences in water distribution, plant inputs and soil

horizon development. Due to the spatial variability at this site in Toolik Lake, Alaska,

74

additional long-term field warming and fertilization studies should be conducted at multiple

locations within this region to confirm whether our observations are unique to this site.

75

2.7 References

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biomes. Journal of Ecology 94, 712-724.

Amelung, W., Brodowski, S., Sandhage-Hofmann, A., Bol, R., 2009. Combining biomarker

with stable isotope analyses for assessing the transformation and turnover of soil organic

matter. Advances in Agronomy 100, 155-250.

Baker, E.A., 1982. The Plant Cuticle. Linnean Society Symposium Series 10. In: Cutler,

D.F., Alvin, K.L., Price, C.E. (Eds.), Chemistry and Morphology of Plant Epicuticular

Waxes. Academic Press, London, UK.

Baldock, J.A., Oades, J.M., Waters, A.G., Peng, X., Vassallo, A.M., Wilson, M.A., 1992.

Aspects of the chemical structure of soil organic materials as revealed by solid-state 13

C

NMR spectroscopy. Biogeochemistry 16, 1-42.

Bernards, M.A., 2002. Demystifying suberin. Canadian Journal of Botany 80, 227-240.

Bertilsson, S., Stepanauskas, R., Cuadros-Hansson, R., Granéli, W., Wikner, J., Tranvik, L.,

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Chapter 3: Evaluation of clay mineral and suberin and cutin

protection of lignin in temperate soils from surface horizons

3.1 Abstract

Soil organic matter (SOM) contains around 1580 Gt of carbon (C) and is one of the

largest global reservoirs of C but the fundamental factors governing SOM stabilization

remain unclear. Recent studies have demonstrated that lignin, a major SOM component, may

be protected from degradation through organo-mineral interactions but such associations are

rarely examined at the molecular-level. Previous research also indicated that suberin and

cutin, which are major sources of aliphatic constituents in soil, may interact with lignin

through hydrophobic interactions but it is unclear if such interactions can protect lignin from

degradation in soil. In this study, we employed molecular-level techniques to examine the

preservation patterns of lignin in soil. Hydrofluoric acid (HF) demineralization was used to

evaluate the extent of mineral protection of lignin-derived phenols. Base hydrolysis (BH)

was used to isolate suberin and cutin monomers from agricultural, forest and 2 grassland

soils with varying mineralogy before and after HF demineralization. Copper (II) oxide

oxidation (CuO) was performed before and after HF demineralization and BH to release

lignin-derived phenols, which were analyzed by gas chromatography-mass spectrometry. The

percentage of mineral-protected lignin among the soils ranged from 3.1 ± 0.2% to 95.9 ±

0.2%, where greater protection was observed in the grassland soils which contain higher

montmorillonite clay content likely rendering more surface area for sorption. Extractable

lignin phenol concentrations increased in all four soil samples after HF treatment suggesting

that mineral protection is likely an important protection mechanism of lignin. In the southern

grassland, agricultural and forest soils, suberin and cutin protection of lignin was not

83

observed until after HF demineralization. Suberin and cutin protection was not observed in

the northern grassland soil, indicating that suberin and cutin do not protect lignin in this soil.

Our results suggest that lignin is protected from extraction to a greater extent via organo-

mineral interactions compared to interactions with suberin and cutin. Interactions between

suberin and cutin with lignin in soil should be further examined to understand their

correlation with organo-mineral associations and how they collectively contribute to organic

matter stabilization processes.

3.2 Introduction

Soil organic matter (SOM) plays a key role in carbon (C) sequestration (Batjes, 1996)

but the fundamental factors governing SOM stabilization remain unclear. A proposed

mechanism for SOM stabilization is through association with clay mineral surfaces, which

affords protection from microbial degradation (Baldock and Skjemstad, 2000; Eusterhues et

al., 2003; Kaiser and Guggenberger, 2003; Mikutta et al., 2006). The dissolution of minerals

by hydrofluoric acid (HF) treatment is believed to release mineral-bound organic matter

(OM) which can provide information about organo-mineral associations in soil (Schmidt et

al., 1997; Eusterhues et al., 2003; Mikutta et al., 2006; Rumpel et al., 2006). Interactions

between SOM and clay minerals vary depending on solution properties (pH, ionic strength

and presence of cations), mineralogy and the composition of the OM sorbate (Asselman and

Garnier, 2000; Chi and Amy, 2004; Feng et al., 2005; Mikutta et al., 2007; Ghosh et al.,

2009; Clemente and Simpson, 2013). It has been observed that polymethylene structures

preferentially sorb to kaolinite and montmorillonite surfaces (Feng et al., 2005; Simpson et

al., 2006; Ghosh et al., 2009) while carboxyl groups preferentially sorb to goethite surfaces

(Ghosh et al., 2009). In addition, evidence also suggests that carboxyl and aromatic

84

components can form strong complexes with iron (Fe) and aluminum (Al) oxides through

ligand exchange (Kaiser and Guggenberger, 2000).

Lignin, a major component of SOM, is the second most abundant biopolymer after

cellulose and possesses an ether-linked phenylpropanoid structure (Filley et al., 2002).

Research has suggested that lignin may be preserved through interactions with clay mineral

surfaces (Bahri et al., 2006; Heim and Schmidt, 2007; Clemente et al., 2011; Clemente and

Simpson, 2013; Hernes et al., 2013). Model sorption experiments have found that greater

concentrations of aromatic structures sorb to montmorillonite surfaces compared to kaolinite

surfaces (Feng et al., 2005; Ghosh et al., 2009; Clemente and Simpson, 2013; Genest et al.,

2014). Sorptive interactions between lignin and Fe and Al oxides have also been previously

investigated where lignin has been observed to be protected from degradation via

associations with Fe and Al oxides (Miltner and Zech, 1998; Kleber et al., 2005; Mikutta et

al., 2006). Despite this, the extent of clay mineral protection of lignin in different soil types

has yet to be investigated. Suberin and cutin, which are major sources of aliphatic

constituents found in SOM (Kögel-Knabner, 2002), are believed to be chemically recalcitrant

(Lorenz et al., 2007). Aliphatic components of cutin have been found to preferentially sorb to

montmorillonite clay surfaces (Genest et al., 2014). Previous studies have also suggested that

aliphatic constituents in SOM may interact with lignin in soil through hydrophobic

interactions such as hydrogen bonding (Kaiser and Guggenberger, 2003; Kleber et al., 2007;

Kögel-Knabner et al., 2008). However, it is unclear if interactions with suberin and cutin,

major sources of aliphatic constituents, can protect lignin from degradation in soil.

The primary objective of this work was to examine the physical protection of lignin

by clay minerals and to determine if this extent varies among soils of different mineralogy.

Our goal was to also determine if suberin and cutin, major sources of aliphatic constituents

85

found in SOM, can protect lignin from degradation in soil. An additional aim was to assess

the correlation between the clay mineral and the suberin and cutin protection mechanisms of

lignin by establishing if one protection mechanism plays a more dominant role than the other.

To achieve these objectives, we applied a molecular-level approach to investigate grassland,

agricultural and forest soils. HF demineralization was performed to examine the extent of

clay mineral protection of lignin. Biomarkers were extracted from non-HF-treated and HF-

treated soil residues by base hydrolysis (BH) and copper (II) oxide (CuO) oxidation to yield

hydrolysable aliphatic lipids and lignin-derived phenols respectively (Otto and Simpson,

2007), which were quantified using gas chromatography-mass spectrometry. Lignin

monomer concentrations from BH and CuO oxidation extracts were used to assess the extent

of suberin and cutin protection of lignin. We hypothesized that lignin monomer

concentrations would increase after HF treatment once mineral-bound lignin was released,

signifying strong sorptive interactions between clay minerals and lignin. We also

hypothesized lignin monomer concentrations would be greater in CuO oxidation extracts that

have been base hydrolyzed. Performing BH prior to CuO oxidation is expected to release

lignin that was previously non-extractable due to OM protection (Otto and Simpson, 2007).

Lastly, we predicted that clay mineral associations with lignin are substantially stronger than

OM-OM interactions between suberin and cutin with lignin.

3.3 Materials and methods

3.3.1 Description of soil samples and sampling sites

Four soils were collected from various soil environments: grassland, agricultural and

forest. Sample details are listed in Table 3-1.

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Table 3-1: Selected properties of four soils used in this study.

Sample Soil Type Location Mineral

Horizon

Sand

(%)

Silt

(%)

Clay

(%)

Carbon Content (%) Mineralogy

Pre-HF

treatment

Post-HF

treatment

Northern

grasslanda

Orthic Black

Chernozem

Edmonton,

Alberta,

Canada

Ah 33 40 21 4.4 35.9 Montmorillonite, illite,

small amounts of

chlorite and kaolinite

Southern

grasslanda

Orthic Brown

Chernozem

Lethbridge,

Alberta,

Canada

Ah 46 31 16 2.1 32.4 Montmorillonite, illite,

small amounts of

chlorite and kaolinite

Agriculturalb Orthic

Melanic

Brunisol

Ottawa,

Ontario,

Canada

Ap 57 16 27 1.7 8.6 Feldspar, chlorite,

vermiculite, illite, small

amounts of

interstratified minerals

Forestc

Orthic

Humo-Ferric

Podzol

British

Columbia,

Canada

Ah 74 19 7 1.9 8.6 Iron and aluminum

sesquioxides

acompiled from (Otto and Simpson, 2006; Clemente et al., 2011)

bcompiled from (Ma et al., 2003; Clemente et al., 2012; Mitchell and Simpson, 2013)

ccompiled from (Keser and Pierre, 1973; Sanborn and Lavkulich, 1989; de Montigny and Nigh, 2009)

87

The northern grassland soil (Orthic Black Chernozem) was collected from the University of

Alberta Ellerslie Research Station in Edmonton, Alberta, Canada (Clemente et al., 2012)

while the southern grassland soil (Orthic Brown Chernozem) was collected from Lethbridge,

Alberta, Canada (Clemente and Simpson, 2013). These grassland soils both developed in an

arid to semi-arid climate and had a pH range of 6.0-6.9. Both soils possess a loam to clay

loam texture and contain montmorillonite and illite as well as chlorite and kaolinite to a

lesser extent (Otto and Simpson, 2006; Clemente and Simpson, 2013). Western Wheatgrass

(Agropyron smithii) was the dominant vegetation overlying both grassland soils (Otto and

Simpson, 2006). Although both soils originated from the Ah mineral horizons of their

respective sites, the C content of the northern grassland soil is slightly higher compared to

that of the southern grassland soil. The mean annual temperature for the grassland soil sites

ranges from 1.7°C to 3.3°C, while the mean annual precipitation for the northern and

southern grassland soil sites are reported to be 452 mm and 413 mm respectively (Janzen et

al., 1998; Otto and Simpson, 2006). The agricultural soil (Orthic Melanic Brunisol) was

sampled in 2007 from Agriculture and Agri-Food Canada’s Central Experimental Farm

located in Ottawa, Ontario, Canada, which had been under maize (Zea Mays L.) monoculture

for 14 years (Clemente et al., 2012). This sample is classified as a sandy clay loam and

contains feldspar, chlorite, vermiculite and illite and other interstratified minerals (MacLean

and Brydon, 1963; Clemente et al., 2012). The agricultural soil site had a mean annual

temperature of 5.8°C and a mean annual precipitation of 880 mm. The forest soil (Orthic

Humo-Ferric Podzol) was sampled in August 2013 from the Silviculture Treatments for

Ecosystem Management in the Sayward (STEMS) long term research installation set up by

the British Columbia Ministry of Forests and Range, near Gray Lake on Vancouver Island,

British Columbia, Canada (de Montigny and Nigh, 2009). Douglas fir (Pseudotsuga

88

menziesii) and western hemlock (Tsuga heterophyllsa) were the overlying vegetation in this

second-growth forest (Churchland et al., 2013), where the mean annual precipitation was

1503 mm and the mean annual temperature was 8.8°C. The forest soil sample possesses a

loamy to sandy-loam texture and contains mostly Fe and Al sesquioxides (Sanborn and

Lavkulich, 1989; de Montigny and Nigh, 2009). All four soil samples were sampled from the

0-15 cm depth. After collection, these soils were air-dried, passed through a 2 mm mesh

sieve, freeze-dried, finely ground using a mortar and pestle and stored at room temperature.

3.3.2 Determination of carbon (C) content

Soil C content was analyzed using a Thermo Flash 2000 elemental analyzer. Samples

were combusted at 950°C and the evolved gases were measured using a thermal conductivity

detector. Inorganic C was measured and was not detected in any of the soil samples.

Therefore, elemental C values represent total organic C (Otto and Simpson, 2006; Pautler et

al., 2010; Clemente and Simpson, 2013). The C content of soil samples were measured

before and after HF demineralization and are reported in Table 3-1.

3.3.3 Biomarker extractions and HF demineralization

Sequential chemical extractions were performed on the whole soil samples to remove

unbound biomarkers by solvent extraction, hydrolysable lipids by BH and finally, lignin-

derived phenols by CuO oxidation (Otto and Simpson, 2007). For solvent extraction, a 15 g

sample of each whole soil sample was sonicated three times for 15 min in 30 ml of DCM,

DCM: MeOH (1:1 v/v) and MeOH respectively. Extraction was repeated 4 times for each

sample to produce 60 g of solvent-extracted soil. The solvent extracts were discarded and the

combined residues were air-dried prior to demineralization and BH (Figure 3-1).

89

Figure 3-1: Flowchart of the extraction sequence used to isolate the extracts and residues. Whole soils were subject to solvent extraction to

remove free lipids. Extract 1 was isolated from solvent extraction and CuO oxidation. Extract 3 was isolated from solvent extraction, HF

demineralization and CuO oxidation. Extracts 2 and 4 were isolated in a similar fashion as extracts 1 and 3 respectively, except with the

addition of the base hydrolysis (BH) procedure.

90

HF demineralization was used to partially dissolve clay minerals that are suspected to

protect SOM (Rumpel et al., 2006) from chemolytic methods such as BH (Mikutta et al.,

2006) and CuO oxidation (Clemente and Simpson, 2013). To perform HF demineralization,

150 ml of 10% (v/v) HF solution was added to 30 g of residue from solvent extraction. The

suspensions were shaken for 24 h at room temperature, centrifuged and the supernatants were

discarded. This process was repeated nine additional times for a total of ten HF extractions.

The residues were washed with deionized water to remove salts and residual HF and then

freeze-dried.

To perform BH, the residues (2 g of solvent-extracted residues or 0.5 g of HF-treated

residues) were heated at 100°C for 3 h with 20 ml of 1 M methanolic KOH. The extracts

were then sonicated with DCM: MeOH (1:1 v/v), centrifuged and acidified to pH 1 using 6 M

HCl. Ester-bound lipids were isolated by liquid-liquid extraction with diethyl ether. After

concentration by rotary evaporation, the ether extracts were dried with Na2SO4, transferred to

2 ml vials and then dried under N2 gas. Residues from BH were air-dried for subsequent

extraction with CuO oxidation. To isolate lignin-derived phenols, 1 g of copper (II) oxide

(CuO), 100 mg of ammonium (II) iron sulfate hexahydrate [Fe(NH4)2(SO4)2•6H2O] and 15

ml of 2 M NaOH were added to the residues (0.5 g of base-hydrolyzed residues or 0.1 g of

HF-treated residues) which were heated at 170°C for 2.5 h. After acidification to pH 1 with 6

M HCl, extracts were stored for 1 h at room temperature in the dark to prevent reactions of

cinnamic acids. The extracts were centrifuged, liquid-liquid extracted in triplicate with

diethyl ether, concentrated by rotary evaporation and dried under N2 gas.

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3.3.4 Derivatization and gas chromatography-mass spectrometry (GC-MS)

Extracts from solvent extraction, BH and CuO oxidation were derivatized with 0.1 ml

of N,O-bis-(trimethylsilyl) trifluoroacetamide (BSTFA) and pyridine at 70°C for 1.5 h. Prior

to this step, 0.5 ml of N,N-Dimethylformamide dimethyl acetal was used to derivatize BH

extracts at 60°C for 30 min. Derivatized extracts were dried under N2 gas, and diluted with

hexane before gas chromatography-mass spectrometry analysis on an Agilent model 6890N

gas chromatograph coupled to an Agilent model 5973 quadrupole mass selective detector

with an Agilent model 7683 autosampler. Gas chromatography separation was achieved on a

HP-5MS fused silica capillary column (30 m x 0.25 mm inner diameter x 0.25 μm film

thickness). The gas chromatograph operating conditions were as follows: initial temperature

held at 65°C for 2 min, temperature increase from 65°C to 300°C at a rate of 6°C/min with a

final temperature of 300°C held for 20 min. Helium was used as a carrier gas with a flow rate

of 1.2 ml/min. Sample injection was achieved with a 2:1 split ratio with the injector

temperature at 280°C. The mass spectrometer was operated in electron impact ionization

mode at 70 eV ionization energy and scanned from m/z 50 to 650. Individual compounds

were assigned by comparison of mass spectra with analytical standards, NIST08 and W9N08

MS libraries and published data. The external quantification standard for ester-bound lipid

extracts was methyl tricosanoate while for lignin-derived phenol extracts, the trimethylsilyl

derivatives of vanillin and vanillic acid were used.

3.3.5 Lignin-derived phenol analysis and calculation of % mineral-protected

lignin and % suberin- and cutin- protected lignin

Eight main lignin-derived phenol monomers were identified and quantified (Hedges

and Ertel, 1982; Otto and Simpson, 2006; Clemente and Simpson, 2013): vanillyls (V;

vanillin, acetovanillone and vanillic acid), syringyls (S; syringaldehyde, acetosyringone and

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syringic acid) and cinnamyls (C; p-coumaric acid and ferulic acid). VSC is defined as the

total extractable concentrations of the vanillyl, syringyl and cinnamyl monomers.

The percentage of mineral-protected lignin was calculated by determining the

difference between the concentrations of lignin monomers in the non-HF-treated and HF-

treated residues, as denoted in Eqn. 1 as [pre-HF] and [post-HF] respectively (Figure 3-1).

% 𝑚𝑖𝑛𝑒𝑟𝑎𝑙 − 𝑝𝑟𝑜𝑡𝑒𝑐𝑡𝑒𝑑 𝑙𝑖𝑔𝑛𝑖𝑛 = ([𝑝𝑜𝑠𝑡 − 𝐻𝐹] − [𝑝𝑟𝑒 − 𝐻𝐹]

[𝑝𝑜𝑠𝑡 − 𝐻𝐹]) × 100%

= (𝑝𝑟𝑜𝑡𝑒𝑐𝑡𝑒𝑑 𝑙𝑖𝑔𝑛𝑖𝑛

𝑝𝑟𝑜𝑡𝑒𝑐𝑡𝑒𝑑 𝑎𝑛𝑑 𝑢𝑛𝑝𝑟𝑜𝑡𝑒𝑐𝑡𝑒𝑑 𝑙𝑖𝑔𝑛𝑖𝑛) × 100% [𝐸𝑞𝑛. 1]

The post-HF extracts included mineral-protected and unprotected lignin while the pre-HF

extracts only included unprotected lignin. For each lignin-derived phenol, the post-HF

concentration was calculated using the concentration from extract 3 while the pre-HF

concentration was calculated using the concentration from extract 1. This calculation was

repeated to compare the concentrations between extracts 2 [pre-HF] and 4 [post-HF].

The percentage of suberin- and cutin-protected lignin was calculated by determining

the difference between the concentrations of lignin monomers in the non-base-hydrolyzed

and base-hydrolyzed extracts, as indicated in Eqn. 2 as [pre-BH] and [post-BH] respectively

(Figure 3-1).

% 𝑆𝑢𝑏𝑒𝑟𝑖𝑛 𝑎𝑛𝑑 𝑐𝑢𝑡𝑖𝑛 − 𝑝𝑟𝑜𝑡𝑒𝑐𝑡𝑒𝑑 𝑙𝑖𝑔𝑛𝑖𝑛 = ([𝑝𝑜𝑠𝑡 − 𝐵𝐻] − [𝑝𝑟𝑒 − 𝐵𝐻]

[𝑝𝑜𝑠𝑡 − 𝐵𝐻]) × 100%

= (𝑝𝑟𝑜𝑡𝑒𝑐𝑡𝑒𝑑 𝑙𝑖𝑔𝑛𝑖𝑛

𝑝𝑟𝑜𝑡𝑒𝑐𝑡𝑒𝑑 𝑎𝑛𝑑 𝑢𝑛𝑝𝑟𝑜𝑡𝑒𝑐𝑡𝑒𝑑 𝑙𝑖𝑔𝑛𝑖𝑛) × 100% [𝐸𝑞𝑛. 2]

The post-BH extracts included suberin- and cutin-protected and unprotected lignin while the

pre-BH extracts only included unprotected lignin. For each lignin-derived phenol, the post-

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BH concentration was calculated using the concentration from extract 2 while the pre-BH

concentration was calculated using the concentration from extract 1. This calculation was

repeated to compare the concentrations between extracts 3 [pre-BH] and 4 [post-BH].

3.4 Results and discussion

3.4.1 Carbon (C) content

The C content increased in all soils after HF treatment (Table 3-1) due to enrichment

of SOM by partial demineralization, which is consistent with results from earlier studies

(Eusterhues et al., 2003; Mikutta et al., 2006; Rumpel et al., 2006). After HF treatment, the C

content in the northern grassland soil increased by about 8 times. HF treatment of the

southern grassland soil resulted in a 16 times increase in C content, which was the greatest

increase out of all four soils, indicating that HF demineralization of this soil was most

efficient. The C content of the agricultural and the forest soils increased approximately 4

times, exhibiting the lowest HF demineralization efficiencies out of all four soils (Table 3-1).

3.4.2 Extraction yields of lignin-derived phenols and mineral protection of

lignin

CuO oxidation yielded consistently high concentrations of vanillic acid in all soils

(Table 3-2) with pre-HF concentrations ranging from 20.6 ± 4.9 μg/g soil in the southern

grassland soil to 100.5 ± 5.3 μg/g soil in the forest soil. The post-HF concentrations ranged

from 218.7 ± 24.3 μg/g soil in the agricultural soil to 421.1 ± 27.5 μg/g soil in the forest soil.

Acetovanillone in the southern grassland soil experienced the greatest increase in

concentration after HF treatment: the post-HF concentration of 81.9 ± 11.8 μg/g soil was

almost 27 times greater than the pre-HF concentration of 3.3 ± 1.7 μg/g soil (Table 3-2). Out

of all lignin monomers from each soil, ferulic acid from the southern grassland soil yielded

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Table 3-2: Concentrations in μg/g soil of eight main lignin-derived phenols released after CuO oxidationa by comparing residues 1 (pre-HF)

and 3 (post-HF). Values were determined from triplicate samples (n = 3), unless otherwise indicated, followed by standard error. Compounds Concentration (μg lignin-derived phenol/g soil)

Northern grassland Southern grassland Agricultural Forest

Pre-HF Post-HF Pre-HF Post-HFb Pre-HF Post-HF Pre-HF Post-HF

Vanillin 10.0 ± 2.9 73.8 ± 23.3 6.3 ± 3.1 118.9 ± 12.9 7.3 ± 5.7 6.2 ± 1.5 21.1 ± 2.6 7.0 ± 0.1

Acetovanillone 6.2 ± 0.9 58.5 ± 6.8 3.3 ± 1.7 81.9 ± 11.8 2.3± 0.7 6.1 ± 2.2 15.5 ± 1.6 16.0 ± 2.4

Vanillic Acid 31.8 ± 2.1 296.9 ± 11.5 20.6 ± 4.9 391.5 ± 51.0 40.7 ± 3.2 218.7 ± 24.3 100.5 ± 5.3 421.1 ± 27.5

Sum of vanillyls 48.0 ± 3.7 429.2 ± 26.9 30.2 ± 6.0 592.3 ± 53.9 50.3 ± 6.6 231.0 ± 24.4 137.1 ± 6.1 444.1 ± 27.6

Syringaldehyde 20.5 ± 2.8 95.6 ± 24.8 16.2 ± 5.3 163.2 ± 5.1 28.2 ± 4.0 58.1 ± 24.8 4.5 ± 0.4 10.3 ± 0.1

Acetosyringone 9.8 ± 1.1 63.9 ± 11.1 6.4 ± 1.7 89.3 ± 3.6 18.4 ± 2.4 51.1 ± 19.8 2.2 ± 0.3 5.5 ± 0.7

Syringic Acid 22.7 ± 2.9 195.6 ± 15.2 14.3 ± 2.5 232.0 ± 28.3 37.8 ± 3.3 205.7 ± 19.9 7.9 ± 0.8 23.1 ± 1.6

Sum of syringyls 53.0 ± 4.2 355.1 ± 31.1 36.9 ± 6.1 484.5 ± 29.0 84.4 ± 5.7 314.9 ± 37.5 14.6 ± 0.9 38.9 ± 1.7

p-Coumaric Acid 23.1 ± 1.3 168.5 ± 6.7 17.3 ± 2.1 319.4 ± 5.8 73.1 ± 7.7 297.6 ± 10.3 34.2 ± 1.6 155.0 ± 5.7

Ferulic Acid 29.1 ± 1.1 239.5 ± 8.5 35.4 ± 4.1 623.0 ± 45.5 26.1 ± 2.0 113.9 ± 9.6 23.8 ± 2.0 91.5 ± 1.6

Sum of cinnamyls 52.2 ± 1.7 408.0 ± 10.8 52.7 ± 4.6 942.4 ± 45.9 99.2 ± 8.0 411.5 ± 14.1 58.0 ± 2.6 246.5 ± 5.9

Sum of VSC 153.2 ± 5.8 1192.3 ± 42.5 119.8 ± 9.7 2019.2 ± 76.5 233.9 ± 11.8 957.4 ± 46.9 209.7 ± 6.7 729.5 ± 28.2 a(Hedges and Ertel ,1982)

bPost-HF concentrations for the southern grassland soil were averaged from duplicate samples (n = 2)

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the highest post-HF concentration: 623.0 ± 45.5 µg/g soil. Out of all four soils, the southern

grassland soil exhibited a 17 % increase in total VSC concentration after HF treatment. In

contrast, the forest soil only experienced a 3% increase (Table 3-2). By comparing residues 2

and 4 (Figure 3-1), the percent concentration increase after HF treatment was most evident

for vanillyl monomers in all four soils, except for the forest soil. In this soil, the cinnamyl

monomers exhibited the greatest increase in concentration, out of the three VSC classes

(Figure 3-2). The percent concentration increases after HF treatment among the VSC classes

of each soil are comparable between residues 1 and 3 (Table 3-2) and residues 2 and 4

(Figure 3-2).

NG SG AGR FOR0

20

40

60

80

100

% m

inera

l p

rote

cte

d-lig

nin

Soil sample

Vanillyls (V)

Syringyls (S)

Cinnamyls (C)

Figure 3-2: Percentage (%) of mineral protected-lignin in each VSC class from triplicate

samples (n = 3) of all four soils after HF treatment (comparison of extracts 2 and 4). NG,

Northern grassland soil; SG, Southern grassland soil; AGR, Agricultural soil; FOR, Forest

soil. Total vanillyls = vanillin, acetovanillone, vanillic acid; total syringyls = syringaldehyde,

acetosyringone, syringic acid; total cinnamyls = p-coumaric acid and ferulic acid. Error bars

indicate standard error.

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After HF treatment, the total VSC concentrations of lignin monomers in all four soils

increased (Table 3-2 and Figure 3-2). HF demineralization likely facilitated the release of

lignin monomers that were previously bound to minerals. After HF demineralization, a

greater amount of lignin was extracted via CuO oxidation, leading to increased VSC

concentrations. This corroborates the claims of strong sorptive interactions between clay

minerals and lignin monomers reported previously (Miltner and Zech, 1998; Dai and

Johnson, 1999; Baldock and Skjemstad, 2000; Kaiser et al., 2002; Eusterhues et al., 2003;

Gonçalves et al., 2003; Mikutta et al., 2006; Rumpel et al., 2006; Clemente and Simpson,

2013). The percentage of mineral-protected lignin was calculated for each monomer of each

soil on the basis of total extract yield using Eqn.1, by comparing residues 1 and 3. They

ranged from 3.1 ± 0.2% in the forest soil to 95.9 ± 0.2% in the southern grassland soil (Table

3-3). This calculation also yielded negative percentages of mineral-protected lignin for

vanillin in the agricultural and forest soils (Table 3-3), signifying its post-HF concentration

was lower than its pre-HF concentration. This may have been due to C loss after HF

demineralization (Dai and Johnson, 1999; Rumpel et al., 2006; Eusterhues et al., 2007). For

example, Rumpel et al. (2006) reported C losses of 12-23% from the Ah horizon of a beech

forest soil after HF treatment while Eusterhues et al. (2007) observed C losses of 21-32%

from the A horizon of beech and oak forest soils after HF treatment. Dai and Johnson (1999)

observed a C loss of 51% from a B horizon forest soil, likely from the preferential removal of

O-alkyl and carboxyl C groups after HF treatment. However, their solid-state 13

C nuclear

magnetic resonance spectrum of the HF-treated B horizon soil also revealed a 4% decrease in

the aromatic C region (110-160 ppm), indicating that aromatic moieties may have been lost

during HF treatment as well.

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Table 3-3: Percentages (%) of mineral protectiona of eight main lignin-derived phenols released after CuO oxidation

b, calculated by

comparing the average yield of triplicate samples (n = 3), in mg/g soil from residues 1 (pre-HF) and 3 (post-HF), followed by standard error.

Northern grassland Southern grasslandc Agricultural Forest

Vanillyls

Vanillin 86.4 ± 0.4 94.7 ± 0.2 -18.5d ± 1.0 -200.6

d ± 0.4

Acetovanillone 89.3 ± 0.2 95.9 ± 0.2 62.5 ± 0.4 3.1 ± 0.2

Vanillic Acid 89.3 ± 0.1 94.7 ± 0.2 81.4 ± 0.1 76.1 ± 0.1

Syringyls

Syringaldehyde 78.6 ± 0.3 90.1 ± 0.1 51.5 ± 0.5 55.9 ± 0.1

Acetosyringone 84.6 ± 0.2 92.8 ± 0.1 64.0 ± 0.5 60.2 ± 0.2

Syringic Acid 88.4 ± 0.1 93.8 ± 0.2 81.6 ± 0.1 65.9 ± 0.1

Cinnamyls

p-Coumaric Acid 86.3 ± 0.1 94.6 ± 0.1 75.4 ± 0.1 77.9 ± 0.1

Ferulic Acid 87.8 ± 0.1 94.3 ± 0.1 77.1 ± 0.1 74.0 ± 0.1 a(([Post-HF]-[Pre-HF])/[Post-HF]) x 100%

bHedges and Ertel (1982)

cPost-HF concentrations for the southern grassland soil were averaged from duplicate samples (n = 2)

dThese negative values may have been due to carbon loss after HF demineralization (Dai and Johnson, 1999; Rumpel et al., 2006; Eusterhues

et al., 2007).

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The preferential degradation of lignin monomers has been well-documented in the

literature, where several studies have reported that vanillyl monomers are less susceptible to

degradation than other VSC classes (Ertel and Hedges, 1984; Hedges et al., 1988; Dignac et

al., 2005; Bahri et al., 2006; Hernes et al., 2013). In our study, vanillyl monomers were

observed to have the highest percentage of mineral protection out of all VSC classes. This is

likely because vanillyl monomers have more accessible bonding sites for clay mineral

interactions due to less methoxy groups (Bahri et al., 2006; Heim and Schmidt, 2007).

Consequently, vanillyl monomers are more likely to be protected from degradation by a

reduced accessibility to decomposers (Miltner and Zech, 1998; Heim and Schmidt, 2007).

This may explain why vanillyl monomers have been found to be more environmentally

persistent than syringyl and cinnamyl monomers (Ertel and Hedges, 1984; Dignac et al.,

2005; Kleber et al., 2005; Bahri et al., 2006; Heim and Schmidt, 2007).

Another consideration is that direct binding sites on clay minerals are finite (Burford

et al., 1964; Feller et al., 1992; Pennell et al., 1995; Kaiser and Guggenberger, 2000). The

capacity for minerals to adsorb OM is governed by mineral-specific properties such as

surface area, porosity and charge characteristics (Krull et al., 2003; Kleber et al., 2005). In

this study, the grassland soils exhibited higher amounts of mineral protection, as

demonstrated by increases in lignin monomer concentrations after HF treatment (Figure 3-2).

These soils contain higher montmorillonite clay content, which indicates there is higher

specific surface area available for sorption (Miltner and Zech, 1998; Heim and Schmidt,

2007; Clemente et al., 2012; Hernes et al., 2013). The southern grassland soil had slightly

higher amounts of mineral-protected lignin compared to the northern grassland soil even

though both soils have the same mineralogy (Table 3-1). In contrast, the agricultural and

forest soils exhibited lower amounts of mineral protection than the grassland soils. The

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agricultural soil contained large amounts of feldspar and chlorite while the forest soil is

dominated by Al and Fe sesquioxide clays, which indicates that there is less mineral surface

area available for sorption (Clemente et al., 2012). The proportion of mineral-protected lignin

was lower in the forest soil than the agricultural soil which is in agreement with earlier

studies (Eusterhues et al., 2005; John et al., 2005; Kögel-Knabner et al., 2008). Previous

research suggests that SOM may be protected from microbial degradation through strong

associations between Fe and Al sesquioxides because they form strong bonds by ligand

exchange and provide the largest surface area in acidic soils (Kaiser and Guggenberger,

2003; Kleber et al., 2005). The aromatic carboxyl and phenolic group-rich fractions sorb

strongly onto Fe and Al sesquioxides (Kaiser and Guggenberger, 2000; Filley et al., 2002).

However, we found that the forest soil in this study, mostly dominated by Fe and Al

sesquioxides, contains less mineral-protected lignin, compared to the grassland soils which

are mostly dominated by montmorillonite clays (Table 3-1). This could signify that organo-

mineral associations between lignin and montmorillonite clays are stronger than those with

Fe and Al sesquioxides. Despite this, previous research has indicated that highly crystalline

Fe oxides have lower sorption capacities than poorly crystalline Fe oxides (Kleber et al.,

2005; von Lützow et al., 2007). The forest soil may have contained highly crystalline Fe

oxides leading to a considerably lower sorption capacity compared to the grassland soils,

which are dominated by montmorillonite clays.

3.4.3 Suberin- and cutin-protected lignin

In addition to mineral protection mechanisms, lignin in soil could be stabilized by

interactions with other SOM constituents such as cutin and suberin through OM-OM

interactions (Kögel-Knabner et al., 1994; Almendros et al., 1996; Thevenot et al., 2013).

Similar to organo-mineral associations, a number of factors may control the interactions

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between nonpolar aliphatic SOM constituents and aromatic components in soil, including:

the type of cations present, pH conditions, cation bridging and hydrogen bonding

mechanisms. In addition, the physical conformation and varying hydrophobicity of aliphatic

moieties are also contributing factors (Almendros et al., 1996; Kaiser and Guggenberger,

2003; Wang and Xing, 2005; Kleber et al., 2007; Thevenot et al., 2013).

The percentage of suberin- and cutin- protected lignin was determined for lignin

monomers in each soil according to Eqn. 2 and the results are illustrated in Figure 3-3. The

pre-HF concentrations of lignin monomers released after BH and CuO oxidation from

extracts 1 and 2 were used to calculate suberin and cutin protection of lignin in the presence

of clay minerals. These were compared with post-HF concentrations of lignin monomers

released after BH and CuO oxidation from extracts 3 and 4 to determine suberin and cutin

protection in the absence of clay minerals.

Lignin monomer yields in the non-HF-treated residues (1 and 2) of all four soils

decreased after BH, resulting in negative percent changes (Figure 3-3). On the contrary,

lignin monomer yields in the HF-treated residues (3 and 4) increased after BH, particularly in

the southern grassland (Figure 3-3b) and agricultural (Figure 3-3c) soils, resulting in positive

percent changes. In the forest soil (Figure 3-3d), the concentrations of vanillin and

acetovanillone in the non-HF-treated residues increased after BH and these concentrations

further increased after HF treatment. The positive percent changes in lignin monomer

concentrations in the HF-treated residues may signify suberin and cutin protection of lignin.

In the southern grassland, agricultural and forest soils, suberin and cutin protection of lignin

was not detected until after the dissolution of clay minerals. In the agricultural soil, suberin

and cutin protection was mostly observed with the vanillyl monomers. This supports the prior

suggestion that bonding sites are more accessible in vanillyl monomers (Bahri et al., 2006).

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Figure 3-3: Changes in average concentrations of lignin monomers from triplicate samples

(n = 3) in the four soils: (a) Northern grassland; (b) Southern grassland; (c) Agricultural; (d)

Forest, suggesting suberin and cutin protection of lignin with mineral interference

(comparison of extracts 1 and 2) and without mineral interference (comparison of extracts 3

and 4). Error bars indicate standard error.

The percent changes in concentration of the cinnamyl monomers (p-coumaric acid

and ferulic acid) in all the soils except for the agricultural soil decreased further after HF

treatment. In the northern grassland soil, lignin monomer concentrations from base-

hydrolyzed extracts were lower in the HF-treated residues than in the non-HF-treated

residues (Figure 3-3a). Unlike the observations made from the three other soils, suberin and

cutin protection of lignin was not observed in the northern grassland soil, even in the absence

of clay minerals. Based on these observations, it is unlikely that suberin and cutin protect

lignin in the northern grassland soil because the amount of extractable lignin did not increase

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even after suberin and cutin monomers were extracted by BH. However, we do not have

direct evidence from this study to propose which other types of interactions, in addition to

organo-mineral interactions, may protect lignin from degradation in the northern grassland

soil.

Despite this, we still observed suberin and cutin protection of lignin in the southern

grassland, agricultural and forest soils. Our hypothesis related to suberin and cutin protection

of lignin complements findings from recent research which provides further evidence to

explain how lignin interacts with other macromolecules in soil via OM-OM interactions

(Thevenot et al., 2013). An isolation procedure was employed to extract a milled wall

enzymatic lignin fraction from an agricultural loamy soil. Combining evidence from earlier

research with their observations, Thevenot et al. (2013) indicated the possibility of strong

covalent bonding between lignin and lipid moieties which could originate from suberin and

cutin (Marseille et al., 1999; Quénéa et al., 2005; Dignac and Rumpel, 2006; Mendez-Millan

et al., 2012) and proposed that lignin-aliphatic interactions in soil could be responsible for

protecting lignin against biodegradation.

3.4.4 Implications for multilayer arrangement of organo-mineral interactions

Our data provide further evidence of interactions between clay minerals and lignin in

soil which is consistent with other studies (Miltner and Zech, 1998; Eusterhues et al., 2003;

Mikutta et al., 2006; Rumpel et al., 2006; Clemente and Simpson, 2013). Genest et al. (2014)

examined organo-clay complexes using 1H high resolution-magic angle spinning nuclear

magnetic resonance spectroscopy and demonstrated that aromatic moieties are tightly bound

to montmorillonite surfaces, which is in agreement with our findings. Genest et al. (2014)

also indicated that aromatic moieties in soil may not be accessible at the solid-liquid interface

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because they are likely to be protected within hydrophobic regions. This is also consistent

with our observations which suggest that macromolecules with hydrophobic domains such as

suberin and cutin may protect aromatic moieties such as lignin from degradation. Results

from our study suggest that organo-mineral interactions between lignin and clay minerals are

comparatively stronger than OM-OM interactions between lignin and macromolecules such

as suberin and cutin. Previous research has proposed a layered architecture of the sorption of

OM onto mineral surfaces, such that sorption occurs over multiple layers and not all the

adsorbed molecules are in contact with the mineral surface (Kaiser and Guggenberger, 2003;

Eusterhues et al., 2005; Wang and Xing, 2005; Kleber et al., 2007). Kaiser and Guggenberger

(2003) postulated that increased surface loading does not necessarily imply increased

sorption onto the mineral surface because sorbing molecules could form multiple layers

through hydrophobic interactions or cation bridging (von Lützow et al., 2006; Kleber et al.,

2007). Wang and Xing (2005) and Eusterhues et al. (2005) both observed constant mineral

surface coverage under different OM loading levels in separate investigations and

hypothesized that this is most likely due to OM-OM interactions, resulting in a multilayer

arrangement. Kleber et al. (2007) compiled observations from earlier work (Wershaw, 1993;

Chien et al., 1997; Engebretson and von Wandruszka, 1997; Engebretson and von

Wandruszka, 1998; von Wandruszka et al., 1998; Ferreira et al., 2001; Martin-Neto et al.,

2001; Kleber et al., 2007) to propose a conceptual model of organo-mineral interactions in

soils: SOM sorbs onto mineral surfaces in a zonal sequence, which consists of a contact zone,

a hydrophobic zone and a kinetic zone (observations from our work do not pertain to the

kinetic zone so it will not be further discussed). In the contact zone, the polar functional

groups of organic compounds form strong associations with the mineral surface and are

consequently protected from microbial degradation. The properties of this association depend

on the density and reactivity of the functional groups present on the mineral surface. In the

104

hydrophobic zone, organic compounds are weakly bound to one another via OM-OM

interactions. These molecules do not directly interact with the mineral surface and are more

readily desorbed and decomposed. These compounds possess alkyl or aromatic functional

groups which may originate from macromolecules like cutin (Kleber et al., 2007). The

conceptual model proposed by Kleber et al. (2007) can be used to explain the findings from

our study. After HF demineralization, higher concentrations of lignin monomers were

extracted which suggests that lignin, a macromolecule with polar functional groups, is likely

to be protected from degradation through association with the mineral surface in the contact

zone. The positive changes in concentrations of lignin monomers after HF demineralization

and BH observed in our study provide further evidence of weak interactions among organic

compounds in the hydrophobic zone (Kleber et al., 2007). This premise is further supported

by the isolation of milled wall enzymatic lignin performed by Thevenot et al. (2013), as

mentioned earlier. Our indirect observations of suberin and cutin protection of select lignin

monomers in certain types of soils support the postulation by Thevenot et al. (2013) of

lignin-aliphatic complexes in soil but additional analyses are needed to confirm this

hypothesis.

3.5 Conclusions

The mineral protection mechanism of lignin was demonstrated by the increases in all

lignin monomer concentrations in all four soils after HF treatment. The grassland soils,

notably the southern grassland soil, exhibited the highest amount of mineral protection

because it possesses the highest montmorillonite clay content, rendering more surface area

available for sorption. In contrast, lignin in the forest soil was the least protected because it

was dominated by Fe and Al sesquioxides, with less available surface area for sorption. Out

of all VSC classes, vanillyl monomers (vanillin, acetovanillone and vanillic acid) were the

105

most stabilized through organo-mineral associations, which concurs with literature findings.

Suberin and cutin protection of lignin, only evident after HF demineralization, was mostly

observed in the southern grassland and agricultural soils and in certain monomers from the

forest soil. These collective observations support the conceptual model of a multilayer

arrangement of organic compounds sorbed onto mineral surfaces. Aromatic components in

SOM such as lignin are strongly bound to mineral layers and are consequently protected from

degradation. However, lignin that interacts with suberin and cutin through weak hydrophobic

interactions may be more prone to degradation because it is not directly adsorbed to the

mineral surface. Our study demonstrates that organo-mineral interactions between clay

mineral surfaces and lignin are likely to be stronger than hydrophobic interactions between

suberin and cutin with lignin. These findings suggest that organo-mineral associations play a

more dominant role in OM preservation compared to OM-OM interactions. Further

investigation on OM-OM interactions, such as those between lignin and suberin and cutin, is

warranted to establish the mechanisms behind these interactions and how they ultimately

contribute to OM stabilization processes.

106

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112

Chapter 4: Conclusions and Future Directions

4.1 Summary

In Chapter 2, soil organic matter (SOM) was characterized in whole soils from Toolik

Lake, Alaska, USA, after 32 years of soil warming and nitrogen+ phosphorus (N+P)

fertilization. Biomarker methods and solid-state 13

C nuclear magnetic resonance (NMR)

techniques were used to analyze SOM composition as described in sections 2.3.4 and 2.3.5

respectively. With warming, N+P fertilization and warming +N+P fertilization, cutin-derived

compounds from leaves increased in the upper soil horizon while suberin-derived compounds

from roots decreased in the lower soil horizon, but these changes were not statistically

significant. In addition, SOM degradation was not enhanced by warming, N+P fertilization or

the combined treatment.

In Chapter 3, four soils of differing clay mineralogy from three Canadian provinces

were subjected to hydrofluoric (HF) acid demineralization and were characterized using

biomarker methods (section 3.3.3). Lignin was found to be mainly protected from extraction

via clay mineral interactions in all four soils. Suberin and cutin protection of lignin was also

observed, but not until after the dissolution of clay minerals via HF demineralization. Of the

three lignin phenol classes, vanillyl monomers were found to be the most stabilized by

interactions with clay minerals and associations with suberin and cutin.

4.1.1 Molecular-level characterization of Arctic soils (Chapter 2)

Upper and lower horizon soils were collected from Toolik Lake, Alaska, USA, after

32 years of warming and N+P fertilization. Biomarker methods were used to determine the

degree of SOM degradation, while solid-state 13

C NMR spectroscopy was employed to offer

113

an overview of SOM alteration in response to warming, N+P fertilization and warming +N+P

fertilization. In the solvent extracts of all these Arctic soils, long-chain n-alkanols and n-

alkanoic acids were comparatively more abundant than short-chain components, which

indicates there was more plant-derived than microbial-derived inputs (Lichtfouse et al., 1995;

Otto and Simpson, 2005; Simoneit, 2005). Low concentrations of simple sugars (glucose,

galactose and mannose) along with high O-alkyl carbon (C) signals in the solid-state 13

C

NMR spectra signified large amounts of cellulose inputs in these Arctic soils (Hatcher et al.,

1983; Salloum et al., 2002). Cutin-derived compounds in the upper horizon soils increased

but not significantly with warming, N+P fertilization and warming +N+P fertilization while

long-chain n-alkanes derived from epicuticular waxes did not vary. Coupled with NMR

analysis results which did not reveal any enrichment in alkyl compounds, this suggests there

was only a slight accumulation of cutin-derived inputs in the upper horizon under elevated

temperatures and increased nutrient availability. Cutin degradation was not promoted by any

of the treatments, which may be due to the chemical recalcitrance of cutin. Suberin-derived

inputs in the lower horizon slightly decreased with warming, N+P fertilization and the

combined treatment. The solid-state 13

C NMR spectra of the lower horizon soils showed

reductions in the alkyl C signals but not in the aromatic and phenolic C signals. The low

intensities of the aromatic and phenolic C signals from the solid-state 13

C NMR spectra

corresponded to the low concentrations of lignin-derived phenols from the copper (II) oxide

(CuO) oxidation extracts. Lignin oxidation was found to be suppressed in these Arctic soils,

likely due to low fungal activity as evidenced by the absence of ergosterol, a fungal

biomarker, from the solvent extracts. However, in the upper horizon of block 4, lignin

oxidation was observed to be most enhanced by N+P fertilization and least enhanced by

warming +N+P fertilization. The biomarker data demonstrated that there was considerable

spatial heterogeneity among the four replicate blocks, which may be attributable to the

114

topographical characteristics surrounding the site. The spatially complex landscape, reflected

through topographical controls such as water distribution, plant growth and soil horizon

development (Burke et al., 1999; Spielvogel et al., 2016), likely contributed to the different

alteration patterns of SOM composition among the blocks.

4.1.2 Clay mineral, suberin and cutin protection of lignin (Chapter 3)

The northern grassland soil was collected from the University of Alberta Ellerslie

Research Station in Edmonton, Alberta, while the southern grassland soil was collected from

Lethbridge, Alberta. The agricultural soil was sampled from Agricultural and Agri-Food

Canada’s Central Experimental Farm in Ottawa, Ontario, while the forest soil was sampled

from the Silviculture Treatments for Ecosystem Management in the Sayward (STEMS) long-

term research installation on Vancouver Island, British Columbia. HF demineralization was

performed to assess the extent of clay mineral protection of lignin-derived phenols. Base

hydrolysis was used to isolate suberin and cutin monomers. CuO oxidation was performed

before and after base hydrolysis and HF demineralization to determine the extent of suberin

and cutin protection of lignin. HF treatment enhanced the extractability of lignin-derived

phenols via CuO oxidation in all the soils, which supports previous observations of strong

sorptive interactions between clay minerals and lignin (Mikutta et al., 2006; Rumpel et al.,

2006). The highest percentage of mineral-protected lignin was observed in the grassland

soils, likely due to their higher montmorillonite clay content which provided greater surface

area for sorption (Miltner and Zech, 1998). The forest soil exhibited the lowest percentage of

mineral-protected lignin likely because the iron (Fe) and aluminum (Al) oxides contained

within this soil had a lower sorption affinity for lignin. The extractability of vanillyl

monomers increased the most in the grassland and agricultural soils after HF

demineralization, which suggests that the vanillyl monomers were the most stabilized

115

through organo-mineral interactions of the three lignin monomer classes. This concurs with

literature findings where vanillyl monomers have been reported to be more persistent in soil

compared to syringyl and cinnamyl monomers (Bahri et al., 2006). In the northern grassland

soil, the amount of extractable lignin did not increase after base hydrolysis, which indicates

that it is unlikely that suberin and cutin protect lignin from degradation in this soil. The

extractability of lignin-derived phenols from the base-hydrolyzed extracts of the southern

grassland, agricultural and forest soils did not increase until after the dissolution of clay

minerals by HF treatment. This suggests that organo-mineral interactions may play a more

dominant role in the protection of lignin compared to interactions with suberin and cutin.

4.2 Limitations and future work

The research presented in this thesis demonstrates that molecular-level techniques

showed alterations in SOM composition in response to environmental changes such as

elevated temperature and increased nutrient availability. This work also offered insight into

how SOM can be stabilized by associations with clay minerals and by interactions between

SOM components. Despite this, it is important to acknowledge the following limitations of

this research and to consider these recommendations for future work:

1. In the Arctic soils from Toolik Lake, Alaska (Chapter 2), the variability in the

biomarker data of the four replicate blocks in response to each treatment

demonstrated substantial spatial heterogeneity among the blocks. In this experiment,

only one soil sample was collected from the upper and lower soil horizons of each

treatment in each block. In the future, multiple samples from each block of each

treatment should be collected to gain a more representative picture of how each block

may respond to warming, N+P fertilization and warming +N+P fertilization. A

116

greater number of sampling points within each block is expected to minimize the

variability in the responses of each treatment (Schöning et al., 2006).

2. Based on the soils gathered from the treatments of the four replicate blocks, SOM

degradation in the Arctic soils from Toolik Lake, Alaska, was not enhanced after 32

years of warming and N+P fertilization (Chapter 2). However, it is unclear whether

this observation was unique, based on the topography of where these treatment blocks

were situated. The presence of underlying permafrost in Arctic soils may have led to

the impermeability of water infiltration (Sturm et al., 2005). As a result, water-

saturated conditions in the active layer may have inhibited SOM degradation

processes (Ping et al., 1998; Sistla et al., 2013). For example, in tundra ecosystems,

the greatest accumulation of soil C stocks has been reported to occur in low-lying

poorly-drained areas, while less soil C stocks tend to accumulate in upland areas with

more efficient drainage (Giblin et al., 1991; Michaelson et al., 1996; Hobbie et al.,

2000). The establishment of additional replicate blocks for the warming, N+P

fertilization and the warming +N+P fertilization treatments in upland and lowland

areas with contrasting drainage conditions at Toolik Lake may clarify whether the

suppression of SOM decomposition observed in this study was specific to the

topographic position of the treatment plots.

3. In the mineral protection experiment (Chapter 3), after HF demineralization, the

extractability of lignin monomers was highest in the grassland soils which contained

montmorillonite clays and lowest in the forest soil which contained mainly Fe and Al

oxides. This finding may indicate that montmorillonite clays, compared to Fe and Al

oxides, likely contain higher amounts of mineral-protected lignin. However, further

investigations on various types of Fe and Al oxides should be conducted until it can

be ascertained that montmorillonite clays have greater sorption affinity for lignin than

117

Fe and Al oxides. Previous reports have shown that highly crystalline Fe and Al

oxides have lower specific surface area and hence lower sorption capacities compared

to poorly crystalline Fe and Al oxides (Kleber et al., 2005; von Lützow et al., 2007).

For example, in an experiment conducted by Mikutta et al. (2006), a greater amount

of mineral protected-SOM was extracted from poorly crystalline Fe oxides than from

highly crystalline Fe oxides. Therefore, additional soils with differing crystalline

phases of Fe and Al oxides should be analyzed together with montmorillonite clays

according to the extraction procedure outlined in Chapter 3. This will allow for an

improved understanding of how the sorption affinity of montmorillonite for lignin

compares with various crystalline phases of Fe and Al oxides.

4. Of the three lignin monomer classes, the extractability of vanillyl monomers

increased the most after HF demineralization in the grassland and agricultural soils,

which signifies the vanillyl monomers exhibited the greatest amount of mineral

protection. Vanillyl monomers have been reported to be more environmentally

persistent than syringyl and cinnamyl monomers (Bahri et al., 2006). However, the

chemical mechanism behind why vanillyl monomers are more persistent is unclear

and would benefit from a mechanistic study. Batch equilibration experiments could

be conducted to examine the sorption affinity of the eight main lignin-derived phenol

monomers (vanillin, acetovanillone, vanillic acid, syringaldehyde, acetosyringone,

syringic acid, p-coumaric acid and ferulic acid) to clay minerals with different

mineralogical properties such as specific surface area and crystal structures (Xing et

al., 1996; Xing, 1997; Kaiser and Guggenberger, 2003; Cecchi et al., 2004). To

determine the degree of sorption of each lignin monomer, quantified data from high

performance liquid chromatography can be used to construct sorption isotherms and

to determine partition coefficients (Cecchi et al., 2004). The results of such

118

experiment would allow for a better understanding of the roles that vanillyl, syringyl

and cinnamyl monomers within the lignin macromolecule may play in sorption to

clay mineral surfaces.

The proposed work will provide more concrete evidence about whether or not our

observations of the alteration of SOM composition were reflective of landscape

characteristics. Further exploration into various types of crystallinity phases of Fe and Al

oxides along with sorption experiments between lignin monomers and clay minerals would

offer more detailed insights into how the extent of mineral protection may differ among

mineral surfaces and specific SOM components.

4.3 Research implications

With the onset of climate change, previous research has shown that warming and

increases in nutrient availability in soil may enhance SOM degradation in Arctic ecosystems.

The work presented in this thesis indicates that changes in temperature and nutrient

availability altered SOM composition but did not enhance SOM degradation overall.

However, such an observation may be ecosystem-specific. Our molecular-level data further

revealed that in an Arctic ecosystem where the landscape is spatially complex, SOM

composition may be altered differently in response to long-term warming and fertilization.

This suggests that the importance of landscape patterns should not be understated because

they can potentially contribute to the changes in SOM biogeochemistry that are expected to

occur in a warmer and more nutrient-rich environment. This information will serve as a

platform for developing more comprehensive climate change mitigation strategies in Arctic

ecosystems.

119

Moreover, this thesis offers further evidence that the associations between SOM

components and mineral surfaces play critical roles in the preservation of SOM. Our

observations suggest that clay mineral interactions may play a dominant part in the protection

of SOM. These findings also highlight the potential role that interactions between SOM

components may play in SOM protection mechanisms. This research demonstrates that clay

mineral associations, along with interactions between SOM components, collectively

contribute to SOM stabilization processes. This knowledge will provide a more mechanistic

understanding of how SOM stabilization can maximize C sequestration in soils, which may

potentially reduce C emissions into the atmosphere and alleviate contributions to the positive

climate feedback.

120

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122

Appendices


soil) of n-alkanes identified from the total solvent extracts of the upper and lower horizon soils of the

control, warming, N+P fertilization and warming +N+P fertilization treatments. All values are reported as mean ± standard error (n = 2).

Block Treatment

Control Warming N+P Fertilization Warming + N+P Fertilization

Upper Lower Upper Lower Upper Lower Upper Lower

n-Heneicosane 1 13.8 ± 1.1 11.5 ± 0.7 39.8 ± 0.3 25.5 ± 0.6 16.7 ± 0.7 16.9 ± 0.1 25.2 ± 1.4 30.9 ± 1.5

2 16.7 ± 0.4 5.9 ± 0.1 7.6 ± 0.2 12.8 ± 0.1 8.2 ± 0.5 7.9 ± 0.6 6.8 ± 0.3 9.4 ± 0.7

3 16.4 ± 1.8 9.3 ± 0.2 19.1 ± 0.2 5.3 ± 0.1 21.7 ± 0.1 6.0 ± 0.1 19.2 ± 2.3 NA

4 20.8 ± 2.6 5.7 ± 0.4 8.2 ± 0.2 32.9 ± 0.1 3.3 ± 0.2 3.4 ± 0.2 3.6 ± 0.1 5.8 ± 0.2

n-Docosane 1 7.15 ± 0.1 6.1 ± 0.7 16.2 ± 1.0 9.8 ± 0.5 7.1 ± 0.3 6.9 ± 0.2 24.5 ± 0.7 12.8 ± 1.0

2 10.3 ± 0.7 2.9 ± 0.1 4.6 ± 0.3 7.2 ± 0.4 5.6 ± 3.6 4.5 ± 0.4 3.5 ± 0.1 5.8 ± 0.1

3 11.3 ± 1.2 5.7 ± 0.2 20.6 ± 1.9 2.7 ± 0.1 5.6 ± 20.2 3.6 ± 0.1 9.9 ± 0.6 NA

4 6.9 ± 0.8 2.4 ± 0.2 4.6 ± 0.4 12.7 ± 1.6 1.8 ± 0.1 2.1 ± 0.1 2.0 ± 0.1 3.3 ± 0.1

n-Tricosane 1 20.4 ± 0.4 22.4 ± 2.2 83.5 ± 4.1 46.4 ± 0.6 22.1 ± 2.1 33.6 ± 0.3 40.2 ± 1.7 58.3 ± 3.0

2 41.8 ± 2.6 13.2 ± 0.5 13.9 ± 2.3 22.6 ± 0.5 11.3 ± 0.7 16.1 ± 1.2 10.5 ± 1.2 20.2 ± 1.0

3 21.2 ± 2.1 18.2 ± 0.6 85.0 ± 4.7 11.5 ± 0.2 16.6 ± 2.1 12.5 ± 0.2 26.6 ± 2.4 NA

4 18.1 ± 1.6 9.1 ± 0.4 26.9 ± 0.9 52.6 ± 0.6 7.0 ± 0.1 6.6 ± 0.2 6.7 ± 0.3 9.7 ± 0.1

n-Tetracosane 1 12.6 ± 0.1 6.6 ± 0.5 16.3 ± 1.4 17.2 ± 0.1 10.5 ± 1.1 12.5 ± 0.4 16.9 ± 0.9 20.8 ± 2.0

2 12.2 ± 0.5 6.6 ± 0.4 6.7 ± 0.9 8.5 ± 2.5 7.2 ± 1.2 4.6 ± 0.7 3.1 ± 0.1 8.2 ± 0.6

3 13.3 ± 0.8 8.1 ± 0.3 17.0 ± 0.1 3.2 ± 0.1 8.6 ± 1.0 3.1 ± 0.4 11.2 ± 0.7 NA

4 9.5 ± 1.3 3.0 ± 0.2 5.2 ± 0.7 10.0 ± 0.9 2.0 ± 0.1 2.1 ± 0.2 3.2 ± 0.1 3.0 ± 0.5

n-Pentacosane 1 17.9 ± 0.4 19.3 ± 1.3 63.3 ± 1.0 39.3 ± 1.9 24.5 ± 1.5 28.9 ± 0.6 45.6 ± 0.6 46.9 ± 2.1

2 29.2 ± 1.4 14.5 ± 1.1 16.2 ± 1.3 23.1 ± 1.0 22.2 ± 0.4 15.2 ± 0.7 17.7 ± 1.8 19.9 ± 0.8

3 35.2 ± 1.0 18.1 ± 1.7 35.2 ± 1.4 11.1 ± 0.1 15.4 ± 0.2 11.8 ± 0.5 53.4 ± 7.5 NA

4 18.6 ± 1.4 9.3 ± 0.6 15.7 ± 0.2 40.4 ± 2.0 7.9 ± 0.1 7.3 ± 0.6 6.8 ± 0.1 10.8 ± .5

n-Hexacosane 1 14.4 ± 0.3 3.1 ± 0.2 19.0 ± 0.6 15.2 ± 0.6 12.4 ± 0.8 4.0 ± 0.2 15.5 ± 0.6 10.9 ± 0.2

2 9.9 ± 1.7 1.3 ± 0.2 10.5 ± 0.1 7.1 ± 0.9 8.8 ± 0.7 5.8 ± 0.8 7.1 ± 0.8 4.7 ± 0.1

3 13.8 ± 1.9 8.1 ± 0.1 20.0 ± 1.4 7.5 ± 0.2 7.2 ± 0.9 3.4 ± 0.2 27.8 ± 4.7 NA

4 9.5 ± 0.8 3.4 ± 0.2 4.0 ± 0.6 30.1 ± 2.5 2.9 ± 0.2 2.5 ± 0.1 2.6 ± 0.3 12.8 ± 1.5

n-Heptacosane 1 48.4 ± 5.0 35.5 ± 3.1 90.3 ± 2.2 74.7 ± 4.7 61.8 ± 2.1 48.7 ± 0.1 131.6 ± 0.5 98.1 ± 9.9

2 67.3 ± 2.5 28.1 ± 1.0 37.8 ± 2.9 42.0 ± 1.1 44.8 ± 4.3 27.2 ± 1.3 46.7 ± 5.7 42.7 ± 2.4

3 68.6 ± 7.8 41.9 ± 1.2 119.0 ± 15.6 24.4 ± 0.2 32.5 ± 2.5 26.3 ± 0.7 126.9 ± 17.7 NA

4 61.5 ± 7.8 19.1 ± 2.7 32.7 ± 1.2 83.3 ± 1.9 26.3 ± 3.2 15.8 ± 1.5 26.7 ± 4.2 25.8 ± 3.5

n-Nonacosane 1 87.3 ± 9.7 43.8 ± 1.3 152.4 ± 4.0 152.1 ± 3.5 116.7 ± 7.1 63.2 ± 1.2 178.7 ± 4.7 149.9 ± 12.4

2 81.9 ± 41.8 46.5 ± 3.2 61.5 ± 8.2 78.6 ± 3.6 62.9 ± 6.3 48.9 ± 2.4 51.4 ± 7.2 87.8 ± 14.4

123

3 114.3 ± 21.8 65.7 ± 6.2 144.0 ± 33.1 69.6 ± 0.1 30.0 ± 1.0 37.4 ± 0.5 132.5 ± 4.5 NA

4 71.2 ± 4.7 34.1 ± 2.2 51.8 ± 4.0 175.9 ± 19.1 48.5 ± 6.8 27.2 ± 0.1 30.0 ± 2.0 37.2 ± 1.1

n-Hentriacontane 1 169.0 ± 5.6 38.2 ± 4.8 154.4 ± 11.0 200.8 ± 1.8 163.6 ± 16.9 81.0 ± 10.4 265.6 ± 16.1 183.2 ± 19.8

2 136.2 ± 5.1 54.5 ± 3.8 64.5 ± 4.5 129.5 ± 5.9 97.7 ± 8.3 89.9 ± 8.3 88.0 ± 13.7 163.7 ± 49.1

3 181.6 ± 32.8 83.3 ± 6.3 168.7 ± 33.3 54.8 ± 4.8 91.4 ± 8.0 50.3 ± 4.0 244.8 ± 12.4 NA

4 202.7 ± 10.3 44.1 ± 5.0 72.4 ± 2.2 163.6 ± 17.0 76.8 ± 11.5 41.3 ± 1.9 46.5 ± 4.1 63.0 ± 11.4

n-Tritriacontane 1 53.3 ± 2.8 79.6 ± 1.7 90.9 ± 8.7 83.1 ± 3.3 114.6 ± 7.2 76.8 ± 0.4 195.6 ± 2.2 86.3 ± 2.7

2 66.5 ± 10.4 25.9 ± 0.5 37.5 ± 0.3 39.8 ± 1.9 47.4 ± 1.0 35.0 ± 2.6 68.2 ± 6.8 24.5 ± 12.3

3 74.1 ± 8.3 54.2 ± 3.0 77.1 ± 0.8 37.7 ± 2.3 68.7 ± 4.5 53.7 ± 0.1 311.9 ± 39.5 NA

4 112.6 ± 7.5 25.2 ± 0.6 29.5 ± 1.1 84.5 ± 0.8 39.6 ± 2.0 17.3 ± 1.9 30.5 ± 1.0 23.3 ± 5.1

Total n-alkanes

1 444.3 ± 12.7 266.0 ± 6.7 726.1 ± 15.4 664.1 ± 7.3 550.0 ± 20.1 372.4 ± 33.7 939.4 ± 17.1 698.1 ± 25.9

2 472.1 ± 43.6 199.5 ± 5.3 260.7 ± 10.2 371.3 ± 7.8 316.0 ± 12.0 255.1 ± 9.3 302.3 ± 18.0 386.8 ± 52.7

3 549.7 ± 41.2 312.5 ± 9.6 705.7 ± 49.8 227.7 ± 5.3 297.8 ± 9.9 208.2 ± 4.1 964.2 ± 46.3 NA

4 531.4 ± 16.1 155.4 ± 6.1 251.0 ± 5.1 685.8 ± 25.9 216.1 ± 13.9 125.8 ± 3.2 158.6 ± 6.3 194.7 ± 13.1

NA = not analyzed (the lower horizon soil sample for the warming +N+P fertilization treatment of block 3 was unavailable)

124


soil) of n-alkanols identified from the total solvent extracts of the upper and lower horizon soils of the

control, warming, N+P fertilization and warming +N+P fertilization treatments. All values are reported as mean ± standard error (n = 2).

Block Treatment



n-Pentadecanol 1 8.3 ± 0.6 1.1 ± 0.2 2.9 ± 0.1 1.6 ± 0.1 3.5 ± 0.3 1.6 ± 0.1 16.3 ± 1.8 2.8 ± 0.4

2 7.3 ± 0.9 0.4 ± 0.1 5.8 ± 0.3 1.0 ± 0.1 10.1 ± 0.7 0.9 ± 0.1 4.1 ± 0.1 0.8 ± 0.1

3 5.8 ± 0.6 0.8 ± 0.1 22.9 ± 2.0 0.4 ± 0.1 83.2 ± 5.1 0.9 ± 0.1 12.6 ± 1.7 NA

4 10.1 ± 0.4 0.5 ± 0.1 4.4 ± 0.6 3.2 ± 0.2 0.9 ± 0.1 0.3 ± 0.1 0.7 ± 0.1 0.6 ± 0.1

n-Hexadecanol 1 6.5± 0.1 5.1 ± 0.4 5.7 ± 0.2 4.1 ± 0.2 10.8 ± 0.2 4.6 ± 0.2 27.1 ± 0.8 7.2 ± 0.7

2 4.4 ± 0.6 1.6 ± 0.1 11.4 ± 1.1 5.2 ± 0.2 12.8 ± 0.9 1.2 ± 0.2 13.4 ± 0.7 2.9 ± 0.2

3 19.9 ± 1.7 2.9 ± 0.1 21.4 ± 3.8 2.2 ± 0.1 33.7 ± 2.3 2.2 ± 0.1 37.0 ± 5.6 NA

4 16.4 ± 0.7 2.8 ± 0.3 3.5 ± 0.9 4.8 ± 0.5 3.7 ± 0.2 1.2 ± 0.1 1.6 ± 0.3 1.9 ± 0.1

n-Octadecanol 1 7.4 ± 0.3 8.6 ± 0.5 12.3 ± 0.2 7.4 ± 0.6 7.6 ± 0.2 7.4 ± 0.1 16.3 ± 1.0 10.1 ± 0.4

2 7.7 ± 0.4 3.8 ± 0.2 4.9 ± 0.2 6.7 ± 0.1 6.2 ± 0.1 3.7 ± 0.3 6.6 ± 0.4 5.8 ± 0.5

3 10.1 ± 1.6 6.9 ± 0.3 12.9 ± 0.1 4.5 ± 0.1 9.7 ± 0.4 4.9 ± 0.1 19.9 ± 3.1 NA

4 9.8 ± 0.3 3.3 ± 0.2 3.6 ± 0.2 8.2 ± 0.4 3.3 ± 0.4 2.4 ± 0.1 2.8 ± 0.1 3.5 ± 0.1

n-Eicosanol 1 49.7 ± 2.7 91.7 ± 3.2 53.1 ± 1.1 61.1 ± 0.7 61.3 ± 6.6 78.0 ± 0.5 68.9 ± 3.7 81.2 ± 4.1

2 41.3 ± 1.5 43.1 ± 1.2 30.4 ± 1.2 58.7 ± 0.2 20.2 ± 0.5 34.3 ± 0.1 42.5 ± 2.0 66.3 ± 4.9

3 63.7 ± 7.0 82.1 ± 3.5 34.7 ± 2.1 44.1 ± 0.1 54.5 ± 7.9 70.4 ± 0.9 43.8 ± 6.6 NA

4 47.3 ± 3.2 32.0 ± 2.1 21.8 ± 2.1 52.9 ± 2.0 21.7 ± 0.4 22.1 ± 1.0 21.3 ± 1.0 29.3 ± 1.0

n-Heneicosanol 1 12.9 ± 0.5 12.1 ± 0.8 26.4 ± 1.6 19.3 ± 1.1 20.0 ± 1.7 16.1 ± 0.4 28.2 ± 1.5 22.1 ± 0.9

2 18.5 ± 0.2 7.9 ± 0.4 11.5 ± 0.4 14.4 ± 0.3 10.6 ± 0.6 10.3 ± 1.6 10.8 ± 1.1 11.5 ± 0.9

3 21.0 ± 2.5 11.6 ± 0.4 30.5 ± 0.6 9.0 ± 0.1 18.1 ± 1.3 8.5 ± 0.1 20.0 ± 2.7 NA

4 14.7 ± 1.8 4.8 ± 0.4 8.8 ± 0.6 19.3 ± 0.9 7.1 ± 0.1 4.5 ± 0.3 6.4 ± 0.5 6.5 ± 0.3

n-Docosanol 1 89.6 ± 0.3 104.3 ± 1.8 221.1 ± 34.2 131.0 ± 0.2 198.3 ± 23.3 115.0 ± 0.1 287.0 ± 21.4 137.8 ± 6.6

2 125.3 ± 3.2 76.0 ± 1.9 86.3 ± 6.8 102.5 ± 0.3 116.0 ± 1.8 64.0 ± 0.5 155.8 ± 9.3 100.4 ± 7.3

3 126.3 ± 3.5 93.7 ± 3.4 172.6 ± 3.7 82.1 ± 0.5 120.0 ± 16.7 65.6 ± 1.3 254.0 ± 41.9 NA

4 108.5 ± 6.4 45.1 ± 2.9 109.1 ± 1.5 122.7 ± 4.3 76.7 ± 0.7 43.4 ± 2.0 75.0 ± 2.6 53.9 ± 1.7

n-Tricosanol 1 144.8 ± 15.1 17.2 ± 0.8 44.3 ± 0.2 28.4 ± 0.5 57.8 ± 4.2 20.8 ± 0.4 63.2 ± 3.3 31.7 ± 3.0

2 42.0 ± 3.7 12.5 ± 0.5 92.3 ± 16.7 18.8 ± 0.2 24.8 ± 0.7 12.7 ± 0.3 24.3 ± 2.0 16.1 ± 1.9

3 115.9 ± 2.7 16.2 ± 0.5 534.2 ± 19.1 11.9 ± 0.2 48.2 ± 7.5 11.5 ± 0.4 59.5 ± 5.5 NA

4 191.9 ± 2.6 7.9 ± 0.6 32.0 ± 5.2 29.0 ± 1.4 19.7 ± 1.0 7.2 ± 0.5 13.8 ± 0.3 9.5 ± 0.3

n-Tetracosanol 1 81.0 ± 2.2 73.5 ± 3.3 192.2 ± 0.9 119.9 ± 2.9 209.2 ± 25.6 93.4 ± 1.1 283.1 ± 17.3 127.8 ± 5.9

2 150.5 ± 5.8 59.0 ± 0.6 121.2 ± 8.6 90.0 ± 1.0 195.5 ± 3.9 46.9 ± 0.8 173.7 ± 13.0 79.2 ± 6.5

3 144.2 ± 17.6 74.4 ± 3.6 291.6 ± 12.1 48.6 ± 0.1 172.7 ± 6.8 44.3 ± 0.9 356.5 ± 64.5 NA

4 159.9 ± 8.5 37.1 ± 2.9 140.0 ± 7.8 127.4 ± 2.8 101.9 ± 8.7 32.6 ± 1.3 82.1 ± 11.0 46.0 ± 2.9

n-Pentacosanol 1 24.9 ± 2.4 13.9 ± 1.7 36.2 ± 5.8 19.8 ± 1.8 51.0 ± 2.4 16.4 ± 1.1 58.8 ± 3.6 31.3 ± 9.4

2 29.5 ± 2.8 10.1 ± 1.3 27.1 ± 1.0 16.8 ± 0.6 40.0 ± 1.7 9.6 ± 1.1 30.4 ± 0.7 12.8 ± 0.2

125

NA = not analyzed (the lower horizon soil sample for the warming +N+P fertilization treatment of block 3 was unavailable)

3 46.0 ± 6.7 14.9 ± 0.8 56.2 ± 4.3 8.9 ± 1.7 41.6 ± 4.6 8.0 ± 1.6 94.4 ± 15.8 NA

4 45.0 ± 3.1 6.5 ± 0.3 19.9 ± 1.6 26.2 ± 1.0 13.8 ± 0.1 7.6 ± 0.3 16.8 ± 0.3 8.4 ± 0.1

n-Hexacosanol 1 63.3 ± 2.1 52.5 ± 6.4 151.8 ± 0.6 87.3 ± 0.4 157.4 ± 8.5 64.4 ± 1.3 392.4 ± 25.1 107.2 ± 7.9

2 114.3 ± 2.1 41.4 ± 0.1 107.2 ± 14.6 68.3 ± 0.4 418.8 ± 38.5 38.5 ± 0.2 174.3 ± 22.3 59.1 ± 4.4

3 106.4 ± 12.5 57.1 ± 4.7 414.2 ± 17.5 39.0 ± 1.1 183.9 ± 22.1 33.0 ± 0.4 402.4 ± 73.0 NA

4 155.1 ± 12.3 26.2 ± 1.5 109.8 ± 2.7 114.3 ± 9.4 68.8 ± 0.1 26.8 ± 0.9 67.1 ± 0.3 41.7 ± 0.8

n-Heptacosanol 1 18.6 ± 0.1 9.5 ± 1.2 31.2 ± 0.3 15.4 ± 2.5 50.9 ± 3.2 10.6 ± 0.2 78.1 ± 12.3 25.4 ± 4.1

2 40.1 ± 4.3 7.8 ± 1.3 23.4 ± 0.8 14.9 ± 0.7 52.4 ± 2.9 7.8 ± 0.1 37.7 ± 5.0 9.0 ± 1.4

3 34.6 ± 2.1 11.4 ± 1.0 67.0 ± 5.2 7.5 ± 0.2 32.2 ± 4.7 8.3 ± 1.2 95.7 ± 16.3 NA

4 37.7 ± 2.8 4.1 ± 0.5 20.0 ± 0.6 42.7 ± 2.1 17.1 ± 0.2 6.1 ± 0.5 13.8 ± 1.6 8.1 ± 0.3

n-Octacosanol 1 86.3 ± 11.8 53.6 ± 5.3 151.3 ± 1.2 112.3 ± 2.6 194.4 ± 19.3 71.7 ± 13.8 622.2 ± 39.5 146.8 ± 7.6

2 143.1 ± 8.2 50.3 ± 2.1 97.3 ± 19.7 88.7 ± 2.4 408.2 ± 47.3 50.0 ± 0.1 257.8 ± 37.4 69.9 ± 6.8

3 130.3 ± 2.8 64.3 ± 0.6 381.8 ± 47.4 53.3 ± 1.5 95.5 ± 4.8 42.4 ± 2.1 758.2 ± 128.1 NA

4 118.2 ± 19.1 30.2 ± 2.1 127.2 ± 0.4 115.4 ± 9.2 116.5 ± 6.2 35.8 ± 1.8 73.2 ± 2.6 48.6 ± 1.5

n-Nonacosan-10-ol 1 21.5 ± 0.3 9.6 ± 0.1 35.4 ± 1.9 25.0 ± 3.6 93.3 ± 3.4 12.3 ± 0.9 123.0 ± 2.8 14.7 ± 0.7

2 21.2 ± 0.5 11.6 ± 0.2 42.4 ± 1.0 9.8 ± 0.6 80.9 ± 0.9 8.8 ± 0.6 57.8 ± 0.9 9.0 ± 0.4

3 44.9 ± 3.8 13.1 ± 0.4 72.9 ± 1.0 6.3 ± 0.1 38.7 ± 3.6 8.9 ± 0.8 158.7 ± 13.4 NA

4 46.8 ± 1.5 8.5 ± 1.0 85.7 ± 2.4 27.5 ± 0.2 18.9 ± 1.8 8.3 ± 0.8 18.5 ± 2.7 6.2 ± 1.2

n-Triacontanol 1 54.2 ± 4.3 28.2 ± 1.4 107.2 ± 3.2 66.2 ± 3.3 131.4 ± 5.5 56.6 ± 0.3 279.8 ± 14.2 67.7 ± 0.1

2 69.4 ± 5.5 18.8 ± 0.3 72.7 ± 10.9 29.5 ± 1.8 126.9 ± 9.4 22.2 ± 0.7 93.7 ± 11.3 29.5 ± 2.5

3 78.0 ± 8.9 32.0 ± 2.8 199.6 ± 6.5 21.9 ± 0.4 113.1 ± 11.4 32.7 ± 1.5 307.1 ± 36.1 NA

4 132.3 ± 17.2 20.2 ± 1.6 55.8 ± 3.2 81.0 ± 0.8 46.3 ± 1.3 15.0 ± 1.2 34.3 ± 1.9 23.2 ± 1.5

n-Dotriacontanol 1 23.0 ± 3.5 7.3 ± 1.3 71.1 ± 3.3 29.5 ± 1.4 71.4 ± 0.1 19.8 ± 2.2 119.0 ± 0.9 17.2 ± 7.5

2 53.8 ± 2.9 5.4 ± 0.4 34.7 ± 2.0 14.7 ± 1.3 70.1 ± 2.7 10.2 ± 0.1 45.1 ± 6.0 12.8 ± 0.3

3 51.6 ± 0.2 15.5 ± 2.5 102.6 ± 15.0 12.6 ± 1.8 79.6 ± 10.4 11.0 ± 0.4 179.8 ± 27.0 NA

4 64.8 ± 7.6 11.1 ± 1.3 21.7 ± 2.9 43.3 ± 0.5 13.9 ± 0.6 10.4 ± 0.2 16.8 ± 0.3 10.2 ± 1.7

Short-chain vs. long-chain n-alkanols

Short-chain C15-C19 1 22.1 ± 0.7 14.7 ± 0.7 20.9 ± 0.3 13.1 ± 0.6 21.9 ± 0.4 13.6 ± 0.1 60.0 ± 2.2 23.9 ± 0.9

Long-chain C20-C32 669.9 ± 20.5 473.5 ± 10.2 1121.3 ± 35.1 715.2 ± 7.3 1296.4 ± 42.1 575.2 ± 51.4 2403.6 ± 57.9 810.9 ± 19.7

Short-chain C15-C19 2 19.5 ± 1.1 5.8 ± 0.2 22.0 ± 1.2 12.9 ± 0.3 29.0 ± 1.2 5.8 ± 0.3 24.1 ± 0.8 11.3 ± 0.5

Long-chain C20-C32 849.0 ± 14.0 344.0 ± 6.2 746.5 ± 33.5 527.1 ± 3.7 1564.5 ± 62.0 315.5 ± 2.3 1104.0 ± 48.5 475.7 ± 14.1

Short-chain C15-C19 3 35.7 ± 2.4 10.5 ± 0.3 57.1 ± 4.3 7.1 ± 0.1 126.7 ± 5.7 8.0 ± 0.1 69.5 ± 6.6 NA

Long-chain C20-C32 962.8 ± 29.3 486.4 ± 8.7 2358.0 ± 58.2 345.3 ± 3.5 998.0 ± 35.4 344.6 ± 3.9 2730.1 ± 174.5 NA

Short-chain C15-C19 4 36.3 ± 0.8 6.6 ± 0.4 11.5 ± 1.1 16.2 ± 0.7 7.9 ± 0.5 3.8 ± 0.1 5.1 ± 0.3 7.3 ± 0.1

Long-chain C20-C32 1122.19 ± 34.2 233.6 ± 5.9 751.8 ± 11.4 801.7 ± 14.6 522.4 ± 11.0 220.0 ± 3.7 439.1 ± 12.2 291.8 ± 4.7

Total n-alkanols

1 692.0 ± 20.5 488.2 ± 10.2 1142.2 ± 35.1 728.3 ± 7.3 1318.3 ± 42.1 588.8 ± 51.4 2463.3 ± 57.9 834.7 ± 19.7

2 868.5 ± 14.1 349.8 ± 3.8 768.6 ± 33.5 540.0 ± 3.7 1593.5 ± 62.0 321.3 ± 2.4 1128.1 ± 48.5 487.0 ± 14.1

3 998.6 ± 29.4 497.0 ± 8.7 2415.1 ± 58.4 352.4 ± 3.2 1124.7 ± 35.8 352.6 ± 3.9 2800.0 ± 174.7 NA

4 1158.5 ± 34.2 240.2 ± 5.9 763.4 ± 11.4 817.9 ± 14.6 530.3 ± 11.0 223.8 ± 3.7 444.2 ± 12.2 299.1 ± 4.7

126


soil) of n-alkanoic acids and total aliphatic compounds identified from the total solvent extracts of the upper

and lower horizon soils of the control, warming, N+P fertilization and warming +N+P fertilization treatments. All values are reported as mean

± standard error (n = 2).

Block Treatment



n-Tetradecanoic acid 1 51.1 ± 1.8 6.8 ± 0.6 30.2 ± 0.2 14.2 ± 0.1 32.4 ± 5.4 9.7 ± 0.1 93.0 ± 4.7 18.4 ± 1.0

2 26.4 ± 1.7 3.9 ± 0.2 35.4 ± 0.3 8.9 ± 0.3 26.4 ± 0.8 4.7 ± 0.3 24.2 ± 1.5 5.5 ± 0.5

3 32.8 ± 3.8 5.3 ± 0.3 124.6 ± 12.8 3.9 ± 0.2 357.6 ± 1.7 3.6 ± 0.1 42.9 ± 3.9 NA

4 122.9 ± 1.9 4.2 ± 0.4 22.5 ± 1.7 19.2 ± 0.4 9.6 ± 1.5 2.0 ± 0.1 5.8 ± 0.6 3.5 ± 0.2

n-Pentadecanoic acid 1 8.4 ± 0.6 4.7 ± 0.5 13.7 ± 0.1 7.2 ± 0.5 10.9 ± 1.0 5.6 ± 0.1 27.1 ± 1.5 8.1 ± 0.1

2 12.2 ± 1.1 1.6 ± 0.1 8.2 ± 0.3 5.0 ± 0.3 12.5 ± 0.1 2.0 ± 0.1 6.2 ± 0.4 2.9 ± 0.2

3 11.4 ± 1.0 4.0 ± 0.1 24.7 ± 1.3 2.3 ± 0.1 16.0 ± 1.6 2.9 ± 0.1 16.5 ± 1.9 NA

4 38.3 ± 2.4 2.7 ± 0.1 5.6 ± 0.5 9.8 ± 0.1 3.7 ± 0.2 1.1 ± 0.1 1.9 ± 0.1 1.8 ± 0.1

n-Hexadecenoic acid (C16:1) 1 16.9 ± 0.4 1.8 ± 0.2 19.3 ± 1.5 4.3 ± 0.2 13.6 ± 1.9 2.6 ± 0.1 22.9 ± 3.2 6.0 ± 0.1

2 10.5 ± 0.5 0.7 ± 0.1 9.1 ± 0.6 4.6 ± 0.1 13.0 ± 0.1 1.1 ± 0.1 5.4 ± 0.2 ND

3 16.2 ± 1.9 1.4 ± 0.1 51.5 ± 1.2 1.0 ± 0.1 27.8 ± 0.1 1.0 ± 0.1 25.4 ± 2.8 NA

4 17.6 ± 2.3 1.5 ± 0.1 10.7 ± 0.7 10.0 ± 0.1 3.2 ± 0.1 0.3 ± 0.1 1.6 ± 0.1 0.7 ± 0.1

n-Hexadecanoic Acid 1 116.5 ± 1.3 39.0 ± 0.9 120.2 ± 2.0 65.1 ± 0.7 135.8 ± 22.2 41.7 ± 0.6 232.9 ± 14.4 75.8 ± 4.2

2 86.2 ± 4.1 22.1 ± 0.6 107.7 ± 11.9 49.9 ± 0.9 177.1 ± 0.7 20.1 ± 0.5 110.6 ± 2.2 29.0 ± 2.7

3 148.8 ± 14.8 24.9 ± 1.0 361.1 ± 3.5 17.5 ± 0.4 147.7 ± 19.0 15.9 ± 0.3 249.4 ± 34.3 NA

4 137.4 ± 9.3 22.7 ± 0.5 80.8 ± 3.2 65.9 ± 2.7 52.4 ± 2.8 10.9 ± 1.5 34.3 ± 1.5 17.2 ± 0.3

n-Heptadecanoic Acid 1 10.6 ± 0.1 5.7 ± 0.1 9.3 ± 0.4 6.3 ± 1.0 8.4 ± 1.3 4.7 ± 0.1 13.6 ± 2.4 9.3 ± 0.6

2 7.2 ± 0.1 2.2 ± 0.1 6.6 ± 0.9 7.9 ± 0.2 10.5 ± 0.4 3.1 ± 0.1 5.8 ± 0.1 4.3 ± 0.3

3 13.8 ± 1.9 4.5 ± 0.2 15.7 ± 1.7 1.9 ± 0.1 12.3 ± 0.4 2.9 ± 0.1 19.2 ± 3.0 NA

4 10.6 ± 2.0 2.5 ± 0.1 4.4 ± 0.8 10.9 ± 0.7 3.4 ± 0.1 1.4 ± 0.1 2.3 ± 0.1 2.7 ± 0.1

n-Octadecadienoic acid (C18:2) 1 10.6 ± 0.5 2.6 ± 0.4 26.8 ± 1.1 6.8 ± 0.6 49.8 ± 10.0 3.8 ± 0.3 82.8 ± 6.3 8.4 ± 0.5

2 19.1 ± 0.8 0.8 ± 0.1 25.4 ± 3.7 7.6 ± 0.3 45.3 ± 1.5 1.8 ± 0.1 21.0 ± 1.3 1.7 ± 0.1

3 24.8 ± 3.5 1.3 ± 0.1 177.2 ± 3.5 1.9 ± 0.1 62.3 ± 6.9 1.5 ± 0.1 65.9 ± 6.6 NA

4 71.6 ± 7.4 3.3 ± 0.1 24.3 ± 1.4 20.8 ± 0.3 9.6 ± 0.8 0.8 ± 0.1 7.9 ± 0.5 2.0 ± 0.2

n-Octadecenoic acid (C18:1) 1 39.7 ± 1.0 5.2 ± 0.1 64.9 ± 1.1 15.8 ± 0.5 78.6 ± 13.1 11.9 ± 2.1 143.3 ± 12.3 19.7 ± 1.6

2 45.6 ± 2.1 1.8 ± 0.1 73.1 ± 8.9 19.3 ± 0.1 83.2 ± 1.3 7.1 ± 0.9 41.6 ± 2.0 5.3 ± 0.1

3 63.4 ± 6.4 4.1 ± 0.2 341.6 ± 9.5 3.8 ± 0.2 120.0 ± 15.8 2.5 ± 0.1 114.5 ± 17.8 NA

4 149.9 ± 18.3 8.8 ± 0.4 91.2 ± 0.9 44.3 ± 3.1 28.2 ± 1.4 2.1 ± 0.2 15.4 ± 0.6 4.9 ± 0.3

n-Octadecenoic acid (C18:1) 1 17.0 ± 1.1 5.3 ± 0.9 26.8 ± 0.3 10.0 ± 0.4 23.1 ± 3.6 5.3 ± 0.3 37.6 ± 4.2 10.6 ± 0.8

2 17.4 ± 2.1 1.7 ± 0.1 14.9 ± 1.8 8.1 ± 0.3 24.2 ± 0.3 1.9 ± 0.1 9.5 ± 0.8 2.8 ± 0.1

3 23.3 ± 2.1 4.2 ± 0.1 173.9 ± 12.7 2.4 ± 0.1 40.7 ± 3.8 2.3 ± 0.1 30.0 ± 3.6 NA

4 23.8 ± 2.0 2.3 ± 0.1 20.7 ± 1.0 15.3 ± 0.8 4.1 ± 0.4 0.9 ± 0.1 3.0 ± 0.4 1.8 ± 0.1

n-Octadecanoic acid 1 83.6 ± 2.8 37.3 ± 0.1 75.5 ± 0.8 45.9 ± 0.3 102.5 ± 15.4 32.8 ± 0.5 146.4 ± 5.5 58.7 ± 3.7

127

2 53.3 ± 2.6 22.7 ± 0.4 71.4 ± 11.3 44.9 ± 0.5 105.9 ± 2.6 19.6 ± 0.2 61.9 ± 0.1 26.9 ± 2.6

3 121.7 ± 12.0 23.6 ± 0.9 138.5 ± 0.6 15.4 ± 0.5 75.9 ± 8.4 15.8 ± 0.1 128.6 ± 16.5 NA

4 103.0 ± 5.3 24.4 ± 0.2 32.1 ± 0.5 39.5 ± 1.7 27.9 ± 0.1 13.1 ± 2.3 25.4 ± 2.5 19.4 ± 0.4

n-Nonadecanoic acid 1 48.3 ± 2.5 32.0 ± 2.1 21.6 ± 0.3 28.5 ± 1.0 19.3 ± 2.3 24.5 ± 0.6 31.5 ± 1.7 48.1 ± 3.4

2 23.5 ± 1.1 20.0 ± 0.5 19.1 ± 1.7 48.0 ± 0.1 10.9 ± 0.5 20.2 ± 0.6 20.0 ± 0.3 25.5 ± 2.5

3 41.8 ± 4.2 25.6 ± 1.2 30.5 ± 0.1 12.8 ± 0.1 16.7 ± 0.3 17.8 ± 0.2 27.9 ± 3.8 NA

4 11.6 ± 2.5 17.3 ± 1.1 8.2 ± 0.3 19.6 ± 0.9 9.1 ± 0.1 10.6 ± 0.6 11.3 ± 0.6 21.2 ± 0.7

n-Eicosanoic acid 1 156.3 ± 9.2 164.0 ± 6.7 110.4 ± 1.5 138.4 ± 1.4 98.7 ± 14.6 156.1 ± 0.7 188.2 ± 10.6 200.1 ± 8.8

2 101.4 ± 2.5 102.9 ± 4.1 91.5 ± 6.8 160.8 ± 1.1 107.1 ± 3.1 99.8 ± 0.1 143.9 ± 6.0 148.1 ± 13.1

3 199.3 ± 25.5 124.6 ± 5.6 137.8 ± 1.4 86.2 ± 1.0 96.0 ± 15.0 107.1 ± 0.1 198.8 ± 29.1 NA

4 89.9 ± 5.7 78.6 ± 4.2 43.5 ± 2.9 88.9 ± 2.7 62.5 ± 0.3 54.1 ± 2.9 70.2 ± 1.8 83.1 ± 2.4

n-Heneicosanoic acid 1 75.4 ± 2.0 74.5 ± 2.5 75.2 ± 0.2 90.8 ± 1.2 79.5 ± 12.8 82.9 ± 2.6 88.9 ± 4.9 125.0 ± 4.9

2 64.4 ± 3.2 51.9 ± 1.9 50.2 ± 0.8 96.7 ± 0.1 37.2 ± 1.2 50.6 ± 0.2 56.6 ± 2.5 71.4 ± 7.0

3 104.9 ± 13.5 62.9 ± 2.0 67.6 ± 0.6 37.9 ± 0.8 65.3 ± 11.1 52.0 ± 0.4 68.9 ± 6.4 NA

4 84.7 ± 5.6 38.5 ± 2.3 28.2 ± 0.5 73.0 ± 2.2 29.6 ± 0.5 28.7 ± 1.7 33.6 ± 0.9 46.8 ± 1.7

n-Docosanoic acid 1 303.3 ± 16.8 270.0 ± 4.4 240.8 ± 2.0 274.9 ± 3.4 399.5 ± 69.0 310.0 ± 0.2 483.0 ± 24.9 355.4 ± 9.6

2 210.4 ± 5.9 191.3 ± 7.0 211.3 ± 10.9 267.3 ± 0.5 270.1 ± 4.3 186.0 ± 0.1 360.6 ± 17.4 278.3 ± 21.5

3 416.7 ± 53.2 220.7 ± 6.6 338.6 ± 0.8 206.3 ± 1.3 359.3 ± 46.2 191.9 ± 1.7 524.4 ± 84.2 NA

4 555.9 ± 21.9 144.1 ± 8.8 152.7 ± 1.8 231.0 ± 11.6 159.7 ± 0.7 118.6 ± 6.6 179.2 ± 4.8 159.8 ± 5.6

n-Tricosanoic acid 1 97.8 ± 2.3 107.5 ± 5.3 155.2 ± 1.5 151.8 ± 1.6 220.6 ± 29.3 146.0 ± 1.0 176.3 ± 4.7 194.1 ± 9.8

2 125.6 ± 7.0 84.9 ± 3.1 97.2 ± 2.1 148.7 ± 1.2 84.0 ± 3.6 80.4 ± 2.7 94.3 ± 6.1 118.2 ± 12.6

3 185.8 ± 26.2 94.9 ± 4.4 124.1 ± 1.0 66.1 ± 0.8 186.8 ± 24.0 70.5 ± 0.2 259.9 ± 31.2 NA

4 229.4 ± 12.3 58.2 ± 4.5 78.3 ± 5.8 168.7 ± 7.6 96.9 ± 15.8 50.3 ± 3.3 79.1 ± 13.7 72.8 ± 4.5

n-Tetracosanoic acid 1 304.3 ± 15.9 249.0 ± 1.2 339.7 ± 1.2 297.7 ± 6.8 526.1 ± 85.2 323.0 ± 13.8 571.9 ± 25.7 380.2 ± 32.8

2 247.6 ± 7.3 202.3 ± 7.3 271.1 ± 16.7 295.6 ± 2.9 215.4 ± 4.8 206.6 ± 4.7 374.5 ± 28.5 304.7 ± 37.1

3 464.7 ± 59.9 234.7 ± 6.6 385.0 ± 33.4 192.7 ± 6.7 662.0 ± 83.6 212.4 ± 1.9 544.4 ± 78.1 NA

4 993.7 ± 47.9 162.4 ± 13.5 205.6 ± 3.5 297.1 ± 6.6 184.2 ± 2.3 146.0 ± 7.0 192.5 ± 2.2 189.8 ± 11.4

n-Pentacosanoic acid 1 48.5 ± 1.1 69.0 ± 3.9 137.5 ± 0.7 118.0 ± 1.5 143.5 ± 24.8 116.4 ± 0.2 125.6 ± 7.8 149.2 ± 11.2

2 103.5 ± 2.4 59.5 ± 1.1 57.1 ± 1.9 109.6 ± 1.2 58.8 ± 2.8 63.5 ± 2.5 64.0 ± 3.9 87.7 ± 7.0

3 106.3 ± 18.9 65.1 ± 2.8 84.5 ± 0.6 54.2 ± 0.1 106.4 ± 12.4 48.3 ± 0.8 103.3 ± 8.2 NA

4 148.5 ± 15.9 39.4 ± 3.6 47.2 ± 2.4 154.1 ± 13.6 47.8 ± 3.1 40.4 ± 3.3 38.8 ± 1.2 56.0 ± 3.0

n-Hexacosanoic acid 1 106.8 ± 6.4 135.5 ± 7.9 254.1 ± 1.1 202.9 ± 2.4 413.2 ± 62.9 211.4 ± 0.4 515.8 ± 21.8 250.5 ± 10.5

2 199.0 ± 5.8 112.9 ± 7.0 159.4 ± 13.5 186.9 ± 5.1 238.1 ± 2.0 136.4 ±6.0 258.1 ± 28.0 184.6 ±21.6

3 263.5 ± 41.2 127.1 ± 2.7 308.8 ± 18.3 110.3 ± 0.3 321.9 ± 26.2 112.0 ± 4.1 637.7 ± 95.1 NA

4 500.8 ± 47.1 88.3 ± 9.1 165.9 ± 3.5 215.5 ± 18.9 114.9 ± 1.1 98.1 ± 6.9 106.6 ± 2.6 122.7 ± 4.3

n-Heptacosanoic acid 1 19.8 ± 0.9 31.7 ± 2.9 107.9 ± 1.6 77.3 ± 1.7 85.0 ± 9.5 82.3 ± 16.7 130.6 ± 6.9 92.3 ± 13.7

2 67.0 ± 1.5 31.1 ± 0.2 37.5 ± 6.0 70.8 ± 10.3 51.9 ± 4.5 39.6 ± 0.8 43.8 ± 5.6 52.8 ± 3.2

3 54.4 ± 8.7 40.4 ± 4.0 74.7 ± 9.3 32.0 ± 3.1 43.5 ± 4.5 33.2 ± 2.4 91.9 ± 8.1 NA

4 56.8 ± 5.7 18.3 ± 1.6 32.8 ± 2.8 84.6 ± 5.2 28.4 ± 0.7 25.9 ± 2.6 27.7 ± 0.1 32.7 ± 2.9

n-Octacosanoic acid 1 65.6 ± 2.6 66.4 ± 2.7 230.9 ± 3.4 174.0 ± 8.6 279.1 ± 37.9 131.5 ± 1.0 555.8 ± 8.0 198.6 ± 6.5

2 171.6 ± 4.9 80.0 ± 9.5 126.2 ± 19.3 141.6 ± 5.9 269.0 ± 10.6 90.4 ± 2.1 254.0 ± 39.6 124.7 ± 14.7

128

NA = not analyzed (the lower horizon soil sample for the warming +N+P fertilization treatment of block 3 was unavailable) aTotal aliphatic compounds = total n-alkanes + total n-alkanols + total n-alkanoic acids (Otto and Simpson, 2005)

3 140.3 ± 3.0 82.1 ± 8.2 371.3 ± 36.3 71.9 ± 3.4 135.8 ± 1.3 66.1 ± 2.5 817.3 ± 145.5 NA

4 220.2 ± 17.8 42.6 ± 3.1 100.1 ± 9.4 188.6 ± 15.7 84.3 ± 0.4 68.1 ± 5.6 69.9 ± 3.8 79.7 ± 6.5

n-Nonacosanoic acid 1 33.4 ± 1.6 24.9 ± 4.1 103.8 ± 4.1 56.5 ± 1.4 120.2 ± 18.8 52.1 ± 3.5 148.7 ± 11.4 66.1 ± 10.6

2 53.9 ± 3.8 21.3 ± 1.3 43.5 ± 2.9 52.2 ± 1.6 32.2 ± 0.3 27.8 ± 1.9 44.4 ± 3.5 35.9 ± 5.4

3 57.3 ± 11.6 25.9 ± 4.6 70.6 ± 14.0 22.6 ± 0.8 59.0 ± 7.2 22.6 ± 1.3 207.3 ± 33.1 NA

4 78.7 ± 10.2 17.6 ± 2.5 28.4 ± 3.1 81.7 ± 3.0 28.7 ± 0.1 17.0 ± 2.1 23.1 ± 0.3 22.4 ± 1.2

n-Triacontanoic acid 1 51.1 ± 6.1 38.8 ± 2.0 159.9 ± 3.8 80.2 ± 5.3 196.7 ± 8.0 49.6 ± 0.5 342.4 ± 40.7 77.9 ± 8.1

2 146.3 ± 3.0 32.2 ± 5.3 81.1 ± 10.7 75.8 ± 7.1 228.1 ± 0.1 48.4 ± 3.0 164.1 ± 16.9 64.7 ± 7.4

3 102.0 ± 13.7 44.7 ± 6.4 306.4 ± 31.8 35.4 ± 0.9 161.3 ± 20.6 37.0 ± 2.0 1418.1 ± 215.2 NA

4 150.5 ± 18.2 28.6 ± 2.2 76.0 ± 1.8 108.2 ± 3.1 57.6 ± 2.6 31.9 ± 1.0 43.9 ± 2.1 40.6 ± 1.4

n-Hentriacontanoic acid 1 24.6 ± 0.1 11.9 ± 1.5 34.1 ± 2.2 32.7 ± 3.3 52.4 ± 5.6 23.5 ± 2.8 64.3 ± 7.2 44.5 ± 0.1

2 23.3 ± 3.8 13.1 ± 0.3 21.0 ± 0.4 19.6 ± 1.4 40.8 ± 2.5 15.6 ± 1.6 34.5 ± 2.1 20.8 ± 2.0

3 34.2 ± 2.0 13.6 ± 0.9 78.3 ± 8.6 11.1 ± 0.2 28.1 ± 1.2 14.6 ± 0.7 106.0 ± 8.7 NA

4 56.5 ± 0.7 10.3 ± 1.0 26.2 ± 1.4 34.3 ± 1.4 20.2 ± 1.9 7.7 ± 0.3 18.8 ± 2.6 11.7 ± 2.7

Short-chain vs. long-chain n-alkanoic acids

Short-chain C15-C19 1 402.7 ± 4.7 140.4 ± 2.6 408.4 ± 3.1 204.2 ± 1.9 474.5 ± 32.5 142.4 ± 3.1 831.1 ± 22.2 263.1 ± 6.9

Long-chain C20-C32 1286.7 ± 26.8 1243.3 ± 14.7 1949.4 ± 7.9 1695.0 ± 13.8 2614.5 ± 140.7 1684.9 ± 86.0 3391.5 ± 62.6 2134.01 ± 44.8

Short-chain C15-C19 2 301.4 ± 6.2 77.6 ± 0.9 371.0 ± 19.2 204.3 ± 1.2 508.8 ± 3.6 81.6 ± 1.3 306.4 ± 3.7 104.0 ± 4.5

Long-chain C20-C32 1514.06 ± 16.0 983.3 ± 17.4 1247.1 ± 34.1 1625.6 ± 15.3 1632.7 ± 14.7 1045.2 ± 9.6 1892.8 ± 62.4 1491.9 ± 55.2

Short-chain C15-C19 3 498.0 ± 21.5 99.0 ± 1.9 1439.4 ± 21.2 63.0 ± 0.7 877.1 ± 27.3 66.2 ± 0.4 720.3 ± 43.3 NA

Long-chain C20-C32 2129.6 ± 101.9 1012.2 ± 17.4 2347.6 ± 64.3 926.9 ± 8.5 2225.3 ± 106.7 967.8 ± 6.5 4978.05 ± 304.8 NA

Short-chain C15-C19 4 686.7 ± 23.1 90.0 ± 1.4 300.4 ± 4.3 255.3 ± 4.6 151.2 ± 3.6 43.2 ± 2.8 108.9 ± 3.1 75.4 ± 1.0

Long-chain C20-C32 3165.6 ± 79.1 648.2 ± 20.6 984.9 ± 13.7 1725.7 ± 32.9 914.8 ± 16.7 686.7 ± 14.7 883.6 ± 15.9 918.3 ± 16.7

Total n-alkanoic acids

1 1689.4 ± 27.3 1383.7 ± 14.9 2357.8 ± 8.4 1899.2 ± 14.0 3089.0 ± 144.4 1827.3 ± 86.1 4222.6 ± 66.4 2397.1 ± 45.3

2 1815.5 ± 17.2 1060.9 ± 17.4 1618.1 ± 39.1 1829.8 ± 15.4 2141.5 ± 15.1 1126.8 ± 9.6 2199.2 ± 62.5 1595.9 ± 55.4

3 2627.6 ± 104.2 1235.7 ± 17.5 3786.9 ± 67.7 989.8 ± 8.5 2540.9 ± 110.1 1033.9 ± 6.5 5698.3 ± 307.8 NA

4 3976.5 ± 82.4 816.8 ± 20.6 1285.4 ± 14.3 1981.0 ± 33.2 1065.9 ± 17.1 729.9 ± 15.0 992.5 ± 16.2 993.7 ± 16.8

Total aliphatic compoundsa

1 2825.8 ± 36.4 2137.9 ± 19.3 4226.2 ± 39.3 3291.6 ± 17.4 4957.2 ± 151.7 2788.5 ± 105.8 7625.3 ± 89.9 3929.9 ± 55.8

2 3156.0 ± 48.9 1610.2 ± 18.6 2647.3 ± 52.6 2741.1 ± 17.6 4050.9 ± 65.0 1703.2 ± 13.6 3629.6 ± 81.2 2469.7 ± 77.7

3 4175.8 ± 115.8 2045.2 ± 21.7 6907.7 ± 102.4 1569.9 ± 10.5 3963.4 ± 116.3 1594.7 ± 8.6 9462.1 ± 356.9 NA

4 5666.4 ± 90.7 1212.4 ± 22.3 2299.7 ± 19.0 3484.7 ± 44.6 1812.3 ± 24.6 1079.6 ± 15.7 1595.3 ± 21.3 1487.5 ± 21.8

129


soil) of major compound classes identified in the total solvent extracts (excluding aliphatic compounds) of

the upper and lower horizon soils of the control, warming, N+P fertilization and warming +N+P fertilization treatments. All values are

reported as mean ± standard error (n = 2). Numbers in bold denote statistical significance from the control treatment (P < 0.05).

Block Treatment



Simple carbohydrates

Glucose 1 132.3 ± 3.3 9.1 ± 0.9 147.8 ± 0.8 40.7 ± 1.4 163.7 ± 42.1 8.4 ± 1.5 607.2 ± 55.6 43.6 ± 1.1

2 193.9 ± 1.7 1.8 ± 0.3 220.1 ± 2.9 29.1 ± 0.5 55.8 ± 0.5 4.2 ± 2.4 187.3 ± 12.3 2.6 ± 0.5

3 292.2 ± 55.4 4.7 ± 0.2 795.8 ± 133.9 29.1 ± 2.1 1350.9 ± 172.0 10.0 ± 0.9 109.8 ± 15.4 NA

4 743.3 ± 14.7 32.2 ± 2.6 ND 127.7 ± 2.2 36.0 ± 8.6 2.8 ± 0.1 28.6 ± 2.1 4.7 ± 0.2

Galactose 1 155.7 ± 1.6 3.3 ± 0.5 64.1 ± 4.1 11.7 ± 0.6 44.5 ± 8.4 5.6 ± 0.2 380.7 ± 5.7 20.1 ± 0.9

2 56.3 ± 2.1 1.0 ± 0.1 109.5 ± 5.7 6.5 ± 0.2 32.3 ± 0.2 3.1 ± 0.8 49.4 ± 0.1 1.7 ± 0.3

3 54.9 ± 0.6 1.6 ± 0.2 216.3 ± 34.0 4.1 ± 0.3 996.3 ± 109.7 3.2 ± 0.1 41.6 ± 4.3 NA

4 236.2 ± 15.3 7.7 ± 1.2 30.0 ± 1.2 27.9 ± 0.3 8.5 ± 1.2 1.2 ± 0.1 4.7 ± 0.2 1.9 ± 0.1

Mannose 1 152.2 ± 6.2 11.0 ± 1.0 160.9 ± 2.3 45.1 ± 1.1 187.4 ± 37.8 9.2 ± 1.8 518.6 ± 29.3 48.0 ± 1.3

2 191.1 ± 1.3 2.1 ± 0.2 164.3 ± 11.5 37.2 ± 0.1 44.8 ± 2.0 8.2 ± 4.4 145.6 ± 26.8 3.7 ± 0.3

3 290.2 ± 75.6 6.3 ± 0.4 628.5 ± 129.9 33.0 ± 3.0 1670.5 ± 282.1 11.4 ± 1.3 120.1 ± 20.0 NA

4 542.7 ± 2.4 34.1 ± 2.5 ND 145.8 ± 3.2 34.8 ± 0.7 4.0 ± 0.4 25.9 ± 1.8 5.3 ± 0.1

Total simple carbohydrates

1 440.3 ± 7.2 23.4 ± 1.4 372.8 ± 4.7 97.4 ± 1.8 395.6 ± 57.1 23.2 ± 3.1 1506.4 ± 63.1 111.5 ± 1.9

2 441.3 ± 2.9 4.9 ± 0.3 493.9 ± 13.1 72.9 ± 0.6 132.9 ± 2.0 18.0 ± 5.1 382.3 ± 29.5 7.9 ± 0.7

3 637.3 ± 93.8 12.6 ± 0.5 1640.6 ± 189.6 66.2 ± 3.7 4017.7 ± 348.2 24.5 ± 1.6 271.5 ± 25.6 NA

4 1522.4 ± 21.4 74.0 ± 3.7 220.1 ± 119.1 301.4 ± 3.9 79.3 ± 8.7 7.9 ± 0.5 59.2 ± 2.8 11.9 ± 0.2

Cyclic compounds

Plant Steroids

Cholesterol 1 28.1 ± 0.8 11.4 ± 1.9 23.4 ± 2.6 8.8 ± 0.8 47.8 ± 5.9 15.6 ± 1.7 67.4 ± 4.5 ND

2 30.0 ± 0.1 6.7 ± 0.2 38.5 ± 0.3 9.9 ± 0.7 60.4 ± 2.7 3.8 ± 0.1 33.2 ± 2.0 ND

3 46.9 ± 1.5 5.5 ± 0.2 81.7 ± 1.1 3.2 ± 0.5 52.3 ± 3.6 2.5 ± 0.4 110.0 ± 10.4 NA

4 53.9 ± 5.5 5.7 ± 0.1 21.7 ± 2.6 19.7 ± 1.6 28.9 ± 0.1 4.2 ± 0.5 20.9 ± 1.9 ND

Campesterol 1 56.6 ± 1.1 47.9 ± 6.7 58.4 ± 3.7 86.1 ± 0.9 154.3 ± 24.3 102.0 ± 0.6 140.6 ± 18.5 46.8 ± 10.6

2 28.0 ± 1.6 10.7 ± 0.1 128.8 ± 22.2 21.1 ± 0.8 102.0 ± 10.6 14.1 ± 0.2 73.8 ± 1.7 20.7 ± 1.0

3 123.0 ± 23.7 23.0 ± 1.9 117.7 ± 0.2 9.4 ± 1.1 213.3 ± 29.2 21.3 ± 1.1 133.8 ± 16.6 NA

4 211.0 ± 81.8 26.7 ± 2.8 47.1 ± 4.2 22.8 ± 0.1 33.4 ± 4.0 7.5 ± 0.5 57.1 ± 0.5 5.0 ± 0.6

Ergosterol 1 ND ND ND ND ND ND ND ND

2 ND ND ND ND ND ND ND ND



Stigmasterol 1 51.4 ± 1.4 25.6 ± 1.7 87.1 ± 5.4 59.8 ± 3.4 99.2 ± 14.5 42.9 ± 1.6 144.3 ± 3.4 23.0 ± 4.4

2 48.2 ± 1.9 2.5 ± 0.1 65.8 ± 0.3 11.4 ± 1.4 143.1 ± 7.6 5.6 ± 0.7 76.5 ± 0.1 6.9 ± 1.7

130

3 79.0 ± 8.8 11.7 ± 0.4 157.2 ± 8.6 1.9 ± 0.1 103.1 ± 14.7 3.1 ± 0.2 319.7 ± 22.5 NA

4 101.2 ± 7.8 7.8 ± 0.1 42.7 ± 2.5 18.6 ± 0.8 42.4 ± 2.9 10.9 ± 0.5 32.9 ± 5.1 4.7 ± 0.1

β-Sitosterol 1 241.1 ± 3.9 181.6 ± 9.8 306.8 ± 14.4 300.0 ± 0.3 614.6 ± 97.4 289.6 ± 3.8 1097.9 ± 75.0 261.4 ± 5.3

2 204.1 ± 17.6 55.0 ± 0.1 631.6 ± 32.8 67.4 ± 1.1 1247.2 ± 34.9 54.1 ±0.2 610.2 ± 34.8 92.4 ± 6.2

3 526.7 ± 52.8 71.9 ± 2.8 1133.4 ± 8.0 47.8 ± 0.5 1194.9 ± 141.0 62.1 ± 0.9 1318.7 ± 152.2 NA

4 1778.9 ± 103.2 98.7 ± 2.1 278.2 ± 11.8 137.9 ± 2.1 333.5 ± 9.4 19.8 ± 1.5 192.0 ± 8.7 32.0 ± 0.1

Stigmasta-3,5-dien-7one 1 65.2 ± 1.6 32.7 ± 4.9 89.9 ± 32.0 80.7 ± 1.8 114.9 ± 12.0 63.9 ± 4.6 255.2 ± 22.9 81.2 ± 5.2

2 56.8 ± 14.1 13.9 ± 2.1 83.3 ± 5.1 18.8 ± 0.4 190.3 ± 14.7 9.7 ± 0.5 168.3 ± 22.2 18.7 ± 2.1

3 100.1 ± 8.5 14.1 ± 0.1 304.7 ± 17.2 10.7 ± 1.0 140.7 ± 18.6 12.7 ± 1.2 781.9 ± 101.7 NA

4 141.3 ± 9.7 11.3 ± 0.1 48.9 ± 4.2 51.2 ± 5.2 52.4 ± 3.5 6.1 ± 0.5 55.9 ± 4.6 11.7 ± 0.5

Sitosterone 1 50.7 ± 3.8 30.9 ± 3.0 130.7 ± 1.1 58.8 ± 9.0 148.9 ± 20.7 74.4 ± 3.7 220.3 ± 12.1 ND

2 85.8 ± 2.6 17.2 ± 0.1 89.8 ± 2.9 26.4 ± 0.3 257.7 ± 3.3 19.4 ± 0.2 119.8 ± 5.8 ND

3 73.7 ± 10.1 12.5 ± 0.9 187.8 ± 10.7 13.2 ± 0.8 120.4 ± 8.5 14.5 ± 0.1 502.4 ± 108.7 NA

4 124.1 ± 13.7 16.0 ± 1.4 53.7 ± 9.3 44.4 ± 1.4 57.5 ± 6.5 8.6 ± 1.1 49.2 ± 3.3 ND

Total plant steroids

1 493.0 ± 6.0 330.1 ± 13.4 696.3 ± 35.7 594.3 ± 9.9 1179.7 ± 104.4 588.3 ± 14.5 1925.6 ± 81.7 412.3 ± 13.7

2 453.0 ± 22.8 106.0 ± 2.1 1037.8 ± 40.0 155.1 ± 2.2 2000.7 ± 40.3 106.6 ± 1.0 1081.6 ± 41.7 138.8 ± 6.8

3 949.5 ± 60.1 138.7 ± 3.5 1982.5 ± 23.5 86.2 ± 1.8 1824.7 ± 146.2 116.2 ± 1.9 3166.6 ± 215.0 NA

4 2410.4 ± 133.1 166.3 ± 3.8 492.3 ± 16.6 294.7 ± 6.1 548.2 ± 12.9 57.1 ± 2.1 408.1 ± 11.7 53.3 ± 0.8

Triterpenoids

Oleanolic acid 1 54.4 ± 2.3 51.3 ± 3.6 292.2 ± 7.7 109.4 ± 0.4 226.1 ± 30.1 80.3 ± 2.9 598.3 ± 47.2 130.0 ± 0.7

2 233.8 ± 5.5 31.7 ± 1.4 247.6 ± 17.9 46.4 ± 0.9 319.4 ± 0.7 25.8 ± 2.4 198.8 ± 10.3 32.4 ± 2.0

3 113.9 ± 10.5 28.7 ± 3.6 887.7 ± 11.8 21.9 ± 0.6 166.0 ± 21.3 24.1 ± 1.3 1116.4 ± 185.4 NA

4 111.5 ± 8.7 25.9 ± 0.6 184.0 ± 21.0 76.6 ± 0.1 150.4 ± 1.3 12.4 ± 0.9 76.7 ± 0.2 14.4 ± 0.4

Ursolic acid 1 150.7 ± 2.2 150.6 ± 5.4 804.6 ± 12.5 325.7 ± 0.5 842.5 ± 127.2 233.3 ± 6.2 1950.2 ± 124.2 353.7 ± 2.3

2 597.8 ± 19.7 114.8 ± 4.8 758.1 ± 71.1 154.5 ± 7.0 833.1 ± 12.6 86.5 ± 2.2 616.1 ± 15.8 108.5 ± 3.5

3 343.3 ± 48.8 79.0 ± 6.5 3059.7 ± 35.0 63.2 ± 0.7 579.8 ± 86.0 66.0 ± 0.9 2626.0 ± 379.0 NA

4 349.6 ± 40.5 78.7 ± 3.1 711.4 ± 104.2 238.0 ± 1.0 540.3 ± 4.6 38.9 ± 1.9 224.8 ± 8.3 41.8 ± 1.7

Total triterpenoids

1 205.1± 3.2 202.0 ± 6.5 1096.6 ± 14.6 435.1 ± 0.6 1068.6 ± 130.7 313.7 ± 16.1 2548.5 ± 132.9 483.7 ± 2.4

2 831.7 ± 20.5 146.5 ± 5.0 1005.7 ± 73.3 200.9 ± 7.0 1152.5 ± 177.9 112.2 ± 3.3 814.9 ± 18.8 140.9 ± 4.0

3 457.3 ± 49.9 107.8 ± 7.5 3947.4 ± 37.0 85.1 ± 0.9 745.9 ± 88.6 90.2 ±1.6 3742.5 ± 421.9 NA

4 461.0 ± 41.4 104.7 ± 3.1 895.4 ± 106.3 314.6 ± 1.0 690.6 ± 4.8 51.3 ± 2.1 301.5 ± 8.3 56.2 ± 1.7

Total cyclic compoundsb

1 698.1 ± 6.8 532.0 ± 14.9 1793.0 ± 38.6 1029.4 ± 9.9 2248.2 ± 167.3 902.0 ± 21.7 4474.1 ± 156.0 896.0 ± 13.9

2 1284.7 ± 30.6 252.5 ± 5.5 2043.5.2 ± 83.5 356.0 ± 7.3 3153.3 ± 42.2 218.8 ± 3.4 1896.6 ± 45.8 279.7 ± 7.9

3 1406.7 ± 78.1 246.5 ± 8.3 5929.9 ± 43.8 171.3 ± 2.1 2570.5 ± 171.0 206.3 ± 2.4 6909.0 ± 473.5 NA

4 2871.4 ± 139.4 271.0 ± 4.9 1387.8 ± 107.6 609.3 ± 6.2 1238.9 ± 13.8 108.4 ± 3.0 709.6 ± 14.4 109.5 ± 1.9

131

NA = not analyzed (the lower horizon soil sample for the warming +N+P fertilization treatment of block 3 was unavailable); ND = not detected bTotal cyclic compounds = total steroids + total triterpenoids (Otto and Simpson, 2005) cAliphatic/cyclic ratio = total aliphatic compounds/total cyclic compounds (Otto and Simpson, 2005)

Aliphatic/ cyclic ratioc

1 4.0 ± 0.1 4.0 ± 0.1 2.4 ±0.1 3.2 ± 0.1 2.2 ± 0.2 3.1 ± 0.1 1.7 ± 0.1 4.4 ± 0.1

2 2.5 ± 0.1 6.4 ± 0.2 1.3 ± 0.1 7.7 ± 0.2 1.3 ± 0.1 7.8 ± 0.1 1.9 ± 0.1 8.8 ± 0.4

3 3.0 ± 0.2 8.3 ± 0.3 1.2 ± 0.1 9.2 ± 0.1 1.5 ± 0.1 7.7 ± 0.1 1.4 ± 0.1 NA

4 2.0 ± 0.1 4.5 ± 0.1 1.7 ± 0.1 5.7 ± 0.1 1.5 ± 0.1 10.0 ± 0.3 2.2 ± 0.1 13.6 ± 0.3

132


soil) of major SOM components released from the base hydrolysis of the upper and lower horizon soils of

the control, warming, N+P fertilization and warming +N+P fertilization treatments. All values are reported as mean ± standard error (n = 2).

Numbers in bold denote statistical significance from the control treatment (P < 0.05).

Block Treatment



Benzyls 1 141.4 ± 37.4 45.5 ± 22.4 146.0 ± 10.4 88.9 ± 10.6 183.5 ± 10.3 29.3 ± 3.8 184.9 ± 18.3 75.0 ± 12.8

2 92.3 ± 2.9 42.7 ± 1.4 121.1 ± 3.2 26.6 ± 1.0 142.2 ± 5.4 21.8 ± 0.4 114.7 ± 11.6 15.7 ± 1.8

3 198.1 ± 8.4 79.3 ± 4.4 265.7 ± 121.9 20.8 ± 1.1 202.5 ± 13.5 21.9 ± 0.6 155.9 ± 19.8 NA

4 123.5 ± 5.5 60.2 ± 2.9 144.0 ± 6.9 114.5 ± 3.8 129.6 ± 10.9 20.9 ± 0.4 53.2 ± 3.9 17.8 ± 2.1

Phenols

Vanillin 1 7.6 ± 2.1 32.6 ± 11.4 9.7 ± 2.2 11.9 ± 0.5 19.5 ± 1.4 6.3 ± 0.9 10.9 ± 2.0 6.4 ± 1.2

2 16.5 ± 1.9 8.6 ± 0.2 57.6 ± 4.3 10.6 ± 0.9 24.5 ± 4.5 9.2 ± 1.5 22.0 ± 0.3 5.5 ± 2.3

3 51.6 ± 5.6 13.7 ± 0.1 31.5 ± 5.9 7.2 ± 0.8 79.6 ± 4.7 9.5 ± 0.6 25.9 ± 5.1 NA

4 50.0 ± 2.1 11.6 ± 0.4 11.5 ± 4.9 11.3 ± 2.7 22.4 ± 3.3 6.8 ± 0.3 12.8 ± 0.5 7.4 ± 0.1

Acetovanillone 1 4.6 ± 2.2 3.8 ± 2.3 7.8 ± 0.2 7.2 ± 0.1 19.6 ± 0.8 2.0 ± 0.4 6.9 ± 0.7 5.4 ± 0.8

2 12.3 ± 3.2 10.5 ± 1.0 10.7 ± 0.9 9.0 ± 0.1 11.4 ± 0.5 3.5 ± 0.2 7.7 ± 0.8 2.2 ± 0.5

3 16.5 ± 7.1 16.5 ± 0.8 12.8 ± 1.7 5.7 ± 0.1 23.4 ± 2.9 4.7 ± 0.9 8.9 ± 0.2 NA

4 29.7 ± 1.5 7.5 ± 0.9 11.2 ± 0.9 31.2 ± 5.9 9.3 ± 2.3 3.7 ± 0.3 5.7 ± 1.1 4.7 ± 1.1

Vanillic acid 1 197.8 ± 18.1 88.4 ± 43.5 203.8 ± 3.5 181.7 ± 19.6 364.5 ± 8.7 66.1 ± 12.4 179.4 ± 9.0 125.9 ± 19.2

2 94.5 ± 5.7 102.9 ± 5.6 129.9 ± 1.8 56.6 ± 0.1 115.2 ± 15.0 34.6 ± 0.4 82.3 ± 6.4 31.4 ± 1.7

3 185.0 ± 3.1 110.3 ± 5.2 186.7 ± 4.4 36.1 ± 1.2 311.7 ± 17.8 38.6 ± 1.6 104.8 ± 12.6 NA

4 232.5 ±11.3 110.7 ± 0.6 116.9 ± 9.8 241.0 ± 12.9 101.1 ± 9.0 27.2 ± 0.9 64.9 ± 7.5 36.1 ± 0.2

Syringic acid 1 110.2 ± 6.3 32.2 ± 13.2 79.0 ± 1.5 73.7 ± 9.8 138.4 ± 13.5 24.6 ± 4.9 70.4 ± 5.4 52.0 ± 10.6

2 37.2 ± 4.5 30.6 ± 1.6 47.8 ± 14.9 19.3 ± 0.9 57.3 ± 0.5 11.1 ± 0.3 34.7 ± 1.6 10.4 ± 0.5

3 118.4 ± 0.1 43.0 ± 2.2 71.4 ± 0.6 11.9 ± 0.6 123.4 ± 10.0 15.3 ± 0.7 60.7 ± 4.7 NA

4 88.6 ± 4.5 45.4 ± 2.2 38.2 ± 0.3 88.2 ± 2.4 49.7 ± 3.3 8.8 ± 0.3 23.5 ± 2.6 12.0 ± 1.1

p-coumaric acid 1 361.2 ± 88.5 106.0 ± 66.9 221.2 ± 8.2 193.2 ± 27.1 331.8 ± 0.2 92.5 ± 17.1 393.0 ± 35.2 138.7 ± 21.1

2 123.4 ± 9.4 83.7 ± 3.8 645.3 ± 19.6 41.2 ± 0.2 378.1 ± 6.8 33.5 ± 1.3 230.8 ± 7.1 27.3 ± 0.4

3 612.8 ± 23.9 138.8 ± 7.9 372.2 ± 14.1 34.7 ± 2.0 614.4 ± 5.1 39.9 ± 2.6 483.6 ± 37.8 NA

4 1137.1 ± 9.1 143.0 ± 1.2 151.2 ± 1.3 162.2 ± 5.2 217.4 ± 18.5 26.9 ± 2.2 53.8 ± 7.5 27.1 ± 0.6

Ferulic acid 1 439.6 ± 97.0 313.3 ± 202.8 264.9 ± 6.5 359.2 ± 56.7 1118.6 ± 48.2 158.3 ± 40.1 336.7 ± 0.6 252.1 ± 36.8

2 184.4 ± 9.4 289.0 ± 5.7 768.8 ± 10.3 149.0 ± 2.0 259.7 ± 4.7 70.9 ± 2.2 186.4 ± 0.1 99.0 ± 3.6

3 616.6 ± 9.6 394.1 ± 7.7 230.5 ± 8.6 99.9 ± 0.4 921.3 ± 5.1 106.4 ± 5.1 224.7 ± 8.3 NA

4 1433.5 ± 19.2 319.6 ± 9.2 218.1 ± 0.6 487.4 ± 34.0 219.8 ± 9.4 61.7 ± 3.0 113.1 ± 15.8 63.5 ± 4.4

Total phenols

1 1126.2 ± 132.8 581.5 ± 218.7 793.2 ± 11.3 835.4 ± 66.6 1999.3 ± 50.8 353.3 ± 45.6 1010.4 ± 36.8 583.5 ± 47.8

2 476.7 ± 15.6 530.5 ± 9.3 1687.9 ± 27.8 285.6 ± 2.3 865.7 ± 17.9 167.8 ± 3.4 573.0 ± 9.9 178.5 ± 5.0

3 1620.8 ± 27.7 725.6 ± 12.8 877.0 ± 19.3 198.8 ± 2.6 2104.2 ± 22.4 215.9 ± 6.2 930.5 ± 41.4 NA

4 2988.2 ± 24.7 647.8 ± 9.7 553.8 ± 11.4 1021.4 ± 37.4 630.9 ± 23.3 139.2 ± 3.9 279.4 ± 19.2 152.7 ± 5.1

133

Steroids and terpenoids

β-sitosterol 1 51.0 ± 4.0 41.6 ± 21.0 20.4 ± 5.7 22.6 ± 0.9 91.3 ± 4.8 8.5 ± 2.2 25.9 ± 1.4 19.8 ± 8.4

2 29.4 ± 6.7 33.8 ± 6.3 31.7 ± 2.9 16.8 ± 1.5 65.0 ± 0.5 10.1 ± 1.4 40.3 ± 6.3 15.2 ± 1.2

3 35.4 ± 0.6 43.3 ± 12.8 60.9 ± 4.4 12.7 ± 0.5 81.9 ± 16.0 12.3 ± 0.3 77.2 ± 11.1 NA

4 110.0 ± 1.8 25.0 ± 7.1 27.0 ± 1.4 43.8 ± 5.4 27.6 ± 4.0 7.3 ± 0.3 13.0 ± 1.3 8.2 ± 2.3

Aliphatic lipids

α-Hydroxyalkanoic

acids C16-C28

1 370.2 ± 32.6 911.6 ± 63.4 355.9 ± 86.5 375.1 ± 33.7 1430.6 ± 96.8 99.4 ± 4.4 290.2 ± 5.3 355.2 ± 49.2

2 144.9 ± 5.8 428.5 ± 6.0 219.8 ± 2.9 192.1 ± 11.0 388.5 ± 20.3 107.3 ± 1.7 224.8 ± 4.7 157.8 ± 8.3

3 305.9 ± 21.8 572.0 ± 58.0 288.5 ± 17.1 127.8 ± 6.5 414.6 ± 4.5 140.2 ± 3.1 421.9 ± 35.1 NA

4 259.0 ± 8.6 193.5 ± 24.9 179.0 ± 1.7 430.8 ± 16.4 195.8 ± 12.0 78.7 ± 0.3 130.2 ± 7.5 90.6 ± 3.8

n-Alkanols

Microbial or plant

origin C16-C18

1 47.3 ± 13.8 57.8 ± 14.4 42.2 ± 3.1 28.2 ± 3.8 117.4 ± 13.7 13.7 ± 2.3 38.4 ± 1.3 33.7 ± 11.8

2 38.1 ± 8.2 34.6 ± 4.2 32.2 ± 3.4 28.8 ± 2.4 23.0 ± 0.9 13.8 ± 1.7 43.3 ± 0.7 15.7 ± 0.8

3 37.2 ± 6.1 50.1 ± 3.0 49.1 ± 3.0 20.9 ± 3.8 50.8 ± 15.9 14.9 ± 2.1 52.5 ± 2.1 NA

4 41.5 ± 4.8 15.8 ± 1.8 35.8 ± 4.7 50.6 ± 7.8 34.5 ± 4.6 10.5 ± 1.0 26.5 ± 1.1 12.0 ± 2.1

Suberin or plant waxes

C20-C30

1 464.3 ± 37.7 850.9 ± 54.5 397.5 ± 52.6 462.1 ± 38.3 1182.5 ± 56.6 157.2 ± 16.6 558.9 ± 24.4 357.9 ± 33.4

2 279.9 ± 9.6 399.9 ± 36.4 321.1 ± 28.4 273.6 ± 10.6 369.0 ± 47.3 144.4 ± 10.7 361.9 ± 12.4 178.3 ± 9.0

3 318.4 ± 34.4 534.9 ± 39.5 613.5 ± 19.6 174.0 ± 7.5 688.2 ± 84.2 166.1 ± 5.8 486.8 ± 27.1 NA

4 602.5 ± 94.8 268.8 ± 27.2 428.7 ± 68.7 745.4 ± 68.3 384.1 ± 10.2 113.7 ± 5.2 241.4 ± 27.0 137.9 ± 8.1

Total n-alkanols

1 511.7 ± 40.2 908.7 ± 56.3 439.6 ± 52.7 490.3 ± 38.5 1299.8 ± 58.2 170.8 ± 16.8 597.2 ± 24.5 391.6 ± 35.4

2 318.0 ± 12.6 434.4 ± 36.6 353.3 ± 28.6 302.4 ± 10.9 392.0 ± 47.3 158.2 ±10.8 405.1 ± 12.4 194.0 ± 9.1

3 355.6 ± 34.9 584.9 ± 39.6 662.6 ± 19.8 195.0 ± 8.4 739.0 ± 85.7 181.0 ± 6.2 539.3 ± 27.2 NA

4 644.0 ± 94.9 284.6 ± 27.2 464.5 ± 68.8 796.0 ± 68.7 418.6 ± 11.2 124.2 ± 5.3 267.9 ± 27.0 149.9 ± 8.3

n-Alkanoic acids

Microbial or plant

origin C16-C18

1 338.2 ± 71.4 526.2 ± 60.1 342.3 ± 11.6 460.8 ± 41.1 1149.8 ± 35.4 154.5 ± 11.4 554.9 ± 14.8 381.4 ± 51.0

2 423.0 ± 20.6 386.0 ± 13.2 679.5 ± 55.5 193.1 ± 3.1 739.2 ± 75.6 178.2 ± 10.3 634.5 ± 58.5 148.1 ± 17.4

3 681.0 ± 42.1 536.8 ± 30.3 1140.7 ± 25.2 156.9 ± 6.7 926.4 ± 49.2 184.6 ± 3.0 1169.8 ± 111.7 NA

4 488.8 ± 19.2 354.4 ± 11.0 298.3 ± 15.6 522.4 ± 21.1 425.6 ± 38.8 97.6 ± 8.4 318.5 ± 29.5 171.0 ± 8.5

Suberin, cutin or plant

waxes C20-C30

1 2942.8 ± 82.9 4768.6 ± 235.7 2167.6 ± 267.5 2774.4 ± 171.6 8953.6 ± 219.7 1063.5 ± 205.9 2295.4 ± 28.5 2230.0 ± 140.5

2 1187.7 ± 46.6 3245.5 ± 55.2 1697.0 ± 55.7 1470.1 ± 35.7 1070.5 ± 40.3 894.6 ± 17.9 1279.1 ± 26.0 1213.2 ± 25.7

3 1870.5 ± 89.6 4413.7± 151.8 1715.9 ± 35.2 973.8 ± 25.6 3022.7 ± 107.1 1132.0 ± 31.4 1803.3 ± 78.9 NA

4 2009.4 ± 34.8 1768.5 ± 61.2 1144.9 ± 39.4 3003.6 ± 222.3 1308.2 ± 36.6 652.2 ± 13.3 896.2 ± 29.1 626.7 ± 22.6

Total n-alkanoic acids

1 3280.9 ± 109.4 5294.8 ± 243.2 2509.9 ± 267.7 3235.2 ± 176.4 10103.4 ±

222.6

1218.0 ± 206.2 2850.3 ± 32.1 2611.4 ± 149.5

2 1610.7 ± 50.9 3631.4 ± 56.7 2376.4 ± 78.7 1663.2 ± 35.8 1809.7 ± 85.7 1072.8 ± 20.6 1913.6 ± 64.0 1361.3 ± 31.0

3 2551.5 ± 99.0 4950.5 ± 154.8 2856.6 ± 43.3 1130.7 ± 26.4 3949.1 ± 117.9 1316.6 ± 31.5 2973.1 ± 136.8 NA

4 2498.2 ± 39.7 2122.8 ± 62.2 1443.2 ± 42.3 3526.0 ± 223.3 1733.8 ± 53.3 749.8 ± 15.7 1214.7 ± 41.4 797.6 ± 24.2

134

n-Alkane α,ω-dioic acids

α,ω-C16 1 100.0 ± 18.7 121.3 ± 17.1 125.1 ± 3.0 138.3 ± 15.1 402.5 ± 9.9 46.0 ± 5.1 110.1 ± 2.3 96.4 ± 18.7

2 58.2 ± 3.2 118.7 ± 2.9 70.9 ± 5.5 61.3 ± 1.0 76.1 ± 9.0 32.3 ± 0.3 72.8 ± 3.2 38.2 ± 3.1

3 61.9 ± 2.0 112.6 ± 9.2 101.7 ± 4.6 30.2 ± 0.9 107.3 ± 1.8 27.8 ± 0.1 126.4 ± 1639 NA

4 76.7 ± 3.5 51.2 ± 3.1 45.6 ± 4.4 118.1 ± 1.0 73.5 ± 4.1 20.5 ± 0.5 48.0 ± 3.5 23.9 ± 2.1

α,ω-C18 1 140.2 ± 24.7 210.1 ± 40.2 134.0 ± 1.0 161.8 ± 8.7 431.1 ± 103.9 61.3 ± 3.8 111.2 ± 0.5 91.0 ± 20.2

2 62.7 ± 11.8 171.7 ± 6.2 68.6 ± 7.4 91.9 ± 3.3 52.5 ± 4.0 49.3 ± 1.9 67.7 ± 5.2 65.2 ± 3.4

3 62.4 ± 1.3 166.6 ± 4.9 249.0 ± 15.0 47.3 ± 0.1 117.0 ± 12.7 43.4 ± 2.5 277.6 ± 17.8 NA

4 480.7 ± 49.8 76.7 ± 6.9 111.8 ± 55.6 250.2 ± 14.6 79.1 ± 7.4 31.1 ± 0.7 56.4 ± 4.9 34.3 ± 3.1

α,ω-C20 1 199.5 ± 22.3 301.0 ± 10.5 239.3 ± 8.1 248.7 ± 32.1 827.0 ± 7.3 108.4 ± 13.2 194.3 ± 6.3 187.0 ± 37.9

2 143.2 ± 19.9 332.2 ± 12.8 159.1 ± 18.9 202.1 ± 11.0 56.6 ± 1.1 101.8 ± 9.2 125.2 ± 18.3 125.8 ± 9.9

3 149.1 ± 19.0 308.7 ± 28.8 131.3 ± 4.2 97.4 ± 5.1 255.0 ± 30.6 83.3 ± 5.8 157.5 ± 2.9 NA

4 194.0 ± 15.9 114.6 ± 21.1 112.8 ± 22.6 306.4 ± 17.9 132.5 ± 10.3 70.2 ± 3.1 113.7 ± 9.6 70.8 ± 1.0

α,ω-C22 1 255.4 ± 27.3 318.3 ± 21.7 315.5 ± 116.0 326.0 ± 141.3 1487.0 ± 77.2 72.1 ± 4.2 163.1 ± 0.2 309.8 ± 40.5

2 129.6 ± 16.1 425.0 ± 0.1 163.4 ± 22.9 228.2 ± 11.2 47.1 ± 7.4 108.9 ± 3.9 97.0 ± 11.2 71.2 ± 56.0

3 152.4 ± 10.5 341.8 ± 68.5 103.0 ± 4.4 97.6 ± 5.6 206.5 ± 4.5 90.5 ± 6.2 108.3 ± 14.5 NA

4 142.7 ± 0.6 135.0 ± 16.7 82.6 ± 11.5 261.7 ± 24.5 99.1 ± 7.9 77.7 ± 1.7 87.7 ± 6.4 72.0 ± 11.7

Total n-alkanedioic

acids

1 695.2 ± 46.9 950.7 ± 49.9 813.9 ± 116.4 874.7 ± 145.9 3147.6 ± 130.0 287.8 ± 15.2 578.8 ± 6.8 684.1 ± 61.9

2 393.7 ± 28.3 1047.7 ± 14.5 462.1 ± 31.1 583.5 ± 16.1 232.3 ± 12.4 292.3 ± 10.1 362.7 ± 22.3 300.4 ± 57.0

3 425.8 ± 21.9 929.6 ± 75.1 584.9 ± 16.8 272.4 ± 7.6 685.7 ± 33.5 245.1 ± 8.8 669.8 ± 28.7 NA

4 894.2 ± 52.4 377.5 ± 27.9 352.8 ± 61.3 936.4 ± 33.7 384.1 ± 15.5 199.6 ± 3.7 305.8 ± 13.0 201.0 ± 12.4

Mid-chain hydroxy and epoxy acids

x,15-Dihydroxy C15

acids

1 21.5 ± 7.1 44.6 ± 1.7 8.8 ± 1.5 ND 49.7 ± 0.4 ND 11.4 ± 1.1 13.3 ± 7.1

2 16.9 ± 1.8 27.3 ± 5.9 17.4 ± 2.8 ND 21.5 ± 3.9 ND 26.2 ± 3.7 8.2 ± 1.0

3 18.3 ± 3.1 24.1 ± 0.3 30.8 ± 1.6 7.6 ± 1.1 51.8 ± 17.4 15.1 ± 7.1 54.0 ± 14.7 NA

4 15.2 ± 1.1 11.6 ± 0.7 22.1 ± 5.1 27.3 ± 2.3 17.3 ± 0.9 4.3 ± 0.1 7.8 ± 0.1 5.2 ± 0.6

7- or 8-Hydroxy C16

α,ω-dioic acid

1 91.0 ± 14.6 289.6 ± 18.2 141.1 ± 9.2 235.7 ± 108.2 894.8 ± 160.9 60.5± 24.8 155.3 ± 2.8 277.5 ± 96.3

2 281.9 ± 11.1 349.8 ± 17.5 333.5 ± 67.9 254.7 ± 26.4 228.5 ± 45.0 84.7 ± 16.3 270.0 ± 24.5 99.3 ± 6.8

3 224.6 ± 58.0 246.9 ± 6.9 406.0 ± 41.3 90.5 ± 13.5 471.1 ± 104.8 63.8 ± 16.1 320.1 ± 29.2 NA

4 469.1 ± 15.7 56.8 ± 11.1 444.6 ± 20.0 555.8± 50.6 271.2 ± 34.7 49.8 ± 6.0 160.0 ± 10.4 64.0 ± 13.8

x,16-Dihydroxy C16

acids

1 147.5 ± 27.2 247.2 ± 17.9 232.2 ± 1.9 416.2 ± 158.0 849.4 ± 14.1 87.1 ± 15.1 120.9 ± 2.3 439.2 ± 136.0

2 320.8 ± 86.0 247.8 ± 59.1 234.1 ± 52.7 238.8 ± 40.0 451.3 ± 60.5 86.4 ± 30.5 701.8 ± 142.1 162.5 ± 7.6

3 288.7 ± 74.7 265.6 ± 33.9 1053.2 ± 27.1 131.5 ± 28.1 322.5 ± 44.1 90.8 ± 24.5 753.0 ± 106.4 NA

4 ND 90.9 ± 14.7 450.4 ± 112.4 739.9 ± 129.0 301.6 ± 41.2 68.2 ± 10.8 160.1 ± 21.7 61.0 ± 3.9

9, 10-Epoxy-18-

hydroxy C18 acid

1 197.8 ± 58.6 668.3 ± 349.9 197.8 ± 3.4 231.8 ± 4.7 688.2 ± 35.8 122.1 ± 45.2 134.3 ± 17.5 169.8 ± 50.7

2 125.6 ± 15.2 284.3 ± 34.4 216.9 ± 1.7 132.5 ± 0.1 205.2 ± 34.8 66.0 ± 0.6 174.1 ± 21.2 80.2 ± 7.5

3 225.1 ± 17.2 397.1 ± 54.4 331.6 ± 19.6 71.1 ± 11.8 329.3 ± 1.0 64.5 ± 4.8 340.5 ± 24.5 NA

4 234.1 ± 7.9 111.3 ± 20.1 218.6 ± 24.6 469.2 ± 122.2 182.1 ± 26.1 53.9 ± 4.5 113.6 ± 15.6 75.1 ± 24.0

135

x,18- Dihydroxy C18

acids

1 52.0 ± 14.5 54.1 ± 19.9 14.8 ± 2.8 27.8 ± 3.0 ND 9.9 ± 2.7 27.9 ± 5.4 16.9 ± 0.2

2 14.2 ± 2.1 34.5 ± 2.4 18.6 ± 0.2 9.0 ± 0.8 29.4 ± 5.2 4.4 ± 0.3 20.6 ± 2.8 10.6 ± 2.6

3 27.9 ± 5.9 55.6 ± 1.7 25.8 ± 0.8 10.3 ± 0.1 33.8 ± 8.6 12.4 ± 2.1 30.9 ± 2.1 NA

4 32.2 ± 2.9 30.5 ± 4.8 22.9 ± 3.3 38.5 ± 2.1 25.9 ± 3.0 8.4 ± 1.7 15.6 ± 2.4 10.6 ± 1.2

9,10,18-Trihydroxy C18

acid

1 67.6 ± 27.1 620.8 ± 411.5 37.1 ± 4.7 145.7 ± 32.2 405.4 ± 71.8 17.7 ± 3.6 45.5 ± 3.5 104.8 ± 27.4

2 30.8 ± 2.0 72.4 ± 9.9 30.3 ± 1.1 50.8 ± 2.5 32.5 ± 1.5 21.8 ± 1.7 35.1 ± 2.9 28.4 ± 1.6

3 44.7 ± 5.1 170.5 ± 28.4 42.9 ± 5.0 39.5 ± 0.9 72.3 ± 10.3 37.1 ± 0.8 37.1 ± 2.9 NA

4 74.4 ± 2.2 74.1 ± 5.9 66.0 ± 5.2 199.6 ± 111.3 53.6 ± 7.7 24.3 ± 0.5 34.5 ± 10.4 28.4 ± 7.5

Total mid-chain-

substituted acids

1 577.3 ± 73.4 1924.6 ±

541.1

631.8 ± 11.5 1057.3 ±

194.3

2921.4 ± 183.5 297.3 ± 53.9 495.2 ± 19.0 1021.7 ± 176.5

2 790.3 ± 88.1 1016.2 ± 71.6 850.8 ± 86.0 685.7 ± 48.0 968.4 ± 83.3 263.4 ± 34.6 1227.7 ± 145.9 389.3 ± 13.1

3 829.3 ± 96.5 1159.9 ± 70.5 1890.3 ± 53.4 350.6 ± 33.4 1280.8 ± 115.8 283.7 ± 30.6 1535.6 ± 114.0 NA

4 1006.5 ± 104.4 375.1 ± 28.4 1224.5 ± 117.0 2030.4 ±

215.7

851.7 ± 60.5 208.9 ± 13.3 491.5 ± 30.7 244.3 ± 29.0

ω-Hydroxyalkanoic acids

ω-C16 1 314.9 ± 57.0 428.4 ± 51.8 600.0 ± 23.8 445.5 ± 52.8 1245.9 ± 26.3 146.4 ± 22.0 437.8 ± 12.0 295.9 ± 103.7

2 246.0 ± 10.9 366.1 ± 2.7 284.0 ± 15.1 217.6 ± 2.1 226.2 ± 28.6 112.2 ± 0.3 299.2 ±10.2 129.0 ± 4.8

3 232.8 ± 23.6 397.4 ± 39.9 469.5 ± 17.4 124.9 ± 12.4 365.7 ± 16.1 105.0 ± 9.5 540.0 ± 16.2 NA

4 299.2 ± 10.9 175.9 ± 18.6 296.1 ± 15.8 554.9 ± 54.2 272.3 ± 21.1 78.7 ± 3.4 159.4 ± 12.3 93.6 ± 1.7

ω-C18:1 1 34.8 ± 6.0 66.1 ± 0.3 12.2 ± 3.9 29.1 ± 11.0 117.7 ± 20.0 6.6 ± 2.1 17.2 ± 4.1 17.6 ± 0.2

2 16.7 ± 2.4 50.0 ± 10.1 18.0 ± 1.8 25.1 ± 1.6 25.0 ± 2.0 6.9 ± 1.1 41.4 ± 2.2 13.8 ± 0.9

3 25.6 ± 6.1 53.4 ± 5.9 43.5 ± 6.3 11.0 ± 2.2 35.1 ± 5.8 7.4 ± 1.5 67.1 ± 11.5 NA

4 30.3 ± 3.6 19.2 ± 0.1 37.2 ± 15.0 33.3 ± 2.8 22.0 ± 4.7 6.8 ± 0.7 12.4 ± 1.2 7.9 ± 1.6

ω-C22 1 109.9 ± 11.2 732.0 ± 191.2 592.9 ± 38.0 635.8 ± 175.1 2376.1 ± 202.2 249.8 ± 19.3 100.1 ± 10.6 397.9 ± 122.3

2 173.8 ± 49.8 481.5 ± 181.6 151.5 ± 39.2 277.2 ± 26.0 76.0 ± 12.9 80.4 ± 23.8 128.5 ± 21.4 132.0 ± 5.9

3 186.0 ± 39.5 667.9 ± 101.6 131.5 ± 0.6 107.1 ± 22.4 1191.1 ± 14.5 53.5 ± 8.5 224.9 ± 39.5 NA

4 378.1 ± 23.7 64.2 ± 0.5 389.5 ± 56.2 736.4 ± 32.1 99.4 ± 15.5 42.3 ± 6.9 105.7 ± 6.4 54.8 ± 22.0

Total ω-

hydroxyalkanoic acids

1 459.6 ± 58.4 1226.6 ±

198.1

1205.1 ± 45.0 1110.4 ±

183.3

3739.7 ± 204.9 402.7 ± 29.3 555.1 ± 16.5 711.5 ± 160.4

2 436.5 ± 51.0 897.6 ± 181.9 453.5 ± 42.0 519.9 ± 26.1 327.1 ± 31.4 199.5 ± 23.8 469.1 ± 23.8 274.8 ± 7.6

3 444.4 ± 46.4 1118.7 ±

109.3

644.5 ± 18.5 243.0 ± 25.7 1591.9 ± 22.4 165.9 ± 12.9 832.0 ± 44.2 NA

4 707.5 ± 26.3 259.2 ± 18.6 722.8 ± 60.3 1324.7 ± 63.1 393.6 ± 26.6 127.8 ± 7.8 277.6 ± 13.9 156.3 ± 22.2

Suberin and cutin monomersa

Suberin ∑S 1 568.8 ± 37.0 1351.2 ± 192.7 1147.7 ± 122.4 1210.4 ± 227.3 4690.1 ± 216.6 430.3 ± 23.7 457.6 ± 12.3 894.7 ± 134.3

2 446.6 ± 56.0 1238.8 ± 182.0 474.1 ± 49.2 707.5 ± 30.3 179.7 ± 14.9 291.0 ± 25.8 350.8 ± 30.3 329.0 ± 57.1

3 487.6 ± 45.1 1318.3 ± 125.9 365.7 ± 6.1 302.1 ± 23.6 1652.5 ± 34.2 227.4 ± 12.0 490.7 ± 42.2 NA

4 714.8 ± 28.5 313.8 ± 26.9 584.9 ± 61.7 1304.6 ± 44.2 330.9 ± 20.2 190.3 ± 7.8 307.1 ± 13.2 197.6 ± 25.0

136

NA= Not analyzed (the lower horizon soil sample for the warming +N+P fertilization treatment of block 3 was unavailable); ND = not detected aCalculated according to Otto and Simpson, 2006a; Suberin ΣS = ω-hydroxyalkanoic acids C20-C32 + ,-dioic acids C20-C32 + 9,10-ep C18 dioic acid; Cutin ΣC = mid-chain hydroxy C14,

C15, C17 acids + C16 mono- and dihydroxy acids and ,-dioic acids; Suberin or cutin ∑SvC = ω-hydroxyalkanoic acids C16, C18+ C18 di- and trihydroxy acids + 9,10-ep- ω-OH C18 acid +

α,ω-diacids C16, C18; Total suberin and cutin = ∑SC = ∑S + ∑C + ∑SvC; ∑C16 = ω-hydroxy C16 acid + α,ω-dioic C16 acid + ∑C16 mid-chain-substituted epoxy and hydroxy acids; ∑C18 = ω-

hydroxy C18 acid + α,ω-dioic C18 acid + ∑C18 mid-chain-substituted epoxy and hydroxy acids; Mid-chain-substituted acids ∑Mid = ∑C14-C18 mid-chain-substituted epoxy and hydroxy

acids; Total suberin and cutin ∑SC = ∑S + ∑C+ ∑SvC

Cutin ∑C 1 260.0 ± 26.3 581.4 ± 29.1 382.1 ± 9.5 651.9 ± 180.8 1793.9 ± 161.6 147.6 ± 27.1 287.6 ± 3.4 730.1 ± 153.6

2 619.7 ± 70.9 624.9 ± 65.7 585.0 ± 83.3 493.4 ± 47.0 701.3 ± 97.7 171.1 ± 33.0 997.9 ± 130.6 270.1 ± 9.0

3 531.6 ± 88.1 536.7 ± 26.5 1490.0 ± 50.9 229.7 ± 27.9 845.5 ± 134.6 169.7 ± 26.2 1127.1 ± 129.4 NA

4 665.7 ± 103.4 159.0 ± 15.2 917.1 ± 117.0 1323.0 ± 109.5 590.1 ± 47.3 122.3 ± 10.4 327.9 ± 24.2 130.1 ± 26.8

Suberin or cutin ∑SvC 1 907.3 ± 83.4 2169.2 ± 477.2 1121.0 ± 25.9 1180.1 ± 71.4 3324.8 ± 144.9 409.9 ± 60.4 883.9 ± 22.6 792.5 ± 115.3

2 554.3 ± 24.3 1097.9 ± 75.6 707.3 ± 19.7 588.2 ± 5.4 646.8 ± 37.3 293.0 ± 30.9 710.9 ± 20.8 365.4 ± 9.6

3 680.4 ± 28.4 1353.2 ± 93.0 1264.0 ± 30.0 334.3 ± 16.4 1060.5 ± 32.4 297.6 ± 27.2 1419.6 ± 49.4 NA

4 1227.7 ± 52.9 539.0 ± 36.2 798.2 ± 65.0 1663.9 ± 151.5 708.4 ± 29.4 223.7 ±12.6 439.9 ± 20.0 273.9 ± 19.3

Suberin ∑S/Cutin ∑C 1 1.27 ± 0.12 1.32 ± 0.30 1.51 ± 0.09 1.31 ± 0.19 1.57 ± 0.08 1.53 ± 0.22 1.15 ± 0.03 1.13 ± 0.18

2 0.85 ± 0.08 1.36 ± 0.14 0.92 ± 0.07 1.20 ± 0.06 0.61 ± 0.06 1.26 ± 0.15 0.62 ± 0.05 1.10 ± 0.09

3 0.97 ± 0.09 1.41 ± 0.11 0.59 ± 0.02 1.13 ± 0.08 1.43 ± 0.11 1.15 ± 0.11 0.75 ± 0.05 NA

4 1.03 ± 0.07 1.22 ± 0.09 0.81 ± 0.08 1.01 ± 0.08 0.80 ± 0.04 1.21 ± 0.07 0.97 ± 0.05 1.17 ± 0.12

Total suberin and cutin

∑ SC

1 1732.1 ± 94.9 4101.8 ± 515.5 2650.8 ± 125.4 3024.4 ± 299.1 9808.7 ± 306.6 987.8 ± 70.3 1629.1 ± 26.0 2417.3 ± 234.4

2 1620.6 ± 93.5 2961.5 ± 207.8 1766.3 ± 98.7 1789.2 ± 56.2 1527.8 ± 105.6 755.1 ± 52.0 2059.5 ± 135.7 964.5 ± 58.6

3 1700.0 ± 93.5 3208.2 ± 158.7 3119.7 ± 59.4 866.0 ± 40.1 3558.5 ± 142.6 694.7 ± 39.6 3037.4 ± 144.8 NA

4 2608.2 ± 116.9 1011.8 ± 47.6 2300.2 ± 147.4 4291.5 ± 192.1 1629.5 ± 59.3 536.3 ± 18.1 1074.9 ± 34.0 601.6 ± 41.4

Cutin degradation ratiosa

ω-C16/∑C16 1 0.48 ± 0.1 0.39 ± 0.08 0.55 ± 0.02 0.38 ± 0.07 0.37 ± 0.02 0.43 ± 0.07 0.53 ± 0.01 0.26 ± 0.08

2 0.27 ± 0.03 0.34 ± 0.04 0.31 ± 0.03 0.28 ± 0.02 0.23 ± 0.01 0.36 ± 0.05 0.23 ± 0.03 0.30 ± 0.01

3 0.29 ± 0.04 0.39 ± 0.03 0.23 ± 0.01 0.33 ± 0.04 0.29 ± 0.08 0.38 ± 0.04 0.31 ± 0.05 NA

4 0.30 ± 0.03 0.47 ± 0.05 0.24 ± 0.04 0.28 ± 0.03 0.30 ± 0.02 0.37 ± 0.02 0.30 ± 0.02 0.39 ± 0.07

ω-C18/∑C18 1 0.07 ± 0.01 0.05 ± 0.02 0.03 ± 0.01 0.05 ± 0.02 0.07 ± 0.01 0.03 ± 0.01 0.05 ± 0.01 0.05 ± 0.01

2 0.07 ± 0.01 0.08 ± 0.02 0.05 ± 0.01 0.08 ± 0.01 0.07 ± 0.01 0.05 ± 0.01 0.12 ± 0.01 0.07 ± 0.01

3 0.07 ± 0.02 0.06 ± 0.01 0.06 ± 0.01 0.06 ± 0.01 0.06 ± 0.01 0.04 ± 0.01 0.09 ± 0.02 NA

4 0.04 ± 0.01 0.06 ± 0.01 0.08 ± 0.03 0.04 ± 0.01 0.06 ± 0.01 0.05 ± 0.01 0.05 ± 0.01 0.05 ± 0.01

∑Mid/∑SC 1 0.33 ± 0.05 0.45 ± 0.1 0.24 ± 0.01 0.34 ± 0.07 0.30 ± 0.02 0.29 ± 0.06 0.30 ± 0.01 0.41 ± 0.08

2 0.49 ± 0.06 0.34 ± 0.03 0.48 ± 0.01 0.38 ± 0.03 0.63 ± 0.07 0.35 ± 0.05 0.58 ± 0.08 0.40 ± 0.03

3 0.48 ± 0.06 0.36 ± 0.03 0.61 ± 0.01 0.40 ± 0.04 0.36 ± 0.04 0.40 ± 0.05 0.50 ± 0.04 NA

4 0.39 ± 0.04 0.37 ± 0.03 0.53 ± 0.06 0.47 ± 0.05 0.52 ± 0.04 0.38 ± 0.03 0.45 ± 0.03 0.39 ± 0.05

137

Table A6: Solid-state 13

C CPMAS-NMR integration results with relative contribution (%) of the four main carbon structures and calculated

alkyl/O-alkyl ratios for the Control treatments of the upper horizon soils of each block. These soils were not treated with hydrofluoric acid.

Block Alkyl C

(0-50 ppm)

O-Alkyl C

(50-110 ppm)

Aromatic and Phenolic C

(110-165 ppm)

Carboxylic and Carbonyl C

(165-215 ppm)

Alkyl/O-Alkyl

1 45 39 6 10 1.15

2 43 42 7 8 1.02

3 35 45 12 8 0.78

4 21 54 21 4 0.39

138

Figure A1: Solid-state 13

C CPMAS-NMR spectra of the upper horizon soils of the Control

treatments of each block with the four major spectral regions: alkyl (0-50 ppm), O-alkyl (50-

110 ppm), aromatic and phenolic (110-165 ppm) and carboxylic and carbonyl carbon (165-

215 ppm). These soils were not treated by hydrofluoric acid.

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