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This is to certify that the
dissertation entitled
THE DYNAMICS OF CARBON, NITROGEN AND SOIL
ORGANIC"MATTER '.IN POPULUS PLANTATIONS

presented by

WILLIAM RICHARD HORWATH

has been accepted towards fulﬁllment
of the requirements for

DOCTOR OF PHILOSOPHY degree in CROP AND SOIL SCIENCES
AND FORESTRY

 

 

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Date3/31/1993

 

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MSU Is An Affirmative Action/Equal Opportunity Institution
GWMHJ

 

 

 

THE DYNAMICS OF CARBON, NITROGEN AND SOIL ORGANIC
MATTER IN POPULUS PLANTATIONS

By

William Richard Horwath

A DISSERTATION
Submitted to
Michigan State University
in partial fulﬁllment of the requirements
for the degree of
DOCTOR OF PHILOSOPHY
Departments of Crop and Soil Science
and

Forestry
1993

ABSTRACT

THE DYNAMICS OF CARBON, NITROGEN AND SOIL ORGANIC
MATTER IN POPUL US PLANTATIONS

By

William Richard Horwath

Short-rotation intensive-culture forestry is similar to agricultural
systems requiring increased nutrient input and management. The
expense and environmental concerns associated with fertilizers have
raised questions about the sustainability of these ecosystems. Sustainability
of production oriented ecosystems can be aided by understanding the
mineralization-immobilization potential of the soil microbial biomass. The
soil microbial biomass is central to a complex system of soil organic
fractions that control soil fertility, production and environmental
contamination.

The lack of root turnover studies has led to a inadequate
understanding of the role of root turnover as substrate soil microbial
biomass and organic matter formation. The current study was designed to
determine: (i) the role of below-ground production and turnover in nutrient
cycling processes; (ii) the contribution of leaf and root litter as substrate for
the maintenance of soil organic matter; and (iii) relate soil microbial
biomass and organic matter dynamics to plant carbon and nitrogen
allocation patterns.

Two-year-old Populus euramericana cv. Eugenei trees were labeled
with 140 and 15N in the ﬁeld. The 15N was injected into the stem to label
leaves and roots without labeling the soil. The 14C labeling was done in a

Plexiglas chamber under ambient conditions. The tree-soil system was

sampled for one year. Labeled leaf litter was placed onto unlabeled tree
plots to differentiate the contribution of leaf and root derived carbon and
nitrogen to soil over a two year period.

The 14:0 required two weeks to stabilize in the root system and
averaged 20% of soil respiration. One year latter, 32% of applied 14C and
33% of the injected 15N was recovered. Reserves in the root system were
sufﬁcient to replace ﬁne roots one and one half times. This represented
signiﬁcantly less C than leaf litter, yet similar amounts of 14C were found
in soil from both litters. Leaf litter appeared to dominate N cycling since
15N was not detected in root labeled soil. Kinetic analysis of incubated soil
showed a greater contribution of C and N to soil organic fractions from leaf
litter. The turnover of labile soil 140 in both leaf and root litter labeled soil
was 14-64 days. The 14C in pools of intermediate resistance had a turnover

time of 2-16 years and increased with soil depth.

Copyright by
WILLIAM RICHARD HORWATH
1993

Theresa Marie
and

Derek Ryan

ACKNOWLEDGMENTS

I sincerely thank Dr. Eldor Paul for his guidance and endless
enthusiasm. Dr. Paul provided me with endless hours of fruitful discussion
and consultation concerning the area of expertise that motivated both of us. I
am grateful to Dr. Kurt Pregitzer for his motivation, input and review of my
writings. Dr. Pregitzer's interest in forest ecology was common to both of us.
I appreciate the assistance and encouragement that Dr. Don Dickmann
always seemed to have concerning my potential and research. I thank Dr. G.
Philip Robertson for his help, advice and use of laboratory space. I thank the
Kellogg Biological Station and the Tree Research Center for space and
personnel to assist in my research. In addition, I would like to thank the
National Science Foundation (NSF), Long-term Ecological Research (LTER)
program, NSF and the Department of Energy for the funding and support
which made this research possible. The National Science Foundation also
provided invaluable technical assistance through the Research for
Undergraduate Experience program.

I thank Matt Zwiernik for his technical assistance and console in
interpreting the results of this research. I thank Nancy Richte, Scott Muske,
Lesie Jagger and Melisa Chapla for their technical assistance.

Special appreciation and love go to Theresa my wife and Derek my son,
for their love, support and encouragement.

Finally, I thank my parents John and Regina who have always
supported and encouraged me to do my best. My work is an expression of the
ethics and value which they instilled upon me.

TABLE OF CONTENTS

LIST OF TABLES x

LIST OF FIGURES xix

Chapter

II.

III.

INTRODUCTION

Rational and Rational

Fine Root Turnover

Microbial Biomass and Soil Organic Matter Dynamics
Objective and Goals

Overview of Chapters

Bibliography 1

Hooozcnton-a

THE INJECTION OF NITROGEN-15 INTO TREES TO STUDY

NITROGEN CYCLING IN SOIL

Abstract 16

Introduction 17

Material and Methods 18
Injection Procedure 18
Field Protocol and Sampling 19
Tissue Analysis 21

Results and Discussion 22

Bibliography 28

14C AND 15N PARTITIONING IN TREE-SOIL-SOIL SYSTEMS

Abstract 30
Introduction 31
Material and Methods 32
Expression of Results 37
Results and Discussion 37
Tree and Environmental Variables 37
Root-Soil Respiration 4O

Carbon Partitioning 46
Nitrogen Distribution 48
Bibliography 54

THE FATE OF 14C AND 15N IN HYBRID POPLAR
PLANTATIONS

Abstract 58
Introduction 59
Material and Methods 62
15N Injection 63
14C Labeling 63
Sampling Procedure 63
Plant Tissue and Soil Analysis 64
Microbial Biomass Determinations 65
Litter Exchange Experiment 67
Expression of Results 68
Results 68
Tree Characteristics 68
Distribution of C and N in the Soil Tree-System 70
Distribution of 14C and 15N in the Tree 74
Speciﬁc Activity of 14C 76
Distribution of 15N Activity 80
The Fate of Labeled C and N 84
Dynamics of 14C and 15N in Leaf Litter 89
Discussion 92
Carbon and Nitrogen Allocation 92
The Fate of 14C 93
The Fate of 15N 95
The Turnover of 14C and 15N From Litter 98
Conclusion 102
Bibliography 103

THE DYNAMICS OF ASSIMILATED 14002 IN THE CHEMICAL

FRACTIONS OF THE SHOOTS AND ROOTS OF POPUL US
PLANTATIONS

Abstract 110
Introduction 111
Material and Methods 113
Plant material 113
Sampling Procedure 1 14
Analysis of Chemical Fractions 114
Analysis of the Chloroform Phase 116
Analysis of Sugars 116

Analysis of Proteins 118

viii

VII.

Analysis of Starch
Analysis of the Structural Residues

Results
Distribution of Chemical Fractions
Distribution of 14C Within Chemical Fractions
Redistribution of l‘er Over Time
Distribution of 140 Within Fractions
Fate of Reserves

Discussion

Conclusion

Bibliography

MICROBIAL BIOMASS AND DYNAMICS OF C AND N IN
POP UL US PLANTATIONS

Abstract
Introduction
Material and Methods
Plant 140 and 15N labeling
Microbial C and N Determinations
Long-Term Soil Incubations
Calculations
Results
Microbial Biomass
Long-Term Incubations
Soil C
N Dynamics
Microbial Biomass C and N
Potentially Mineralizable C
Discussion
Microbial Biomass Determinations
Long-term Soil Incubations
Potentially Mineralizable Pools of C
Conclusions
Bibliography

Conclusion

APPENDDI A

APPENDIX B

ix

119
120
120
120

125
130

133
151
156
163
165

170
172
175
175
177
178
179
180
180
191
192
196
199
210
222
222
223
225
227
229

234

239

244

LIST OF TABLES

Table 2.1. Mean leaf N concentrations and the distribution of

15N in the most recently mature leaf of each branch
according to crown aspect 3 days following injection of tree
set 1. Standard error given in parentheses.

Table 2.2. Summary of N and carbon concentrations and the

recovery of 15N in senescent leaves. The 15N recovered is
based on the amount injected. Standard error given in
parentheses.

Table 2.3. Mean labeled N and N concentrations of different
classes of roots collected over time from tree sets 1 and 2.
Standard error given in parentheses.

Table 2.4. Percentage of N in various root classes derived from
15N two and eight weeks after injection.

Table 3.1. The dry weight of tree components in July and
September. Standard error are shown in parentheses.

Table 3.2. Gas exchange and environmental characteristics
during labeling periods in July and September.

Table 3.3. Carbon concentration (%), speciﬁc activity (Bq mg-l
C), % of 140 recovered of total applied (Bq/Bq added), and %

14C of the total amount of 14C recovered for the different
tree components. Standard error given in parentheses.

Table 3.4. Nitrogen concentration (%), atom % 15N excess,
nitrogen derived from applied l5N (%Ndfl5N) and % 15N
recovered of the total amount applied for the various tree
components. Standard error are shown in parentheses.

47

Table 3.5. The interrelationships between the distribution of

15N and 14C of the various tree components from the July
and September labelings. The values are expressed as the

ratio of the distribution of 15N to 140 recovered in the tree
components.

Table 4.1. Height and leaf area measurements taken from
samplings done during the growing season from trees

labeled July 19, 1990 and September 5, 1990. Standard error
of the mean shown in parentheses.

Table 4.2. The dry weight (g) of tree components from July 19,
1990 dual labeled trees sampled over the following year.
Standard error of the mean shown in parentheses.

Table 4.3. The dry weight (g) of tree components from
September 5, 1990 dual labeled trees sampled over the
following year. Standard error of the mean shown in
parentheses.

Table 4.4. The speciﬁc activity (Bq mg’1 C) for the different tree

components sampled during the year following 140 labeling
on July 19, 1990. Standard error of the mean shown in
parentheses.

Table 4.5. The speciﬁc activity (Bq mg'1 C) for the different tree

components sampled during the year following 14C labeling
on September 5, 1990. Standard error of the mean shown in
parentheses.

Table 4.6. Atom percent 15N excess (atom % - natural
abundance) is shown for the different tree components

sampled the year following 15N injection on July 14, 1990.
Standard error of the mean shown in parentheses.

Table 4.7. Atom percent 15N excess (atom % - natural
abundance) for the different tree components sampled

during the year following 15N injection on August 31, 1990.
Standard error of the mean shown in parentheses.

xi

71

81

Table 4.8. Recovery of labeled C in mg* and percent of the
amount applied from tree-soil components sampled the
year following labeling July 19, 1990. Standard error of the
mean shown in parentheses.

Table 4.9. Recovery of labeled C in mg”: and percent of the total
amount applied from tree-soil components sampled the
year following labeling September 5, 1990. Standard error
of the mean shown in parentheses.

Table 4.10. Recovery of labeled N in mg* and percent of the
total amount applied the year following injection July 14,
1990. Standard error of the mean shown in parentheses.

Table 4.11. Recovery of labeled N in mg* and percent of the
total amount applied the year following injection August 31,
1990. Standard error of the mean shown in parentheses.

Table 4.12. The fate of labeled C in leaf litter from both the July
and September labelings. Labeled C in litter, soil and
microbial biomass was sampled over 613 days. Total values
represent soil microbial biomass. Standard error of the
mean shown in parentheses were applicable.

Table 4.13. Fate of labeled N in leaf litter from both the July
and September stem injections. Labeled N in litter, soil and
microbial biomass sampled over 613 days. Total values
represent soil -microbial biomass. Standard error of the
mean shown in parentheses were applicable.

Table 5.1. Concentration (mg g’1) of chemical fractions found
in shoot and root components during growth and
dormancy. July 19, 1990 labeling with subsequent sampling
August 2, 1990 and November 2, 1990. Standard error of the
mean in parentheses.

Table 2. Concentration (mg g'l) of chemical fractions found in
shoot and root components during growth and dormancy.
September 5, 1990 labeling with subsequent sampling
September 20, 1990 and November 4, 1990. Standard error of
the mean in parentheses.

xii

91

121

Table 5.3. Amount of labeled C (%) found in the different
chemical fractions expressed as the percentage of C derived
from 140. July labeling with subsequent August 2, 1990
and November 2, 1990. Standard error of the mean in
parentheses.

Table 5.4. Amount of labeled C (%) found in the different
chemical fractions expressed as the percentage of C derived
from 14C. September 5, 1990 labeling with subsequent
sampling September 20, 1990 and November 4, 1990.
Standard error of the mean in parentheses.

Table 5.5. Concentration (mg g'1)of chemical fractions found

in ﬁne root components the year following 14C labeling.
Data is presented for subsequent sampling of trees on July
25, 1991 for the July 19, 1990 labeling and September 5, 1991
for the September 5, 1990 labeling. Standard error of the
mean in parentheses.

Table 5.6. Amount of labeled C (%) found in the different
chemical fractions expressed as the percentage of C derived
from 1‘er one year following 14C labeling. Data is presented
for subsequent sampling of trees on July 25, 1991 for the
July 19, 1990 labeling and September, 1991 for the
September 5, 1990 labeling. Standard error of the mean in
parentheses.

Table 6.1. Characteristics of the plant-soil system studied.

Table 6.2. Direct microscopic counts of bacterial numbers and
hyphal lengths and biomass C conversions for the ﬁrst two
samplings of labeled trees. Standard error of the mean
shown in parentheses.

Table 6.33. Microbial biomass C determined by the CFE
method for the ﬁrst and second samplings of trees labeled

with 14C and 15N July 19,1990 and September 5,1990.
Standard error of the mean shown in parentheses.

xiii

131

176

182

183

Table 6.3b. Microbial C determined by the CFE method for the
ﬁrst sampling, June 6, 1991, of the litter exchange
experiment. Standard error of the mean shown in
parentheses.

Table 6.4a. Microbial biomass C, 14‘0 and N determined by the
CFI method for all samplings of trees labeled on July
19,1990 and September 5,1990. Standard error of the mean
shown in parentheses.

Table 6.4b. Microbial biomass C, 14C and N determined by the
CFI method for all samplings of the litter exchange
experiment on June 6, 1991 and November 14, 1991.
Standard error of the mean shown in parentheses.

Table 6.5. Average results from all samplings and methods of
determining microbial biomass C. The data has been
adjusted to 100 ug C=CFI. Standard error of the mean
shown in parentheses.

Table 6.6. Microbial biomass C, speciﬁc activity and percent of

applied 14C from long term soil incubations of tree plots
labeled July 19, 1990. Standard error of the mean shown in
parentheses.

Table 6.7. Microbial biomass C, speciﬁc activity and percent of

applied 14C from long term soil incubations of tree plots
labeled September 5, 1990. Standard error of the mean
shown in parentheses.

Table 6.8. Microbial biomass C, speciﬁc activity and percent of

applied 140 from long term soil incubations of the litter
exchange plots. Standard error of the mean shown in
parentheses.

Table 6.9. Microbial biomass N from the long-term
incubations of soil from the tree plots labeled July 19, 1990.
Standard error of the mean shown in parentheses.

xiv

201

Table 6.10. Microbial biomass N from the long-term
incubations of soil from the tree plots labeled September 5,
1990. Standard error of the mean shown in parentheses.

Table 6.11a. Microbial biomass N, 15N and percent of applied

15N from soil incubations of September litter plots.
Standard error of the mean in parentheses.

Table 6.11b. Microbial biomass N, 15N and percent of applied

15N from soil incubations of July litter plots. Standard
error of the mean in parentheses.

Table 6.12. Parameters developed from the modiﬁed
exponential equation applied to the 002 mineralization data
from long-term incubations of soil from the sampling of
labeled tree plots. MRT is mean residence time.

Table 6.13. Amount of C found in the soil microbial biomass of
the trees labeled in July and September 1990 and C
mineralized at 150 (C150). Csoil is total soil C. Biomass C is
C in the soil biomass.

Table 6.14. Parameters developed from the modiﬁed
exponential equation applied to the 002 mineralization data
from long-term incubations of soil from the sampling of leaf
litter exchange plots. MRT is mean residence time. Csoil
is total soil C.

Table 6.14a Represents the amount of C found in the soil
microbial biomass of the leaf litter exchange experiment
and C mineralized at 150 (C150) days. Csoil is total soil C.
Biomass C is C in the soil biomass.

Table 6.15. Parameters developed from the modiﬁed
exponential equation applied to the 14002 mineralization

data from long-term incubations of soil from the sampling
of labeled tree plots. NIRT is mean residence time.

211

212

213

214

216

Table 6.16. Represents the amount of radiolabeled C found in
the soil microbial biomass and 14C mineralized at 150 days
(140150). 14Csoil is total soil 14C. Biomass 14C is 14C in the
soil biomass.

Table 6.17. Parameters developed from the modiﬁed

exponential equation applied to the 14C02 mineralization
data from long-term incubations of soil from the sampling
of the litter exchange plots. MRT is mean residence time.

Table 6.17a. Amount of radiolabeled C found in the soil
microbial biomass of the litter exchange experiment and

14C mineralized at 150 (140150). 14Csoi1 is total soil 14C.
Biomass 14C is 14C in the soil biomass.

Table 6.18. N mineralized was calculated by linear regression.
N150 is the N mineralized 150 days. N301] is total soil N.
Biomass N/N15o is a ratio.

Table A.1. The carbon concentration of tree components
sampled one year after the dual labeling procedure on July
19, 1990. Standard error of the mean shown in parentheses.

Table A2. Carbon concentration of tree components sampled
one year after the dual labeling procedure on September 5,
1990. Standard error of the mean shown in parentheses.

Table A.3. Nitrogen concentration of tree components sampled
one year after the dual labeling procedure on July 19, 1990.
Standard error of the mean shown in parentheses.

Table A.4. Nitrogen concentration of tree components sampled
one year after the dual labeling procedure on September 5,
1990. Standard error of the mean shown in parentheses.

Table B.1. Results of C and 14C mineralization from long-
term laboratory soil incubations done on the last two
samplings of trees labeled July 19, 1990. The soil depth is 0
to 25 cm. Standard error of the mean shown in
parentheses.

xvi

217

218

219

221

241

Table 3.2. Results ofC and 14C mineralization from long-
term laboratory soil incubations done on the last two
samplings of trees labeled July 19, 1990. The soil depth is
25-60 cm. Standard error of the mean shown in
parentheses.

Table B.3. Results of C and 140 mineralization from long-
term laboratory soil incubations done on the last two
samplings of trees labeled July 19, 1990. The soil depth is
60- 100 cm. Standard error of the mean shown in
parentheses.

Table B.4. Results of C and 1‘er mineralization from long-
term laboratory soil incubations done on the last two
samplings of trees labeled September 5, 1990. The soil depth
is 0-25 cm. Standard error of the mean shown in
parentheses.

Table B.5. Results of C and 140 mineralization from long-
term laboratory soil incubations done on the last two
samplings of trees labeled September 5, 1990. The soil depth
is 25-60 cm. Standard error of the mean shown in
parentheses.

Table B.6. Results of C and 14C mineralization from long-
term laboratory soil incubations done on the last two
samplings of trees labeled September 5, 1990. The soil depth
is 60-100 cm. Standard error of the mean shown in
parentheses.

Table B.7. Results from long-term laboratory soil incubations
done on the ﬁrst two samplings of the litter exchange
experiment started December 21, 1990. Litter is from the
September labeled trees. The soil depth is O to 10 cm.
Standard error of the mean shown in parentheses.

Table B.8. Results from long-term laboratory soil incubations
done on the ﬁrst two samplings of the litter exchange
experiment started December 21, 1990. Litter is from the
September labeled trees. The soil depth is 10 to 25 cm.
Standard error of the mean shown in parentheses.

xvii

251

252

Table B.9. Results from long-term laboratory soil incubations
done on the ﬁrst two samplings of the litter exchange
experiment started December 21, 1990. Litter is from the
July labeled trees. The soil depth is O to 10 cm. Standard
error of the mean shown in parentheses.

 

Table B.10. Results from long-term laboratory soil incubations
done on the ﬁrst two samplings of the litter exchange
experiment started December 21, 1990. Litter is from the
July labeled trees. The soil depth is 10 to 25 cm. Standard
error of the mean shown in parentheses.

 

Table 3.11. N mineralization results are shown for the last two
samplings of tree labeled July 19, 1990. All soil depths are
indicated. Standard error of the mean shown in
parentheses..

Table B.12. N mineralization results are shown for the last two
samplings of tree labeled September 5, 1990. All soil depths
are indicated. Standard error of the mean shown in
parentheses.

Table 3.13. Results from N and 15N mineralization during
long-term soil incubations. Soil collected from the 0-10 cm
depth under the September labeled litter. Standard error of
the mean shown in parentheses.

Table B.14. Results from N and 15N mineralization during
long-term soil incubations. Soil collected from the 0-10 cm
depth under the July labeled litter. Standard error of the
mean shown in parentheses.

Table B.15. N mineralization results are shown from both
samplings of September and July litter exchange
experiment. Soil was collected from the 10-25 cm depth.
Standard error of the mean shown in parentheses.

xviii

257

LIST OF FIGURES

Figure 2.1. Diagram of the injection technique.

Figure 3.1. Schematic diagram of the 14C exposure system
used to label eight, two-year-old whole poplar trees. The
trees were approadmately three meters tall when labeled.

Figure 3.2. Root-soil respiration averaged over eight trees for
the period following labeling in July and September.

Values are expressed as mg COz-C m'2 hr'l. Standard
error of the mean are depicted as line bars.

Figure 3.3. Average speciﬁc activity of below-ground
respiration. Each symbol represents the mean of eight root-
soil microcosms.

Figure 3.4. Carbon- 14 respired from the root-soil system.

Figure 4.1. The distribution of C and N (g) in the Populus-soil
system for July 19, 1990 labeled trees sampled August 2,
1990. The values in parentheses represent the amount of C
and N for the November 2, 1990 sampling.

Figure 4.2. Relative distribution of weight (g), 15N (mg) and
14C (mg) within trees labeled on July 19, 1990 (A) and
September 5, 1990 (B).

Figure 5.1. Diagram of the tissue fractionation protocol.

xix

75

115

Figure 5.2a. The distribution of total sugars and labeled 14C-
sugars among the shoot and root components. The July
labeling with subsequent sampling August 2, 1990 and
November 2, 1990 show the acropetal translocation tendency
of sugars synthesized in July during mid-growing season.
Standard error of the mean shown as line bars.

Figure 5.2b. The distribution of total sugars and labeled 14C-
sugars among the shoot and root components. The
September labeling with subsequent sampling September
20, 1990 and November 4, 1990 show the basipetal
translocation of C assimilated in September during autumn
for this hybrid poplar clone. Standard error of the mean
shown as line bars.

Figure 5.3a. The distribution of total starch and labeled 14C--
starch among the shoot and root components. The July
labeling and subsequent sampling August 2, 1990 and
November 2, 1990 show the tendency of this clone to
translocate C to the root system for storage in July during
mid-growing season. The distribution of labeled starch
increases in the root system as dormancy is induced.
Standard error of the mean shown as line bars.

Figure 5.3b. The distribution of total starch and labeled 14C-
starch among the shoot and root components for trees
labeled September 5,1990. The subsequent sampling done
on September 20, 1990 and November 4, 1990 show the
basipetal translocation and loading of the coarse root
system with starch and labeled C assimilated during
autumn. Standard error of the mean shown as line bars.

Figure 5.4a. The distribution of total residue and labeled 14C-
residue among the shoot and root components of trees
labeled July 19, 1990. The subsequent sampling done on
August 2, 1990 and November 2, 1990 show that the shoot
residue comprises a signiﬁcant proportion of the tree.
Standard error of the mean shown as line bars.

XX

141

144

Figure 5.4b. The distribution of total residue and labeled 14:0-
residue among the shoot and root components of trees
labeled September 5, 1990. The subsequent sampling done
on September 20, 1990 and November 4, 1990 show that the
root residue is both increasing in mass and receiving more
labeled C at this time during the season. Standard error of
the mean shown as line bars.

Figure 5.5a. The distribution of 14C within the chemical
fractions of the trees labeled July 19,1990. The subsequent
samplings done on August 2, 1990 and November 2, 1990
reveal that trees labeled during the growing season
synthesize a signiﬁcant amount of the residue fraction as a
result of shoot and root expansion. Standard error of the
mean shown as line bars.

Figure 5.5b. The distribution of 1‘er within the chemical
fractions of the trees labeled September 5,1990. The
subsequent samplings done on September 19, 1990 and
November 4, 1990 show that trees labeled after budset
synthesize less of the residue fraction and allocate more
photosynthates to labile fractions. Standard error of the
mean shown as line bars.

Figure 5.6a. The amount of ﬁne roots and the total
nonstructural carbohydrates (sugars and starch) in the root
system for the July labeling and subsequent harvests (upper
graph). The speciﬁc activities of sugar, starch and residue
fraction of ﬁne roots (< 0.5 mm) for the year following
labeling (lower graph). Standard error of the mean shown
as line bars.

Figure 5.6b. The amount of ﬁne roots and the total
nonstructural carbohydrates (sugars and starch) in the root
system for the September labeling and subsequent harvests
(upper graph). The speciﬁc activities of sugar, starch and
residue fraction of ﬁne roots (< 0.5 mm) for the year
following labeling (lower graph). Standard error of the
mean shown as line bars.

xxi

146

148

Figure 6.1. The comparison of the fumigation and direct
microscopy methods of microbial biomass determination
are shown in a-c. Figures d and e compare the biomass C
and speciﬁc activity of biomass C from the fumigation
methods.

Figure 6.2. The C (Fig. 6.2a) and 140 (Fig. 6.2b)
mineralization data for the ﬁrst (60 days) and second (366
days) long-term incubation of September labeled trees. The
graph lines are simulated values calculated using equation
2. Standard error of the mean shown as line bars.

Figure 6.3. The C (Fig. 6.3a)and 14 C (Fig. 6%)
mineralization data for the ﬁrst (168 days) and second (328
days) long-term incubation of September litter exchange.
The graph lines are simulated values calculated using
equation 2. Standard error of the mean shown as line bars.

Figure 6.4. The decline in speciﬁc activity of mineralized C
during the long-term incubation of soils labeled through
leaf and root derived C. Figure 6.4a and 6.4b are from leaf
litter plots and 6.4c and 6.4d are from plots from the whole
tree labeling. Standard error of the mean shown as line
bars.

Figure 6.5. Cumulative N and 15N mineralization curves
from long-term soil incubations of surface soil (0-10 cm)
from the litter exchange experiment. Figure 6.5a
represents the ﬁrst sampling on June 6, 1991 and ﬁgure
6.5b the sampling done on November 14, 1991 of both
September and July litter. Standard error of the mean
shown as line bars.

Figure 6.6. Nitrogen mineralization data for all the
subsurface (10-25 cm) samplings of the leaf litter exchange
(Fig 6.6a) and all soil depths of the ﬁrst long-term
incubation of July and September labeled tree plots (Fig.
6.6b). Soil depth and dates are indicated. Standard error of
the mean shown as line bars.

xxii

Figure 6.7. The amount of mineralized 15N and microbial

biomass 15N for the surface soil of the September labeled
litter. Figure 6.7a represents the ﬁrst sampling on June 6,
1991 and ﬁgure 6.7b the sampling done on November 14,
1991. Stande error of the mean shown as line bars.

Figure 6.8. Speciﬁc activity of mineralized C and microbial
biomass C for the surface soil of the September labeled
litter. Figure 6.8a represents the ﬁrst sampling on June 6,
1991 and ﬁgure 6.8b the November 14, 1991 sampling.
Standard error of the mean shown as line bars.

xxiii

Chapter 1

INTRODUCTION

Rationale and Justiﬁcation

High production-short rotation plantation forestry has been
suggested as a solution to limited fossil fuels and demand for pulp close to
processing facilities. These plantations often contain hybrids, such as
Populus, requiring nutrient supplies that are in excess of traditionally
managed forests and even higher than some intensively managed
agricultural systems (Ingestad and Agren, 1984). The sustainability of
such high production systems, especially successive rotations, will depend
on management techniques based on understanding litter and soil organic
matter turnover.

The initial decomposition of ﬁne roots and leaf litter involves
microbial immobilization of nitrogen (N) and phosphorous (P). The
resulting detritus, composed of ﬁne root necromass, leaf litter, microbial
populations, and secondary microbial products constitute pools of N and P
of major signiﬁcance to soil nutrient dynamics. The study of the
magnitude and dynamics of litter turnover, especially ﬁne roots, and its
effect on nutrient cycling has become an important consideration in
ecosystem productivity. The processes involved in litter decomposition,
however, have just begun to be understood.

We now realize that roots account for a large proportion of the carbon
(C), N and P budgets of terrestrial plant communities (Harris et al., 1977;
Vogt et al., 1986; Santantonio, 1989), however, we still do not understand
how important death and decay of roots are to the functioning of the soil

2

community. The dynamic nature of ﬁne root turnover has not been
addressed, especially the role of turnover in regulating microbial
populations and soil organic matter dynamics. The general requirements
of high yield Populus systems are known (Dickmann et al., 1988). To
sustain this level of production it is normally necessary to use substantial
amounts of fertilizer (Baker, 1977). By understanding the mineralization-
immobilization potential of the soil biomass and formation and decay of soil
organic matter we may be better able to sustain high production ecosystems
without major fertilizer inputs and minimal losses of nutrients to the

environment.

Fine Root Turnover

Fine root turnover has been measured in fewer than 30 forest stands
world wide (Santantonio and Grace, 1987). The inherently difficult task of
gathering and identifying ﬁne roots, along with inadequate methods of
estimating turnover, are reasons for this dearth of information (Gholz et
al., 1986; Santantonio and Grace, 1987; Raich and Nadelhoffer, 1989;
Santantonio, 1989; Fogel, 1990). Since no standard method exists,
researchers have used a variety of indirect and direct ways to estimate ﬁne
root turnover. Direct methods of coring and excavation include measuring
changes in the live standing crop of roots (Harris et al., 1977 ; McClaugherty
et al., 1982) or measuring both live and dead roots (Persson, 1979; Keyes and
Grier, 1981; Gholz et al., 1986). Indirect methods have been based on N
balance and mineralization budgets (N adelhoffer et al., 1985) and
carbohydrate concentrations of ﬁne roots (Marshal and Waring, 1985).

Root biomass production in coniferous forests was historically

estimated as about half that of leaf litter production (Gray and Williams,

3

1971; Whittaker and Marks, 1975). Fine root production and turnover are
rarely included in these data. Recent investigations have shown that ﬁne
root production in hardwood and coniferous forests account for 1.5 to 5
times as much detritus as leaf litter (Edwards and Harris, 1977; Grier et
al., 1981; Nadelhoffer et al., 1985). However, other authors report that the
contribution of ﬁne roots to detritus is only 3-29% of total detrital input
(Keyes and Grier, 1981; Joslin and Henderson, 1987; Santantonio, 1989,
Pregitzer et al., 1990). Site quality and methodology must be taken into
account when comparing data on ﬁne root production. Fine root turnover
and associated decomposition processes will have signiﬁcant impact on net
primary production and nutrient cycling in hybrid poplar systems
regardless of the uncertainty associated with measuring below ground
production.

The decomposition of ﬁne roots and leaf litter exhibit a distinct
pattern encompassing three phases (Fogel and Cromack, 1977; Berg and
Staaf, 1981; McClaugherty et al., 1984; Berg, 1988). During the ﬁrst phase of
decay, nutrients are mineralized through the decomposition of narrow C:N
cytoplasmic constituents. The ﬁrst phase is short in sequence (days to
months) and leads to the second phase of immobilization. The second
phase lasts up to 3 years and is characterized by slow weight loss attributed
to wide C:N substrates such as lignocellulose, secondary microbial
products and the accumulation of polymerized polyphenols. Following the
second phase, a third phase of mineralization begins and continues until
the recalcitrant components are incorporated into stabilized organic matter
(Aber and Melillo, 1980; McClaugherty et al., 1984; Berg, 1988).

Retranslocation of leaf nutrients is a mechanism whereby trees

conserve nutrients and bypass the root uptake pathway and soil nutrient

4

cycling processes. Poplars can retranslocate up to 70% of N and 50% of P
from leaves prior to senescence (Baker and Blackmon, 1977). However,
little is known of what happens to nutrients in senescing ﬁne roots.
Evidence from ﬁne root studies of Pinus radiata suggests that no
conservation of nutrients occurs (Nambiar, 1987 ). Similar results were
obtained for hybrid poplars (Horwath et al., 1992). Nutrients lost through
root turnover must be recycled through decomposition and subsequent
uptake. Studies have shown that 22-75% of N uptake by trees was allocated
to ﬁne root production (McClaugherty et al., 1982; Meier et al., 1985;
Nadelhoffer et al., 1985). Potentially large amounts of nutrients are
involved in ﬁne root mortality and decomposition.

Litter decomposition and subsequent microbial immobilization is an
important means of conserving nutrients in forest systems (Aber and
Melillo, 1982). This can be especially crucial in preventing leaching losses
following disturbances such as whole tree harvesting, a method employed
in many intensively-cultured forest plantations (Borman et al., 1979;
Vitousek and Melillo, 197 9; Vitousek and Matson, 1985). Litter turnover is a
continuing process taking years to complete. As a result, essential
nutrients will be immobilized during stand establishment and
regeneration, a time when nutrients are in great demand. Fertilization
will be required to compensate for nutrient demand until large detrital and
organic matter pools are established. When these pools are large enough,
they should be able to maintain and buffer soil fertility with minimal
fertilizer inputs. "Organic farming" is based on this concept and recent
evidence demonstrates how successful management of soil organic matter
can result in high agricultural yield with little or no fertilizer input (Doran
et al., 1988).

Microbial Biomass and Soil Organic Matter Dynamics

Controls on the mechanisms of microbial mineralization of organic
matter and associated nutrients are being investigated from a genetic and
cellular perspective (Payne, 1980; North, 1982; Dashman and Stotzky, 1986).
But it has proven difficult to translate in vitro information at the enzyme,
cellular and organismal level into an ecosystem context. Mechanisms
controlling short-term nutrient dynamics cannot be understood or
predicted by summing or averaging the characteristics of heterogeneous
systems. Therefore, conceptual models have been used to simulate C, N
and P dynamics and to divide soil organic matter components into
biologically meaningful fractions.

J ansson (1958) proposed two decomposable organic fractions which
he called active and passive. The active fraction is the most biologically
signiﬁcant pool in soil nutrition while the passive fraction is composed of
recalcitrant and physically protected organic matter. We now consider that
the active fraction contains the soil microbial biomass and easily
mineralizable organic matter. Paul and J uma (1981) further advanced the
concept of organic matter compartmentalization into ﬁve soil N fractions
consisting of microbial biomass N, active N, metabolite N, stabilized N and
old N. The investigators based these pools on decay rates determined by
using tracers and isotopic dilution techniques. The stabilized N and old N
turnover rates were based on carbon dating information (Martel and Paul,
1974).

Models which predict the rates and fluxes of soil nutrients are
coupled to the need of the soil microbial biomass to obtain energy from the
C-H and C-C bond of organic compounds (McGill and Cole, 1981; Frissel

6

and van Veen, 1980; van Veen et al., 1981). The mineralization of N is a
consequence of the microbial metabolism of organic compounds containing
N. The C:N ratio of the substrate is used as an indicator to determine the
potential mineralizable amounts of N. These models imply that the need
for C ultimately controls the fate of N in soil. Other researchers have
suggested that the quality of the organic matter (Bosatta and Staaf, 1982)
and clay effects (Dashman and Stotzky, 1986), rather than C to N ratios,
control N availability. Melillo et al. (1982) suggested this concept for forest
systems when they related the decomposition of leaf litter to lignin/N ratio.

Objectives and Goals

The current study was designed to address and enhance the
understanding of organic matter dynamics in ecosystems managed for
high biomass output such as short-rotation intensive-culture forestry. This
requires information on leaf and ﬁne root litter turnover, plant-soil
interactions, nutrient cycling and soil organic matter formation and
decomposition. Carbon entering the soil from the plant biomass is the
primary ecosystem process driving the cycling of nutrients. The production
and turnover of ﬁne roots is an important C input to soil, however, the
magnitude and importance of root derived C to nutrient cycling processes is
vaguely understood.

The amount and quality of plant C and nutrients returned to soil will
determine the mineralization potential of the soil biomass. It is therefore
important to establish the role of ﬁne root necromass in regulating the
cycling of nutrients and the formation of soil organic matter. The

understanding of these ecosystem processes will enable us to manipulate

 

7

the dynamic nature of soil organic matter formation in a way that sustains
productivity and minimizes chemical fertilizer inputs.

The current study was designed to address the uncertainties
associated with the production and decay of ﬁne roots and the potential

impact on nutrient cycling and long-term soil organic matter maintenance.

The general objectives of this study were the following:

(i) To evaluate the relative role of ﬁne root production and
decomposition in the nutrient cycling process.

(ii) Determine the C and N ﬂux through leaf litter, ﬁne root litter, soil
microbial biomass and soil organic matter fractions of hybrid poplar
plantations.

(iii) Determine the impact of litter decomposition and the
simultaneous mineralization-immobilization of N on a short-term and
long-term basis.

(iv) Relate soil biomass activity and substrate availability to patterns
of plant allocation.

The rationale for these objectives lies in the premise that ﬁne root
necromass will decompose at a slower rate than leaf litter and will exhibit
minimal retranslocation of cytoplasmic constituents before death. This will
lead to less immobilization of nutrients than leaf litter during the
decomposition process. As a result, decomposing ﬁne roots will have less
associated microbial biomass and form less stabilized organic matter than
is formed by leaf litter decomposition.

The determination of plant residue turnover, microbial biomass

activity and soil organic matter formation was accomplished using tracers

8

(14C and 15N). The use of tracers provided a quantitative estimate of above-
and below-ground production and its contribution to nutrient cycling and

soil organic matter formation.

Overview of Chapters
Chapter 2

Chapter 2 reports the results of a pilot study detailing the
development of a method to label Populus trees with 15N without physically
disturbing the soil. The labeling of root and shoot tissue by injecting N into
the vessel elements of the stem was useful in measuring the ﬂux of N from
these litters into soil organic matter fractions. The 15N was preferentially
distributed among age classes of leaves and size classes of roots. The
results from this study proved interesting in that the sink strength of the
different tree tissues could be established. The movement of 15N into ﬁne
root necromass was established by sorting live and dead roots over time.
These results demonstrated the ability to measure the contribution of root

litter to N cycling processes in situ.

Chapter 3

Chapter 3 describes the method used to radiolabel two-year-old hybrid
poplars with 14C02 in the ﬁeld. The combination of 15N injection and 14C
labeling provided a dual tracer approach for studying the fate of C and N in
this system. Trees were labeled on July 19 and September 5, 1990 to
encompass distinct seasonal growth and storage patterns exhibited by this
clone. The speciﬁc objectives addressed in this chapter include: (i) the
measurement of the initial fate of tracers (14C and 15N) two weeks after the

dual labeling; (ii) characterization of 14C respiration from the root system;

9

(iii) description of sampling protocols for tree components and soil in the
ﬁeld; (iv) discussion of the laboratory sample preparation and analytical

determinations.

Chapter 4

Chapter 4 examines the results from all samplings of trees labeled on
July 19 and September 5, 1990. The trees were sampled two weeks after
labeling, in November of 1990 and one year after labeling. Labeled leaf litter
was placed onto unlabeled tree plots in a litter exchange experiment to
determine the role of leaf decomposition and root turnover in nutrient
cycling processes. The litter exchange plots were sampled over the next two
years. Speciﬁc results addressed in this chapter include: (i) estimates of
root turnover by the dilution of labeled C with unlabeled C; (ii) the long-
term fate of 14C and 15N within tree components from the second to third
year of growth; (iii) the potential magnitude of leaf and root derived C input
to soil; (iv) assessing the role of leaf and root derived C in the maintenance

of soil organic matter fractions.

Chapter 5

Chapter 5 examines the partitioning of 14C-photosynthate within the
different chemical fractions of the tree. The chemical fractions included
lipids, sugars, protein, starch and structural residues. Structural residues
were considered to include lignin and the various polysaccharides forming
pectin, hemi-cellulose and cellulose. Speciﬁc results addressed in this
chapter include: (i) characterizing the long-term fate of radiolabeled C
within the different chemical fractions; (ii) exploring the relationships of

translocation and storage of C during different periods of growth and

10

dormancy; and (iii) assessing the importance of reserve C to the subsequent

growth of roots during the next growing season.

Chapter 6

Chapter 6 examines the dynamics of C and N in the soil microbial
biomass and soil organic matter fractions of this hybrid poplar plantation.
Long-term laboratory soil incubations were done on the last two samplings
of labeled trees and ﬁrst two samplings of leaf litter plots to determine the
movement of C and N into the microbial biomass, labile soil C and organic
matter pools of intermediate resistance. Potentially mineralizable C and
turnover time of soil C pools were derived by applying mathematical models
to C mineralization data. Speciﬁc objectives of this chapter include: (i)
determining the ﬂux of C and N to microbial biomass from leaf and root
turnover; (ii) assess the ﬂux of C and N from tree components labeled at
different times of the growing season; (iii) quantify the stabilization of C

into various fractions of soil organic matter.

BIBLIOGRAPHY

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relative contributions of organic matter and nitrogen to forest soils.
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Aber, J. D. and Melillo, J. M. 1982. Nitrogen immobilization in decaying
hardwood litter as a function of initial nitrogen content. Can. J. Bot.
60:2263-2269.

Baker, B. B. and B. G. Blackmon. 1977. Biomass and nutrient accumulation
in a cottonwood plantations-the ﬁrst growing season. Soil Sci. Am. J.
41:632-636.

Berg, B. 1988. Dynamics of nitrogen (15N) in decomposing Scots pine (Pinus
sylvestris) needle litter. Long-term decomposition in a Scots pine
forest. VI. Can. J. Bot. 66:1539-1546.

Berg, B. and H. Staaf. 1981. Leaching, accumulation and release of
nitrogen in decomposing forest litter. Ecological Bulletins
(Stockholm) 33:163-178.

Borman, F. H., Likens, G. E. and Melillo, J. M. 1979. Nitrogen budgets for
an aggrading northern hardwood ecosystem. Science 196:981-983.

Bosatta E. and H. Staaf. 1982. The control of nitrogen turn-over in forest
litter. Oikos 39: 143-151.

Dashman, T. and G. Stotzky. 1986. Microbial utilization of amino acids and
peptide bound on homoionic montmorillonite and kaolinnite. Soil
Biology and Biochemistry. 18:5- 14.

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Dickmann, D. I., K. S. Pregitzer, and P. V. Nguyen. 1988. Net assimilation
and photosynthate allocation of Populus clones grown under short-
rotation intensive culture: Physiological and genetic responses

regulating yield. Annual progress report, Subcontract No. 86x-
959036, USDOE.

Doran, J. W., Frasser, D. G., Culik, M. N., and Liebhardt, W. C. 1988.
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on the soil microbial processes and nitrogen availability. American
J. of Alternative Agriculture. 2: 99-106.

Edwards, N. T. and Harris, W. F. 1977. Carbon cycling in a mixed
deciduous forest ﬂoor. Ecology 58:431-437.

Fogel, R. 1990. Root turnover and production in forest trees. HortScience
25:270-273

Fogel, R. and Cromack, K. jr. 1977. Effect of habitat and substrate quality on
Douglas ﬁr litter decomposition in Western Oregon. Can. J. Bot.
55:1632-1640.

Frissel, M. J. and J. A. van Veen. 1980. Simulation model of the behavior of
N in soil. In: Simulation of nitrogen behavior of soil-plant systems.
Frissel, M. J. and J. A. van Veen (ed.) Centre for agriculture pub.
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Gholz, H. L., L. C. Hendry, and W. P. Cropper jr. 1986. Organic matter
dynamics of ﬁne root in plantations of slash pine (Pinus elliotti) in
North Florida. Can J. For. Res. 16:529-538.

Gray, T. G. R. and Williams, S. T. 1971. Soil micro-organisms. University
Reviews in Botany, Oliver & Boyd, Edinburgh.

Grier, C. G., K. A. Vogt, M. R. Keyes, and R. L. Edmonds. 1981. Biomass
distribution and above- and below-ground production in young and
mature Abies amabilis zone ecosystems of the Washington Cascades.
Can. J. For. Res. 11:155-167.

13

Harris, W. F., R. S. Kinerson, and N. T. Edwards. 1977. Comparison of
belowground of natural deciduous forest and loblolly pine
plantations. Pedobiologia. 17:369-381.

Horwath, W. R., E. A. Paul and K. S. Pregitzer. 1992. Injection of nitrogen-
15 into trees to study nitrogen cycling in soil. Soil Sci. Soc. Am.
56:316-319.

Ingstad, T. and G. I. Agren. 1984. Fertilization for long-term maximum
production. In K. Perttu (ed.): Ecology and management of forest
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Research, Swedish Univ. Agricultural Sciences, Report. 15: 155-165.

Jansson, S. L. 1958. Tracer studies on nitrogen transformations in soil.
Ann. Roy. Agric. Coll. Sweden. 24:101-361.

Joslin, J. D. and Henderson, G. S. 1987. Organic matter and nutrients
associated with ﬁne root turnover in a white oak stand. Forest
Science 33: 338-354.

Keyes, M. R. and C. C. Grier. 1981. Above- and below-ground net production
in 40-year old Douglas-ﬁr stands on low and high productivity sites.
Can J. For. Res. 11:599-605.

Marshall, J. D. and R. H. Waring. 1985. Predicting ﬁne root turnover by
monitoring root starch and soil temperature. Can. J. For. Res.
15:791-800.

Martel, Y. A. and E. A. Paul. 1974. The use of radiocarbon dating of organic
matter in the study of soil genesis. Soil Sci. Am. Proc. 38:501-506.

McClaugherty, C. A., J. D. Aber, and J. M. Melillo. 1982. The role of ﬁne
roots in the organic matter and nitrogen budgets of two forested
ecosystems. Ecology. 63:1481-1490.

McClaugherty, C. A., J. D. Aber, and J. M. Melillo. 1984. Decomposition
dynamics of ﬁne roots in forested ecosystems. Oikos 42:378-386.

14

McGill, W. B. and Cole, C. V. 1981. Comparative aspects of organic C, N, S
and P cycling through organic matter during pedogenesis.
Geoderma 26:267-286.

Meier, C. E., Grier, C. C. and Cole, D. W. 1985. Below- and above-ground N
and P use by Abies amabilis stands. Ecology 66:1928-1942.

Melillo, J. M., Aber, J. D. and Muratore, J. F. 1982. Nitrogen and lignin
control of hardwood leaf litter decomposition dynamics. Ecology
63:621-626.

Nadelhoffer, K. J., J. D. Aber, and J. M. Melillo. 1985. Fine roots, net
primary production, and soil nitrogen availability: a new hypothesis.
Ecology. 66: 1377-1390.

Nambiar, E. K. S. 1987. Do nutrients retranslocate from ﬁne roots. Can. J.
For. Res. 17:913-918.

North, M. J. 1982. Comparative biochemistry of the proteinases of
eukaryotic microorganisms. Microbiol. Rev. 46:308-340.

Paul, E. A., and N. G. Juma. 1981. Mineralization and immobilization of
nitrogen by microorganisms. In Terrestrial nitrogen cycles. F. E.
Clark and T. Rosswall (ed.) Ecol. Bull. (Stockholm) 33:179-195.

Payne, J. W. Regulation of peptide transport in bacteria. 1980. In J. W.
Payne (ed.) Microorganisms and nitrogen sources. p. 335-358. John
Wiley and Sons, Ltd. New York.

Persson, H., 1979. Fine-root production, mortality and decomposition in
forest ecosystems. Vegetatio. 41:101-109.

Pregitzer, K. S., D. I. Dickmann, R. Hendrick, and P. V. Nguyen. 1990.
Whole-tree carbon and nitrogen partitioning in young hybrid poplars.
Tree Physiology. 7:79-93

15

Santantonio, D. 1989. Dry-matter partitioning and ﬁne-root production in
forests--New approaches to a difficult problem. pp. 57-72. In:
Biomass production of fast-growing trees. Pereira, J. S. and J. J.
Landsberg (eds.) Kluwer Academic Publishers.

Santantonio, D. and J. C. Grace. 1987. Estimating ﬁne-root production and
turnover from biomass and decomposition data: a compartment-ﬂow
model. Can. J. For. Res. 17:900-908.

Van Veen, J. A., McGill, W. B., Hunt, H. W. Frissel, M. J. and Cole, C.V. .
1981. Simulation models of the terrestrial nitrogen cycle.- In Clark,
F. E. and Rosswall, T. (eds.), Terrestrial Nitrogen Cycles, Ecosystem
Strategies and Management Impacts. Ecol. Bull. (Stockholm) 33:25-
48.

Vitousek, P. M. and Melillo, J. M. 1979. Nitrate losses from disturbed
forests: patterns and mechanisms. For. Sci. 25:605-619.

Vitousek, RM. and Matson, RA. 1985. Disturbance, nitrogen availability,
and nitrogen losses in an intensively managed loblolly pine
plantation. Ecology 66: 1360-1376.

Vogt, K. C., Grier, C. C., and Vogt, D. J. 1986. Production, turnover, and
nutritional dynamics of above- and belowground detritus of world
forests. In Macfadyen, A. and Ford, E. D. (eds.), Advances in
Ecological Research Vol. 15. Academic Press, New York, New York.

Whittaker, R. F. and Marks, P. L. 1975. Methods of assessing terrestrial
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productivity of the biosphere, Ecol. Stud. 14: 55-118.

Chapter 2

THE INJECTION OF NITROGEN-15 INTO TREES TO STUDY NITROGEN
CYCLING IN SOIL

ABSTRACT

A method was developed to introduce 15N directly into trees to study
internal N translocation and litter turnover. Leaf and root biomass were
labeled by injection of 15N directly into the vessel elements of hybrid
Populus trees during their second growing season. The 15N was uniformly
distributed throughout the canopy and root system. The rate and amount of
15N turnover from plant tissue can be determined by pool transfer or
through differences in plant 15N concentrations. The 15N was detected in
the dead root pool 8 weeks following injection, indicating root turnover.
Results demonstrate the ability to measure the contribution of ﬁne root litter

to N cycling processes without disturbing the soil environment.

16

17
INTRODUCTION

The application of 15N either as fertilizer or plant residue to soil has
made it possible to study nitrogen transformations and processes in both
agricultural and forest systems (Jansson, 1958; Amato and Ladd, 1980). By
following 15N through plant-soil N pools and then calculating isotopic
dilution, a relatively accurate picture of the fate of N added as fertilizer has
been determined (Vitousek and Andariese, 1986; Voroney et al., 1989).
However, there is a lack of information concerning the direct contribution
of N from plant roots to soil N pools. Further, most dilution techniques
disturb either the soil (by introducing labeled residues) or the N pool size.

Successful injection of 15N directly into the plant allows for the study
of N ﬂux ﬁom plant to soil without introducing the label into the soil. Thus,
15N can be followed from plant to litter and ﬁnally to soil N pools. The rate
of N cycling from an in situ plant pool to various soil pools could then be
directly calculated, avoiding some of the problems associated with other
15N methodologies. However, discriminating N ﬂux and plant uptake of
mineralized 15N from previously labeled tissues continues to be a problem
associated with all 15N techniques.

The injection of 15N into trees was based on methods used for the
systemic introduction of nutrients, growth regulators, and pesticides
(Morris, 1951; Graham, 1954; Schreiber, 1969; Filer, 1973; Sterrett and
Creager, 1977; Vreeland et al., 1981; Schulert et al., 1988). Applications of
these methods have included the study of root distribution, interspeciﬁc
plant nutrient transfers, metabolism of growth regulators and pesticides,

and bioavailability and sink-strength of nutrients (Auerbach et al., 1964;

Rue
D01

éISM-D

18

Russell and Ellis, 1968; Woods and Brock, 1964; Sterrett and Hipkins, 1980;
Domir, 1980; Schulert et al., 1988).

The objective of this study was to examine the feasibility of labeling
tree biomass in situ for the study of soil N cycling associated with leaf and
ﬁne root litter turnover. Tracer 15N was injected into trees and carried in
the transpiration stream to the leaves. About two weeks to a month
following application the N was distributed throughout the plant. I report
the development of a method for studying the direct ﬂux of labeled N from
trees to the soil.

MATERIAL AND METHODS

Injection Procedure

Trees of a single Populus clone (P. x euramericana cv. Eugenei) in
their second season of growth were injected with 15N. The N was dissolved
in an artiﬁcial sap solution consisting of 5.0 mM KCl and 0.4 mM malic
acid adjusted to pH 5.4 with KOH (Dickson et al., 1985). The N was added as
15N-(NH4)2$O4 at levels equivalent to 5-10% of the total tree N. The volume
of the 15N solution injected was 100 mL. Unlabeled sap solution was used to
chase and ﬂush the labeled N from the severed vessel elements until uptake
had ceased. The solutions were sterilized by autoclaving at 120° C for 15
minutes or ﬁltering through a 0.2 pm Millipore ﬁlter to avoid introducing
pathogens directly into the tree.

A centered 6 mm diameter hole was drilled 75% of the way through a
5.0 cm diameter stem at 8.0 cm above ground level using a brad point wood
drill bit and rechargeable drill. The conﬁguration was found to be ideal for
trees this size, but may have to be adjusted according to the age and size of
different trees. Hole diameters of 2 mm have also been tested on 3-5 cm

di

tr;

we
in
ill

WE

19

diameter trees with success. The injection method relies on an active
transpiration stream and, therefore, irrigation may be necessary when soil
matrix potential is low. With species other than Populus the ideal protocol
may have to be obtained through experimentation. Populus is diffuse
porous, the injection method may have only limited value in ring porous
taxa.

The hole was ﬁtted with a modiﬁed Tygon tubing connector with an
inserted septum to create a tight seal (Figure 2.1). The cavity was then
ﬂushed to exclude air by connecting a syringe body (30 mL) ﬁlled with
artiﬁcial sap to one of two syringe needles inserted into the septum. Sap
was forced into the cavity until a steady stream of liquid was seen coming
from the second needle. The syringe needles were removed following the
ﬂushing procedure. A gravity-fed reservoir containing the 15N solution
was then connected by inserting the tracer delivery tube tipped with a
syringe needle into the septum (Figure 2.1). The sap reservoir was
suspended 1 m above the hole. Care must be exercised when switching
from the 15N solution to the chase solution to avoid introducing air into the
injection line. The entire operation from drilling the hole to connecting the

labeled N solution took no more than 45 seconds.

Field Protocol and Sampling

Each replicate tree was centered in a 1 m2 soil monolith by
trenching to a depth of 45 cm and wrapping the soil column with two layers
of plastic sheeting two months prior to 15N injection. Tree Set 1 contained 4
replicate trees and they were injected on July 13, 1988 with 500 mg of 39.0
atom % 15N-(NH4,)2SO4 which amounted to approximately 5% of the total

tree N based on previous work by Pregitzer et al. (1990). Tree Set 2 also

 

second year
growth

 

 

ﬁrst year
wtll

 

 

Figure 2.1. Diagram of the injection technique.

sap
reservou

21

contained 4 replicate trees and they were injected on September 12, 1988
with 500 mg of 98.0 atom % 15N-(NH4)2SO4. Soil cores to a depth of 45 cm
were taken from the 1 m2 soil monoliths surrounding each tree at two and
eight weeks after injection. Four cores were taken randomly from each of
the soil monoliths two weeks following injection. Five additional cores were
taken from each monolith eight weeks following injection. In order to avoid
dead roots created by extracting the ﬁrst set of cores, the cores extracted at
eight weeks were all randomly located near the periphery of each monolith.
Each tree was encased in commercial bird netting to trap falling leaves.
Abscised leaves were recovered daily from the trees throughout the growing
season until all leaves had fallen.

Roots were hand picked from soil cores sieved through a 3 mm mesh
screen. Roots were classiﬁed under magniﬁcation according to size and
age class. Sizes were determined on fresh roots with a microcaliper. The
classiﬁcation criteria included root color, texture, and tensile strength.
Fresh dead roots were characterized by a dark stele and fragile cortex. New
white ﬁne roots were characteristically light (white) in color with a smooth
cortex indistinguishable from the stele. Older brown roots had a brownish
ﬁbrous cortex and light stele that easily dissociated from the cortex.

Tissue Analysis

Leaf and root samples were dried in a convection oven at 60°C for 48
hours. Leaves were subsampled and ground in a household blender. The
ground leaf tissue was then ball milled using a rolling glass jar and steel
rod technique (Harris and Paul, 1989). Root samples were homogenized by
cutting into small fragments with a dissecting blade. Samples of known

mass were combusted in a biological sample converter (Roboprep, Europa

22
Scientiﬁc, Crewe, UK) to yield N2 and C02. The gases were then analyzed

with a continuous-ﬂow stable isotope ratio mass spectrometer for N and
15N (AN CA-MS, Europa Scientiﬁc, Crewe, UK) and thermal conductivity
detector for 002 (Carle, Fullerton, CA). Urea-15N (0.60571 atom % 15N)
and 44% carbon (as cellulose) Whatman #1 ﬁlter paper were used for

reference standards (Harris and Paul, 1989).

RESULTS AND DISCUSSION

The correct drill conﬁguration is necessary to avoid killing branches
and leaves as a result of severing too many vessel elements. Also,
accessing too few vessel elements can lead to N toxicity by delivering the N
solution to limited parts of the canopy. In a correct drill conﬁguration, the
injected N will ascend in the transpiration stream and accumulate in the
leaf biomass before equilibrating throughout the entire plant. Table 2.1,
shows no signiﬁcant aspect differences of the injected N or 15N 72 hours
after injection using Tukey's procedure for comparison of means at 0:005.
Following leaf fall, the leaves from Tree Sets 1 and 2 contained only 17%
and 18% of the applied 15N, respectively (Table 2.2). Most of the canopy N
was apparently conserved as has been shown before for this genotype
(Pregitzer et al., 1990).

The amount of 15N injected into a tree will depend on the plant or soil
pool of interest. We believe the maximum amount of 15N-(NH4,)2804 that
can be injected into a tree is approximately 5-10% of the total tree N content.
Our experience indicates that greater concentrations can be toxic to the
canopy. Also, the level of soil N availability may inﬂuence the amount of N

that can be injected. Existing high soil N levels followed by injection of 15N

Table 2.1. Mean leaf N concentrations and the distribution of 15N in the
most recently mature leaf of each branch according to crown aspect 3
days following injection of tree set 1. Standard error of the mean given
in parentheses.

 

 

 

Crown Atom %
Aspect g N kgjl 15N Excess§

N 3.64a (0.18) 0.93a (0.17)
NE 3.91a (0.21) 1.22a (0.24)
E 4.113 (0.22) 0.78a (0.18)
SE 3.93a (0.15) 0.85a (0.23)
S 4.28a (0.33) 1.05a (0.22)
SW 4.20a (0.17 ) 0.80a (0.24)
W 3.49a (0.14) 0.66a (0.17)
NW 3.41a (0.20) 0.89a (0.22)
§ Atom % 1‘5N excess is deﬁned as 15N above background (0.3663%).

Table 2.2. Summary of N and C concentrations and the recovery of 15N in

senescent leaves. The 15N recovered is based on the amount injected.
Standard error of the mean given in parentheses.

Atom % % TBN

 

 

_ N k 1 C k 1 15N Excess§ Recovered
Tree Set 1 1.38 (0.13) 36.70 (0.25) 0.39 (0.05) 17.2
Tree Set2 1.70 (0.05) 37.70 (0.32) 1.07 (0.11) 18.2

 

§ Atom % 15N excess is deﬁned asﬁlFN above background (0.3663%).

24

can lead to premature leaf abscission by a mechanism we do not
understand. This imposes limitations on the detection limits of 15N in soil
organic matter or microbial biomass pools, especially in young trees with
low total N contents. However, the use of several injections of 15N may
overcome limitations of a single injection.

The results of our initial experiments indicate that the enrichment of
the root biomass occurred within 5 days (data not shown) and was
stabilized about 2 weeks following injection, regardless of the time of
addition (Table 2.3). The increased enrichment of the dead roots in Tree Set
2 between 2 to 8 weeks indicates that some turnover of the live root pool
occurred during this period. Both tree sets exhibited a decrease in
enrichment of new white and older brown roots with time. The decrease in
enrichment was assumed to be associated with dilution from the uptake of
unlabeled soil N, growth of new roots, and turnover.

Using data from Table 2.3 for the dead root pool and current ﬁeld
studies on root production (Chapter 4), the potential atom % 15N excess of
soil microbial biomass N is 0.022%. The standard deviation of analysis of
our mass spectrometer is 0.0004 atom % 15N. Detectability of 15N will
depend on the rate of litter turnover and retention time of N in the microbial
biomass and soil organic matter, but appears feasible.

An alternative way to measure the transfer of 15N from plant to soil
is to determine the change in the percent of N in the plant biomass derived
from 15N over time (Table 2.4). This calculation is independent of the
amount and enrichment of the injected N (Rennie and Rennie, 1983). The
precision of this method can be increased by a complete harvest of the plant
biomass (root and shoot). The assumption of this method is that all 15N lost
from the plant through time zero is due to litter turnover and that all 15N

Table 2.3. Mean labeled N and N concentrations of different classes of roots
collected over time from tree sets 1 and 2. Standard error of the mean
given in parentheses. -

 

 

 

 

 

 

Atom % 15N Excess§ g N kg—l
Root Root
Class Size 2 wkl 8 wk 2 wk 8 wk
New < 0.5 0.21 (0.03) 0.08 (0.01) 1.8 (0.0) 2.8 (0.2)
White 0.5-1.0 0.38 (0.10) 0.18 (0.04) 2.0 (0.3) 1.5 (0.1)
Older <0.5 0.19 (0.03) 0.12 (0.02) 1.7 (0.0) 2.8 (0.1)
Brown 0.5-1.0 0.29 (0.03) 0.19 (0.02) 1.0 (0.1) 1.7 (0.1)
1.0-3.0 0.30 (0.04) 0.21 (0.02) 1.0 (0.1) 1.4 (0.1)
>3.0 0.28 (0.02) 0.23 (0.3) 0.8 (0.1) 1.0 (0.3)
Dead <0.5 na na 0.07 (0.01) na na 2.3 (0.1)
0.5-1.0 na na 0.07 (0.02) na na 1.3 (0.1)
1.0-3.0 na na 0.18 (0.08) na na 1.6 (0.1)
W
Atom % 15N Excess g N kgl
Root Root
Class Size 2 wk 8 wk 2 wk 8 wk
New < 0.5 0.19 (0.09) 0.17 (0.06) 2.0 (0.1) 1.6 (0.1)
White 0.5-1.0 0.55 (0.13) 0.30 (0.10) 1.6 (0.2) 1.5 (0.1)
Older <0.5 0.15 (0.04) 0.14 (0.04) 1.6 (0.1) 1.6 (0.1)
Brown 0.5-1.0 0.54 (0.11) 0.21 (0.05) 1.1 (0.1) 1.4 (0.1)
1.0-3.0 0.93 (0.21) 0.22 (0.22) 0.1 (1.8) 0.2 (0.2)
> 3.0 0.70 (0.20) 0.55 (0.19) 1.0 (0.1) 1.3 (0.1)
Dead < 0.5 0.04 (0.02) 0.21 (0.09) 1.6 (0.1) 1.6 (0.1)
0.5-1.0 0.01 (0.01) 0.51 (0.17) na na 1.4 (0.2)

 

§ Atom % 15N excess is deﬁned as 15N above background (0.3663%).

1' Time since injection of 15N.

na not available

Table 2.4. Percentage of N in various root classes derived from 15N two and
eight weeks after injection. l'

 

Time Since Injection 2m; 8%
Tree Tree Tree Tree
Set 1 Set 2 Set 1 Set 2 Change Y
New WE'te Roots
<05 m 0.51 0.19 0.21 0.17 -0.18
0.5-1 0.97 0.56 0.46 0.31 -0.35
Older Brown Roots
<0.5 0.49 0.15 0.31 0.14 -0.10
0.5-1 0.74 0.55 0.49 0.21 -0.29
1.0-3.0 0.77 0.71 0.54 0.45 -0.25
> 3.0 0.72 0.71 0.59 0.56 -0.14
Dead Roots
<0.5 na 0.01 0.18 0.21 +0.18
0.5-1 na na 0.46 0.52 +0.49
1.0-3.0 na 0.04 0.18 0.17 +0.18

 

1' Calculated according to Rennie and Rennie (1983).
Y Change calculated from the average of tree sets one and two at 8 weeks
na not available

27

lost has entered the soil N pool. Obviously, this assumption has limitations
in a ﬁeld situation.

Nitrogen derived from the 15N is similar for both sets of trees 2 and 8
weeks following injection, with the exception that more label was found in
the new white root pool of Tree Set 1 (Table 2.4). The dead root pool for both
tree sets at 8 weeks contains similar amounts of N derived from the injected
15N. Based on the July and September injections, the data indicate the ﬂux
of 15N to the root system is independent of the time of injection. It may be
that 15N can be injected at different times during the growing season to
label the root system. This notion, currently under study, may be useful in
studying the temporal aspects of N cycling. The data also demonstrate the
movement of label from live to dead roots and, therefore, the potential to
measure root turnover.

Analysis of the total root biomass by excavation would be necessary to
calculate total 15N distribution and to obtain the greatest reproducibility
(Waremburg and Paul, 1973; Harris and Paul, 1989). Though the degree of
sampling was limited in this study, it aptly demonstrates the potential of
tracing 15N injected into the stem throughout the tree and into litter pools.
The injection of tracer N provides a relatively easy means to transfer 15N
into the tree and promises to be an effective way to study N transformations
and cycling in undisturbed tree-soil systems. The most powerful aspect of
the injection method is the ability to measure the contribution of a
functioning root system to soil N processes. The potential to label other soil

N pools clearly exists.

Au

Dic

Dor

Gra

Her

BIBLIOGRAPHY

Amato, M. and J. N. Ladd. 1980. Assay for microbial biomass based on
ninhydrin-reactive nitrogen in extracts of fumigated soils. Soil
Biology and Biochemistry 20:107-114.

Auerbach, S. I., J. S. Olson, and H. D. Waller. 1964. Landscape
investigations using 137Ce. Nature. 201:761-764.

Dickson, R. E., T.C. Vogelmann, and P. R Larson. 1985. Glutamine
transfer from xylem to phloem and translocation to developing leaves
of Populus deltoides. Plant Physiol. 77: 412-417.

Domir, S. C. 1980. Metabolism and distribution of 14C-maleic hydrazide and

14C-daminozide injected into red oak. J. Amer. Soc. Hort. Sci. 105:
678-680.

Filer, T. H. Jr. 1973. Pressure apparatus for the injecting chemicals into
trees. Plant Disease Reporter. 57: 338-341.

Graham, B. F. 1954. A technique for introducing radioactive isotopes into
tree stems. Ecology. 35: 415.

Harris, D. and E. A. Paul. 1989. Automated analysis of 15N and 14'0 in
biological samples. Comm. in Soil Sci. and Plant Anal. 20: 935-947.

Jansson, S. L. 1958. Tracer studies on nitrogen transformations in soil.
Ann. Roy. Agric. Coll. Sweden. 24:101-361.

Morris, R. F. 1951. Tree injection experiments in the study of birch dieback.
For. Chron. 27: 319-329.

Pregitzer, K. S., D. I. Dickmann , R. L. Hendrick , and P. V. Nguyen. 1990.
Whole-tree carbon and nitrogen partitioning in young, fast-growing
poplars. Tree Physiology 7: 79-93.

E

81‘

hit

29

Rennie, R. J. and D. A Rennie. 1983. Techniques for quantifying N2 ﬁxation
in association with nonlegumes under ﬁeld and greenhouse
conditions. Can. J. Microbiol. 29: 1022-1035.

Russell, R. S. and F. B. Ellis. 1968. Estimation of the distribution of plant
roots in soil. Nature. 217: 582-583.

Schreiber, L. R. 1969. A method for the injection of chemicals into trees.
Plant Disease Reporter. 53: 764-765.

Schulert, A., A. Zeind, and W. Darby. 1988. Stem injection: the most
efﬁcient labelling procedure for bioavailability investigations. 72nd
Annual Meetings (Fed. Am. Soc. Exp. Biol.). 2: A424. Abstract 747.

Sterrett, J. P. and P. L. Hipkins. 1980. Response of apple buds to pressure
injection of abscisic acid and cytokinin. J. Amer. Hort. Sci. 105: 917-
920.

Sterrett, J. P. and R. A. Creager. 1977. A miniature pressure injector for
deciduous seedlings and branches. HortScience. 12: 156-158.

Vitousek, P. M. and S. W. Andariese. 1986. Microbial transformations of
labeled nitrogen in a clear-cut pine plantation. Oecologia 68: 601-605.

Voroney, R. P., E. A. Paul, and D. W Anderson. 1989. Decomposition of
wheat straw and stabilization of microbial products. Can. J. Soil Sci.
69: 63-77.

Vreeland, P., T. Lugaski, H. Vreeland, and E. Kleiner. 1981. A new isotope
injection procedure for use in forest ecosystems. Environmental and
Experimental Botany. 21: 267-268.

Waremburg,F. R. and E. A. Paul. 1973. The use of 14002 canopy techniques
for measuring carbon transfer through the plant-soil system. Plant
and Soil 38: 331-345.

WOOds, F. W. and K Brock. 1964. Interspeciﬁc transfer of 45Ca and 32P by
root systems. Ecology 45: 886-889.

Chapter 3

14C AND 15N PARTITIONING IN TREE-SOIL SYSTEMS

ABSTRACT

Hybrid poplars were labeled with 14C and 15N to determine the
partitioning of C and N during the second year of growth in the ﬁeld. Eight
trees were pulsed-labeled in a large Plexiglas chamber in July and
September, 1990. Levels of 14C02 and climate within the chamber were
controlled. Nitrogen-15 was stem injected into the trees. Three trees were
harvested two weeks following each labeling period. Carbon dioxide
assimilation values for whole-tree canopies averaged 3.8 umol 002 m'2 s’1
in July and 6.2 umol 002 m'2 s'1 in September. Above-ground dark
respiration amounted to 18% and 24% of net assimilation in July and
September. Carbon-14 root respiration peaked 2 days after labeling and
after 2 weeks 14C had stabilized below-ground, demonstrating that these
trees primarily depend on current photosynthate for growth and
maintenance during the growing season. Root respiration averaged 20% of
total soil respiration in both July and September. In July, 80% of the
recovered 14*C was located above-ground and closely resembled the weight
distribution of the growing shoot. By September, 38% of the recovered 14C
was found in the root system and closely resembled the weight distribution
of the different size classes of roots. The root system contained 8% of the
injected 15N in July and 20% in September.

31
INTRODUCTION

The productivity of trees is inﬂuenced by the partitioning of carbon
and nitrogen. Allocation of C and N in above- and below-ground tree
components affects harvest index and the performance of a species in a
forest stand (Isebrands, 1982). In intensively cultured plantations the
measurement of photosynthesis, dry matter production and respiration are
important in interpreting how stands function. However, the majority of
harvest and 002 exchange studies have been conducted on the shoot
system, while the below-ground plant-soil system has received little
attention (Tranquillini, 1963; Kinerson et al., 1977; Agren et al., 1980;
Linder and Troeng, 1981; Friend et al., 1991) Nonetheless, below-ground
production can account for greater than 50% of primary production in
forests (Harris et al., 1975; Olsen, 1975; Persson, 1979; Keyes and Grier,
1981; Fogel, 1990, Hendrick and Pregitzer, 1993). The ﬂux of C below-
ground represents a considerable cost to trees and should be considered
when managing forests (Dickson, 1989).

Detailed studies of above- and below-ground C ﬂow and N distribution
within the tree-soil system are possible with the use of isotopes (Dickson,
1989; Horwath et al., 1992). Previous 14C studies have estimated whole-tree
C allocation by pulse-labeling potted trees, individual leaves or branches
(Schier, 1970; Isebrands and Nelson, 1983). The advantage of isotope
methods over harvest and 002 exchange methods is that growth and
decomposition which occur simultaneously can be isolated to some extent
by following long-term isotope flux and dilution below-ground
(W arembourg and Paul, 1973). The general objectives of my study were to
label whole-trees in the ﬁeld with 14C and 15N and to quantify whole-tree C

32

and N allocation. The experimental system was designed to maintain COz
concentration and climate at ambient levels.

I was speciﬁcally interested in determining seasonal differences in:
(1) the respiratory cost of the shoots and roots; (2) the fate of 14C and 15N in
whole trees. In particular, I was interested in evaluating the use of 14C to
measure root-soil respiration. Seasonal rhythms in growth and the
allocation of carbohydrates and nitrogen reserves have been documented in
Populus (Isebrands and Nelson, 1983; Pregitzer et al., 1990; Nguyen et al.,
1990), but budgets developed by labeling whole-trees greater than 3 m tall
have never been reported. It is my belief that patterns of C and N allocation
have ecological signiﬁcance and that, if ignored, seasonal changes in
source-sink relationships can greatly confound the interpretation of
research and the application of cultural practices. This chapter represents
my ﬁrst attempt to study whole tree gas-exchange and C and N allocation in

Populus-soil systems using a dual isotope approach in the ﬁeld.

MATERIALS AND METHODS

Trees were grown at the Kellogg Biological Station, ("Long-Term
Ecological Research" National Science Foundation), Michigan State
University, in southwest Michigan. The soil series is Kalamazoo (ﬁne
loamy, mixed, mesic, Typic Hapudalf). The site was prepared by
moldbroad plowing and disking. Cuttings of Populus euramericana cv.
Eugenei 10 cm in length were planted on a 2 x 1 m spacing in the spring of
1989. Later in the ﬁrst growing season, groups of eight trees were selected
for uniform height and diameter characteristics. An undisturbed soil block

(1 m3) surrounding each tree was trenched using a narrow ditching

33
machine (Ditchwitch, Howell, MI). Plywood dividers were inserted into the

trenches and the vertical faces of the monoliths were wrapped in 0.18 mm
vinyl before back-ﬁlling.

The following summer (1990), injection of 15N into trees was initiated
ﬁve days prior to 1‘er exposure. The 15N (100 mg of 98% enriched
(15NH4)2SO4 was injected as an artiﬁcial sap solution into the vessel
elements of the stem tissue (Horwath et al., 1992). Following the 15N
injection, the vinyl sheeting of each soil block was sealed against the
plywood structure with foam sealant (Insta-Foam Products, Inc., Marietta,
GA). Polyvinyl Chloride sheets (3.2 mm) were sealed to the top of the
plywood structure with silicone caulking to create a headspace (15 cm tall x
1 m x 1 m) above the soil surface of each soil block. Tree stems were
wrapped with modeling clay, polyethylene plastic and duct tape before being
sealed to the PVC plates with silicone caulking. Tygon tubing (1.3 cm I.D.)
was routed from each headspace to a C02 trap containing 500 mL 4.0 M
NaOH. The headspace atmosphere was drawn at a rate of 12 L min'1 with
a diaphragm pump (Cole Parmer Inst. Co., Chicago, IL) through an
aeration stone in each base trap to remove C02. Base traps were sampled
daily to determine below-ground CO2 and 14C respiration during and
following the radiolabeling procedure. Subsamples of 1 mL were titrated to
a phenolthalein endpoint (pH 7.0) using excess BaC12 and 0.1 M HCl to
determine total C02-C. Scintillation cocktail (10 mL of Safety Solve,
Research Products International Corp., Mount Prospect, IL) was added to
1.0 mL subsamples from the base traps and analyzed for 14C on a liquid
scintillation spectrometer (Packard Instrument Co., Downers Grove, IL).

A Plexiglas chamber (3.2 m tall x 3 m wide x 4 m long) was

assembled over the eight trees and below-ground partitions (Figure 3.1).

PLEXIGLAS
CHAMBER

  
  

3.2 m AIR
PUMP

COOLING
WATER

1m - -
9‘3.
.-'.';'

Figure 3.1. Schematic diagram of the 14C exposure system used to label
eight, two-year-old whole poplar trees. The trees were approximately
three meters tall when labeled.

35
The chamber was sealed to the plywood structure and PVC sheets with
silicone caulking. A 14C02 generator was connected to a diaphragm pump
(GAST, Benton Harbor, MI) to circulate 14C02 into the chamber. The
14CO 2 generator consisted of a vessel containing 1.0 L of concentrated
H2804 to which was attached a reservoir containing 6.0 moles of Na214CO3
solution. The carbonate solution was dispensed into the acid via a solenoid
valve (Skinner, New Brunswick, CT) as required to maintain CO2 levels
equivalent to the ambient atmosphere outside the chamber. The acid-

carbonate mixture was continuously agitated using a magnetic stirrer to

facilitate the liberation of 14C02.

Chamber CO2 level was monitored with an Infra Red Gas Analyzer,
IRGA (Analytical Development Co. Ltd., Hoddesdon, England).
Temperature sensors (Omega Engineering, Inc., Stamford, CT) were
installed to monitor the soil surface, chamber air, and ambient air
temperatures. Photon ﬂux density was measured during the labeling
periods with a radiometer/photometer (LI-COR, Inc., Lincoln, NE). The
IRGA, solenoid valve and temperature sensors were connected to a data
acquisition and control device (Remote Measuring Systems, Inc., Seattle,
WA) that was interfaced by serial connection to a portable computer.
Computer software was written in Quick Basic (Microsoft, Corp. Redmond,
WA) to control the level of CO2, monitor temperatures and collect data. The
chamber temperature was controlled by forcing air through a water cooled
heat exchanger (self customized). The air turbulence created by the heat
exchanger was sufﬁcient to mix the 14C02 gas uniformly.

Eight trees were labeled July 19, 1990 and September 5, 1990. A total
of 3.04 x 108 Bq of 14CO2 with a speciﬁc activity of 4215 Bq mg"1 C was added
to the labeling chamber in July. In September, a total of 2.85 x 108 BQ of

%

14C02 with a speciﬁc activity of 4300 Bq mg‘1 C was added. Each tree
received on average 1.0 mCi of 14C. Chamber C02 levels were compared to
ambient levels and maintained at 290-350 ppm. Night respiration was
monitored until the following morning to determine above-ground dark
respiration. The following morning C02 from night respiration was
allowed to reassimilate before the Plexiglas chamber was removed.

Two weeks after the addition of 14C, 3 randomly-selected trees were
sampled. The leaves, stem and branches were composited according to the
following designations: ﬁrst year stem, bottom two-thirds (including stem,
branches and leaves); ﬁrst year stem, top one-third (including stem,
branches and leaves); second year branches (including branches and
leaves); and, second year leader (including stem and leaves). The soil block
was excavated by depth (0-25 cm, 25-60 cm and 60-100 cm) into wheel
barrows. The soil from each wheel barrow was weighed and sieved
through a 8 mm sieve to remove coarse roots. The soil was then placed into
a cement mixer, thoroughly mixed and a subsample taken. The sampled
plant material and soils were stored on ice in the ﬁeld and then transferred
to a 4°C cold room.

In the laboratory, roots were sorted by hand into the following size
classes; <05 m, 0.5-1 mm, 1-3 mm, >3 mm, >10 mm. Leaf area was
measured using a Delta-T meter (Decagon Devices, Pullman, WA). Plant
material was dried at 80°C for 48- 96 hr, coarse roots and stems were dried
for longer periods. The unrecovered ﬁne roots (<0.5 to 1 mm) from the ﬁeld-
sieved soil sample composites were recovered by hydronuematic elutriation
(Smucker et al., 1982) to determine the total ﬁne root biomass for each soil
block.

Plant tissue was ground in a Wiley mill to pass a 60 mesh screen.

37

Plant and soil samples were analyzed for C, N and 15N using a gas
chromatography-mass spectrometer (AN CA-MS, Europa Scientiﬁc, Crewe,
U. K.). Ammonium sulfate (0.3663 atom % 15N) and forty-four percent C
(as cellulose) Whatman no. 1 ﬁlter paper were used as a reference
standards. Radiolabeled C from the gas chromatograph-mass
spectrometer exhaust was collected in a C02 absorbing cocktail (J. R.
Harvey Instrument Corp., Hillsdale, NJ) and analyzed for 14C in a Liquid
Scintillation Analyzer according to the procedure of Harris and Paul (1989).

Expression of Results

The 14C results are expressed as speciﬁc activity (Bq mg‘1 C). The
15N results are expressed as atom percent 15N excess above natural 15N
level and N derived from 15N (Ndf15N). The N derived from 15N is the ratio
of atom percent excess of tissue/atom percent excess applied. Standard
error of the mean is shown to demonstrate the usefulness of our labeling

techniques in the ﬁeld.

RESULTS AND DISCUSSION

Tree and Environmental Variables

The Eugenei clones were growing rapidly in height at a rate of 35
mm d'1 for the 3 week period prior to the July labeling. By September,
height growth had ceased as a result of budset. Trees labeled during July
had an average diameter of 4.4 cm (at 10 cm) and a height of 310 cm. Trees
labeled in September were 4.5 cm in diameter and 337 cm in height. The
total leaf area of trees was 6.1 m2 in July and 3.1 m2 in September. A leaf

area decrease in the autumn is not uncommon for this genotype. It is

38

attributable to late season drought stress and Melampsora rust and
Marsonnina leaf spot disease which cause loss of leaves in the lower
portion the canopy. A reduction in leaf biomass and increase in root mass
was characteristic of trees harvested in September (Table 3.1). The increase
in coarse root mass (> 3 mm) is associated with late season translocation
and storage of photosynthate (Pregitzer et al., 1990; Nguyen et al., 1990). As
a result of translocation and partial leaf loss, the root to shoot ratio
increased ﬁ'om 0.34 to 0.62 from July to September.

The mammum light intensity inside the chamber during the labeling
periods was 1250 umol m'2 s'1 in July and 1060 umol m'2 s‘1 in September
(Table 3.2). Net photosynthesis averaged over a photoperiod and tree canopy
was 3.8 umol C02 111'2 s‘1 in July and 6.2 umol CO2 m"2 s’1 in September.
Incident radiation levels inside the labeling chamber were typical of solar
days and reached saturating light levels during midday. Saturating light
levels of 800-1000 umol m'2 s'1 are typical for ﬁeld grown hybrid poplars
(Ceulemans et al., 1980). Under saturating light conditions, net
photosynthesis values of 95-207 umol CO2 m'2 s’1 have been reported for
hybrid poplars (Ceulemans et al., 1980; Nelson et al., 1982; Bassman and
Zwier, 1991). The fact that the average rate of photosynthesis was
somewhat lower than other values reported in the literature is probably
attributable to whole-tree measurements compared with the single leaf or
branch determinations of other studies. During autumn, an increases in
photosynthesis has been observed in mature leaves of hybrid poplars as
determined by 14C02 uptake (Nelson and Isebrands, 1983; Friend et al.,
1991), and I observed this phenomena as well (Table 3.2). Since signiﬁcant
leaf drop had occurred by September, the autumn canopies were composed

of a relatively greater proportion of sun leaves (leaf drop is acropetal in

{9

Table 3.1. The dry weight of tree components in July and September.
Standard error of the mean shown in parentheses.

 

 

 

 

weight % weight weight % weight
_(g) distribution (g) distribution
stem 781 (122) 33 770 (125) 30
branches 432 (96) 18 519 (28) 2)
leaves 528 (52) 23 303 (14) 12
total above- 1742 (270) 74 1592 (158) 62
ground
roots
<5 mm 185 (15) 8 213 (4) 8
.5-1 mm 16 (2) 1 24 (2) 1
1-3 mm 49 (2) 2 53 (3) 2
3-10mm 136 (36) 6 230 (32) 9
>10 mm 116 (38) 5 273 (53) 11
cutting % (18) 4 191 (10) 7
total below- 599 (104) % 984 (25) 38
ground
total 2341 (370) 2576 (182)

 

Table 3.2. Gas exchange and environmental characteristics during
labeling periods in July and September.

 

 

 

shill Sentemlm
Photon Flux Density (max.) umol m-2 s-1 1250 1060
Net Assimilation umol C02 111'2 8'1 3.8 6.2
Above-ground Dark Respiration 0.67 1.47
umol CO2 ln'2 8'1
Dark Respiration (% of 14C added) 18 24
Chamber air °C (min-max.) 19-33 19-34
Ambient air °C (min.-max.) 19-36 18-34

Soil surface °C (min.-max.) 18-24 18-24

 

40

Populus). This may have inﬂuenced average rates of photosynthesis.

Dark respiration measurements included leaves, branches and
stem. Dark respiration expressed on the basis of leaf area was 0.67 umol
CO2 m'2 s'1 in July and 1.47 umol CO2 m'2 S'1 September, or 18 to 24% of
net assimilation (Table 3.2). Dark respiration measurements for hybrid
poplars have been estimated at 1.1-1.6 umol CO2 m"2 8'1 on portions of
intact leaves (Bassman and Zwier, 1991). The data demonstrate that night
respiration during warm periods can be signiﬁcant and should be
addressed when calculating the 0 balance of ﬁeld grown poplars.

Chamber air temperature was maintained between 19 to 34°C and it
closely tracked ambient air temperature ﬂuctuations of 18 to 36°C (Table
3.2). Surface soil temperature in the below-ground headspace ranged from
20 to 24°C for both labelings. To avoid water condensation and unnaturally
dry air inside the chamber, the temperature of the heat exchanger was

maintained at approximately the dew point of ambient air.

Root-Soil Respiration

Below-ground respiration measurements were recorded for each tree
for two weeks following the labeling period (Figure 3.2). The soil surface
respiration rate per tree averaged 240 mg CO2-C 111'2 hr“1 in July and 218
mg 002-0 111'2 hr'1 in September. On average, between 5.2 and 5.8 g of
C02-C m‘2 d'1 were respired from the soil surface of each soil block (1 m3 of
soil). This measurement includes both root, associated rhizosphere
organisms and soil microbial biomass respiration.

The speciﬁc activity (Bq mg'1 C02-C) traces of the below-ground
respiration indicate that a period of approximately 2 weeks was required to

metabolize the translocated 1‘iC-photosynthate in the root-soil system

41

Figure 3.2. Root-soil respiration averaged over eight trees for the period
following labeling in July and September. Values are expressed as mg

CO2-C m‘2 hr'l. Standard error of the mean depicted as line bars.

Figure 3.3. Average speciﬁc activity of below-ground respiration. Each
symbol represents the mean of eight root-soil microcosms.

 

 

Figure3.2
750
PIE an i 0 July
as I September
8 45° (13¢
C?
N ax) _
s ++¢ fungi , t i,
M 150 — + Hlﬁo "D
s <1> _ on,
o e
0 100 200 300 400
time(hours)
Figure3.3.

July Label

 

time (hours)

0 September Label

 

 

time (hours)

43

(Figure 3.3). During both labeling periods, maximum below-ground
speciﬁc activity (90-100 Bq mg'1 CO2-C) occurred approximately 2 days after
labeling. This represents the time needed to translocate radiolabeled
photosynthate for growth and maintenance of the root system. The
secondary rise occurring in the September speciﬁc activity curve may be
attributable to mycorrhizal symbionts sequestering remobilized starch
reserves, microbial utilization of root exudates or root decomposition
(Harris and Paul, 1992). When the speciﬁc activity trace dropped to about
5% of its maximum I assumed that the translocated 14C was no longer
being metabolized and was now in either storage or structural forms
(Harris and Paul, 1992). At this time, 14 days after labeling, the tree-soil
system was harvested.

The 14C recovered from below-ground respiration expressed as a
percent of that applied (Bq/Bq added) is an index of the quantity of
assimilated 14C lost to root respiration. The amount of respired 14C from
roots and associated soil microorganisms was 8.1% in July and 9.8% in

September (Figure 3.4). The exponential equation:

39
B(lapplied

=Q(1—e"kt) (1)
where BQroot is the Bq in root-soil respiration, BQapplied is the total Bq
added during labeling, Q is the accumulation of Bq in the root-soil
respiration component, k is the rate constant for Bq accumulation and t is
time, closely simulated the below-ground speciﬁc activity trace. The
calculated accumulation of 14C in the respired root-soil C pool was 9.1% in
Jilly and 10.8% in September using values extrapolated to 1000 hr. The
Spedﬁc activity trace of the September trees required a longer period to

 

 

TP“
"'1‘—
din--

0.14
0.12

_ _— _ — — — 1
ﬂ
——

llilllll

lllll

 

   

 

 

 

3
S
as
a.
Q 0 1 W " -I
5' 0.08 _:_
--)—
g 0.06 0 July :1:
a 004 July estimated _:_
'3 0 September {-
S 0-02 — - - September estimated '2’
U 0 ,. L l l l 1*—
I l l T
0 2X) 4(1) HI) 8“) 1000

time (hours)

Figure 3.4. Carbon- 14 respired from the root-soil system.

45

reach an asymptote than in July. This indicates that it took longer for 14C
to stabilize in September. The turnover rate of the translocated 14C in the
roots can be calculated from the rate constant for Bq accumulation in
equation 1. The half-life of respired 14C in the below-ground system was 69
hours in July and 99 hours in September.

Root respiration, as measured, represented the fraction of 14C
sustaining the metabolic activities of both the root system and associated
soil organisms. In both the July and September labelings root-soil
respiration accounted for 9 to 12% of the recovered 14C. The majority of the
respired 14C was probably due to the metabolic activity of the root system
since the microbial biomass contained only a small amount of the 140 two
weeks following the labelings (Horwath, unpublished data). As with our
net photosynthesis measurements of the canopy, these values represent the
entire root system and must be taken in light of the 81 to 89% total recovery
of 14C. The assumption concerning incomplete recovery of 14C is that
experimental error was spread randomly through the labeling and
sampling protocols. If the majority of the loss occurred from the bottom of
the unsealed soil block or leaks in the below-ground containment system,
root respiration values could possibly double. However, it is more likely that
the unrecovered 14C was lost through above-ground respiration shortly
after the Plexiglas chamber was removed.

The separate contributions of root and soil biomass respiration to 002
ﬂux from soil is difficult to discriminate. In our study, the proportion of
14C respired from the roots was related to net assimilation of 14C during
the labeling period. Ifwe assume that all of the 14C respired over 500 hours
was due to root respiration (Figure 3.4), then the contribution of root

respiration to total soil respiration was 19% in July and 21% in September.

46

Cropper and Gholz (1991) estimated that two thirds of soil respiration was
from the ﬁne root ﬁaction in mature slash pine trees. Edwards and Harris
(1977) calculated that 35% of forest ﬂoor respiration in a deciduous forest
evolved from the metabolic activity of roots. Our values for root respiration
are lower and represent a direct determination of the portion of gross
assimilation used for the maintenance of the root system over a two week
period following labeling. Figure 3.3 demonstrates that little 14C was
utilized for root respiration and growth two-weeks following labeling, i. e. it
is at least temporarily stabilized in structural or nonstructural compounds.
This clearly shows that these trees primarily depend upon current

photosynthate for growth and maintenance during the growing season.

Carbon Partitioning

Whole-tree 14C distribution varied according to clonal phenology.
During active growth in July, 80% of the 14C was found in above-ground
components compared to 49% after budset in September (Table 3.3). The
14C recovered in July correlates strongly to the weight of the most actively
growing portions of the tree. In September, more 14C was allocated to roots
and recovery closely resembled the biomass distribution in the different root
size classes. The amount of recovered 140 in the below-ground components
increased 3.5 fold to 38% in September (Table 3.3). The ﬁne and coarse root
fractions become signiﬁcantly enriched with 14C in September as compared
to July. In July, the speciﬁc activity of the ﬁne roots was about 25-50% of
speciﬁc activity of the actively growing above-ground tissues. By September,
the root system became a more active sink for 14C with only the leaves
having a higher speciﬁc activity. The total recovery of 14C ranged from 81%

in September to 89% in July and is among the highest recoveries we have

47

Table 3.3. Carbon concentration (%), speciﬁc activity (Bq mg-l C), % of 14C
recovered of total applied (Bq/Bq added), and % 1‘er of the total amount of

14C recovered for the different tree components. Standard error of the
mean in parentheses.

Jul: 15% 1i 1990

 

 

 

 

 

%C Bq Bq/Bq % 11C of
mg-l C added total 14C
recovered
stem 43.0 (0.6) 35.6 (2.5) 31.3 (4.8) 34.7 (2.0)
branches 42.5 (0.5) 36.6 (4.8) 16.8 (1.9) 18.8 (0.5)
leaves 40.4 (0.3) 42.8 (0.7) 23.9 (1.7) 26.9 (0.8)
total above-ground 72.0 (8.3) 80.4 (1.8)
roots
<5 mm 28.1 (2.4) 12.9 (1.6) 1.7 (0.0) 2.0 (0.2)
.5-1 mm 36.1 (2.0) 13.3 (1.8) 0.2 (0.0) 0.2 (0.0)
1-3 mm 38.2 (1.7) 15.9 (3.9) 0.8 (0.1) 0.9 (0.2)
3-10 mm 38.7 (1.0) 21.3 (5.5) 2.6 (0.3) 3.2 (0.5)
>10 mm 40.7 (0.0) 21.8 (5.4) 2.1 (0.2) 2.3 (0.2)
cutting 41.2 (1.2) 20.3 (3.9) 2.0 (0.1) 2.3 (0.1)
soil respiration 45.4 (1.8) 7.7 (0.2) 8.8 (0.9)
total below-ground 17.1 (0.2) 19.6 (1.8)
total tree 89.3 (8.1)
September5-19, 1990 _
stem 40.8 (1.0) 17.3 (1.9) 15.6 (3.7) 19.2 (4.5)
branches 42.0 (2.0) 12.4 (1.8) 7.5 (1.0) 9.3 (1.4)
leaves 39.5 (2.2) 50.8 (4.6) 17.0 (1.3) 20.9 (1.3)
total above-ground 40.1 (3.9) 49.4 (4.4)
roots
<5 mm 28.7 (1.3) 31.5 (6.3) 5.5 (1.3) 6.8 (1.8)
.5-1 mm 31.9 (2.0) 35.2 (4.4) 0.8 (0.1) 0.9 (0.2)
1-3 mm 34.0 (1.2) 32.0 (5.7) 1.6 (0.3) 2.0 (0.3)
3-10 mm 36.6 (0.5) 38.9 (7.3) 8.8 (1.0) 10.9 (1.4)
>10 mm 37.3 (1.2) 38.1 (5.7) 11.1 (3.2) 13.5 (3.5)
cutting 43.0 (0.6) 25.9 (9.2) 3.4 (0.7) 4.3 (1.0)
soil respiration 39.0 (3.5) 9.0 (1.2) 12.2 (1.8)
total below-ground 37.7 (4.2) 50.6 (4.4)

total tree

81.1 (2.2)

 

achieved in the ﬁeld.

The C allocation characteristics of this clone are closely related to the
termination of shoot growth (Michael et al., 1988). After budset, source-
sink relations change and roots became a major sink. Budset and leaf
senescence on branches occurs acropetally in this clone. The onset of
budset in both the terminal and branch shoots shifts the C allocation
pattern from acropetal to basipetal (Isebrands, 1982). Hybrid poplars gain
root sink strength as autumn approaches (Isebrands and Nelson, 1983;
Friend et al., 1991). Similar conclusions have been derived for apple trees
(Quinlan, 1969). During our labeling studies, translocation of radiolabeled
C to the root system nearly quadrupled from July to September.

The speciﬁc activity of labeled tissue proved useful in estimating sink
strength and growth rates. The leaves were signiﬁcant C sinks containing
21-27% of the recovered 14C in July and September. The speciﬁc activity of
the leaves remained higher than all other tissues regardless of the time of
labeling. Similar results for conifer species have been obtained using
different methods of pulse-labeling (Smith and Paul, 1989; Kuhns and
Gjerstad, 1991). A signiﬁcant amount of C exchange must occur in
metabolic pools associated with C ﬁxation and respiratory maintenance in
leaves. This C exchange occurs even after budset when shoot growth has

ceased.

Nitrogen Distribution

The above-ground components of the tree were a major sink for N in
both July and September. In July, the leaves contained 57%, stem and
branches 34% and roots 8% of the recovered 15N (Table 3.4). During
September, the leaves contained 16% of the injected N while stem and

49

Table 3.4. Nitrogen concentration (%), atom % 15N excess, nitrogen derivehr-«a

from applied 15N (%Ndf15N) and % 15N recovered of the total amount
applied for the various tree components. Standard error of the mean in

 

 

parentheses.
Jul: 15M1, 1990
%N Atom % %Ndf N % N
excess of total
_ _ ﬂiplied
stem 0.43 (0.03) 0.51 (0.08) 0.52 19.2 (0.7)
branches 0.59 (0.04) 0.53 (0.08) 0.54 15.3 (0.7)
leaves 2.41 (0.09) 0.46 (0.05) 0.47 57.3 (1.0)
total above-ground 91.7 (0.8)
roots
<5 mm 0.72 (0.06) 0.06 (0.00) 0.06 0.8 (0.0)
.5-1 mm 0.76 (0.06) 0.09 (0.01) 0.09 0.1 (0.0)
1-3 mm 0.73 (0.08) 0.17 (0.03) 0.17 0.6 (0.0)
3-10 mm 0.76 (0.06) 0.26 (0.04) 0.26 2.4 (0.3)
>10 mm 0.62 (0.05) 0.34 (0.07) 0.34 2.1 (0.6)
cutting 0.47 (0.02) 0.48 (0.06) 0.49 2.1 (0.1)
total below-ground 8.2 (0.9)
total tree 99.1 (2.2)
August 318eptember 19, 1990 J _
stem 0.48 (0.01) 0.70 (0.06) 0.71 26.4 (2.1)
branches 0.87 (0.04) 0.78 (0.12) 0.80 36.1 (2.6)
leaves 2.44 (0.03) 0.22 (0.04) 0.23 17.4 (3.6)
total above-ground 79.8 (2.0)
roots
<5 mm 0.71 (0.03) 0.12 (0.02) 0.12 1.9 (0.3)
.5-1 mm 0.82 (0.03) 0.15 (0.02) 0.15 0.3 (0.1)
1-3 mm 0.93 (0.04) 0.21 (0.02) 0.22 1.1 (0.1)
3-10 mm 1.07 (0.06) 0.24 (0.03) 0.25 6.2 (1.0)
>10 mm 0.90 (0.02) 0.27 (0.03) 0.27 6.9 (1.4)
cutting 0.63 (0.05) 0.31 (0.03) 0.31 3.8 (0.4)
total below-ground 20.1 (2.0)

total tree 95.4 (1.8)

50

branches were now the strongest sink for N, containing 60%. The woody
shoot of poplars have been shown to store proteins in ray cells during
dormancy (Sauter et al., 1988, 1989).

The root system contained twice as much 15N in September
compared to July, demonstrating the downward translocation of N typical
of this clone in autumn (Pregitzer et al., 1990). The majority of the
additional 15N found in the root system in September was located in the
coarse root fraction. Fine roots were also more enriched with injected 15N
late in the growing season.

The plant nitrogen derived from the labeled nitrogen (Ndf15N)
expressed as a percentage of atom percent excess of each tree component to
atom percent excess of injected 15N indicates the amount of labeled N in
tree tissue. The N df15N of all the tree components is less than 1% for both
labelings (Table 3.4). This shows that the injected N was a minor
component of the total N in all tissues. The injected 15N was meant only to
tag tree N pools. Recovery of labeled N was greater than 95% of that applied
(bath labelings).

The 15N labeling of poplar trees by injection has been shown to enrich
individual tree components relatively uniformly (Horwath et al., 1992). The
method is useful to determine N sink strength and to preferentially label
leaves and roots for long-term turnover studies. Cooper and Clarkson
(1989) have proposed that N utilization is regulated by a common pool of N.
This would explain the uniform labeling of tissues and preferential loading
of stronger N sinks.

The clonal response to senescence is to store N in woody tissue for
subsequent growth the following growing season (Pregitzer et al., 1990).

The September increase of 15N in coarse roots and branches illustrates this

51

point. The 15N distribution increased in the root pool along with total N.
However, the distribution is not equivalent in different root classes. Uptake
of soil N, or a diminishing sink for N in the ﬁne roots could explain the
preferential enrichment of the woody coarse roots. However, preferential
storage in coarse roots is a more likely answer. The above-ground N df15N
values for the July and September illustrate that the stem and branches are
also a strong sink for the injected 15N during the latter part of the growing
season.

The comparison of the interrelationships between N distribution and
C ﬂow during the growing season are made possible using the dual
labeling approach. Table 3.5 adapted ﬁ'om the 15N and 14C distribution in
Tables 3.3 and 3.4 shows that N and C do not move similarly. The high
recovery of 15N to 140 in leaves during July is not surprising. The ratio of
4.8 in branches in September and general retention of 15N in the above-
ground portion is noteworthy. The root system gains an additional 10% of
the injected 15N by September, but its distribution is masked by the
additional accumulation of 14C changing the ratio of the isotopes little. The
15N injection technique preferentially loads the aboveground tree
components and may require a longer equilibration period to distribute the
N throughout the entire tree. This is possible since the unlabeled N
distribution above-ground is 82% in July and 65% in September (of the total
tree N). The 15N injection technique demonstrates marked changes in N
distribution between the labeling dates and this may reﬂect the storage
pattern of tree components for N at speciﬁc times during the growing
season.

The pronounced seasonal pattern of C and N allocation in Populus

was ﬁrst elucidated by Dickson and Nelson (1982) and Isebrands and

Table 3.5. The interrelationships between the distribution of 15N and 14C of
the various tree components from the July and September labelings.

The values are expressed as the ratio of the distribution of 15N to 14C
recovered in the tree components.

 

Wm Jill! Sentember
stem 0.61 1.69
branches 0.91 4.80
leaves 2.41 1.02
total aboveground 1.27 1.99
roots
<5 mm 0.47 0.35
.5-1 mm 0.45 0.38
1-3 mm 0.75 0.69
3- 10 mm 0.92 0.70
>10 mm 1.00 0.62
cutting 1.05 1.12

total below-ground 0.87 0.70

53
Nelson (1983). Later, Pregitzer et al. (1990) and Nguyen et al. (1990)

demonstrated that the root system of Eugenei is a major site for the storage
of nonstructural carbohydrates during the dormant season and that N
allocation, like C allocation, exhibits a pronounced seasonal pattern of
distribution and storage that reﬂects relative sink strength. This whole-
tree dual tracer study conﬁrms these earlier studies in rich detail. It is
clear that this genotype allocates C and N to the different components of the
tree preferentially, depending on time of year. These inherent patterns of C
and N partitioning can have a major impact on ecosystem studies
attempting to understand the role of the root system in providing substrate
(C) for microbial metabolism and litter decomposition. This topic will be
discussed in subsequent communications, but it is clear that C and N ﬂux
to the roots varies signiﬁcantly over the course of the growing season
depending on relative sink strength.

My results also point to the importance of developing decent
estimates of tree respiration. Above-ground dark respiration accounted for
18-24% of net daily assimilation and root respiration accounted for 19-21% of
soil respiration. Total respiration estimates are useful in determining
gross production and the impact of forest ecosystems on global C budgets.
Finally, I believe this study demonstrates the power of using tracers to
study whole-tree C and N allocation. Clearly, the pronounced seasonal
patterns documented in this chapter need to be tested by working with other

genotypes and species.

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Nelson, N. D. and J. D. Isebrands. 1983. Late-season photosynthesis and
photosynthate distribution in an intensively-cultured Populus
nigraxLaurifolia clone. Photosynthetica 17:537-549.

Nguyen, P. V., D. I. Dickmann, K. S. Pregitzer and R. L. Hendrick. 1990.
Late-season changes in allocation of starch and sugar to shoots,
coarse roots and ﬁne roots in two hybrid poplar clones. Tree Physiol.
7:95-105.

Olsen, J. S. 1975. Productivity of forest ecosystems. In Productivity of world
ecosystems. National Academy of Sciences. Washington D. C.

Persson, H. 1979. Fine root production, mortality and decomposition in
forest ecosystems. Vegetaitio 41:101-109.

Pregitzer, K. S., D. I. Dickmann, R. Hendrick and P. V. Nguyen. 1990.
Whole-tree carbon and nitrogen partitioning in young hybrid poplars.
Tree Physiol. 7:79-93.

57

Quinlan, J. D. 1969. Mobilization of 14C in the spring following autumn
assimilation of 14002 by an apple rootstock. J. Hort. Sci. 44:107-110.

Sauter, J. J. B. V. Cleve, and K. Apel. 1988. Protein bodies in ray cells of
Populus x canadensis Moench robusta. Planta. 173:31-34.

Sauter, J. J ., B. V. Cleve, and S. Wellenkamp. 1989. Ultrastructural and
biochemical results on the localization and distribution of storage
proteins in a poplar tree and in twigs of other tree species.
Holzforschung. 43:1-6.

Schier, G. A. 1970. Seasonal pathway of 14C-photosynthate in red pine '
labeled in May, July and October. For. Sci. 16:1- 13.

Smith, J. L. and E. A. Paul. 1988. Use of an in situ labeling technique for

the determination of seasonal 1‘er distribution in ponderosa pine.
Plant and Soil 106:221-229.

Smucker, A. J. M., S. L. McBurney, and A. K. Srivastava. 1982.
Quantitative separation of roots from compacted soil proﬁles by the
hydropneumatic elutriation system. Agron. J. 74:500-503.

Tranquillini, W. 1963. Die C02-jahresbilanz und stoffproduktion der zirbe
(Pinus cembra). Mitt. Forstl. BundesVersuchsanstalt Mariabrunn.
60:535-546.

Warembourg, F. R. and E. A. Paul. 1973. The use of 14002 canopy

techniques for measuring carbon transfer through the plant-soil
system. Plant an Soil. 38:331-345.

Chapter 4

THE FATE OF 14C AND 15N IN HYBRID POPLAR PLANTATIONS.

ABSTRACT

The growth of plants is often described as a series of competing sinks
for carbon and nutrients. Carbon and nutrient requirements of ﬁne roots
are vaguely understood. The intent of this research was to determine the
role of ﬁne roots in the cycling of C and nutrients in a Populus-soil system.
Two-year-old hybrid poplars were radiolabeled with 14CO2 and stem
injected with 15N. The tree-soil system was sampled during the following
year. Labeled leaf litter was placed on unlabeled tree plots in a leaf litter
exchange designed to differentiate C and N ﬂux from leaf litter and root
turnover. The microbial biomass contained from 0.4-1.2% of the
photoassimilated C and no 15N two weeks after labeling in root labeled soil.
This amount declined steadily during the following year. Conversely, the
leaf litter labeled soils contained more 1‘er and 15N than root labeled soils
and maintained the isotopic C and N levels throughout the ﬁrst year.
During the initial years of plantation establishment, leaf litter plays a more
signiﬁcant role in supplying substrate for microbial biomass activity and

soil organic matter maintenance than does root turnover.

59
INTRODUCTION

The growth of plants is often generalized as a series of competing
sinks for C and nutrients (Wardlaw, 1989). Bloom et al. (1985) maintain
that the allocation of resources will favor an optimal balance between the
shoot and roots eliminating resource limitation. The plasticity of the shoot
to root ratio ultimately will lead to changes in nutrient cycling and soil
organic matter conservation by altering the amount of leaf and root litter
entering the soil. The practice of fertilizing forests to increase production is
becoming more common, especially in short-rotation intensively-cultured
systems (Miller, 1984). The prospect of altering nutrient cycling processes
exists by changing the quality and amount of litter in high production
plantation forests.

Carbon allocation and nutrient requirements of ﬁne root systems in
forests are vaguely understood (Gholz et al., 1986; Santantonio and Grace,
1987; Raich and Naddlehoffer, 1989; Santantonio, 1989; Fogel, 1990). Fine
roots generally account for only 5% of the standing biomass, but often
account for a major proportion of stand production (Santantonio, 1989). The
variability of ﬁne root production estimates is often attributed to
methodology and assumptions for production and mortality (Kurz and
Kimmins, 1987; Vogt et al., 1986; Singh et al., 1984). Fine root production
expressed as a proportion of total net primary production ranged from 3-6%
for fast growing plantations on fertile sites (Santantonio and Grace, 1987;
Pregitzer et al., 1990) to 5-73% for various conifer systems (Agren et al.,
1980; Keyes and Grier, 1981; 1980; Fogel, 1983; Vogt, 1991). Fine root

production and turnover may have a substantial inﬂuence on the amount of

a)

litter returned to soil and the rate of nutrient cycling (Wollum and Davey,
1975; Henderson and Harris, 1975; Fogel, 1990).

Carbon and N allocation between the shoot and root will be dependent
on species and site. Modern forest practice selects for species that
preferentially allocate to shoot production to increase harvestable yield
(Nambiar and Fife, 1991). The change in allocation does not affect
branches, coarse roots or leaves, rather, a tradeoff between stem and ﬁne
roots occurs (Linder and Axelsson, 1982; Axelsson and Axelsson, 1986;
Santantonio and Santantonio, 1987; Santantonio, 1989). This fundamental
change in C and N allocation to ﬁne roots, a potential litter pool, may alter
soil N processes by changing the diversity and amount of litter input. An
understanding of soil N processes is crucial since N often limits growth of
trees (Keeny, 1980; Nommik and Larsson, 1989). The nitrogen cycle is
generally tightly conserved in aggrading natural forest systems (Cole and
Rapp, 1981; Gholz, 1981). In plantations, the change in root litter input may
alter N cycling, especially if root decomposition is a continuous process
rather than temporal (Miller, 1984). Nitrogen availability has been shown
to alter shoot/root allocation in trees (Linder and Troeng, 1981; Axelsson
and Axelsson, 1986; Gower and Vitousek, 1989; Pregitzer et al., 1990).

Nutrient uptake in a forest system is at its maximum during the
establishment period (Miller, 1989). The rotation length of intensively-
cultured forests is sufﬁciently brief to perpetuate the cycle of maximum
nutrient uptake. The consequence of perpetuating this cycle is to maintain
high concentrations of nutrients within the tree. Nutrient uptake may
exceed requirements in young trees (N ambiar and Fife, 1991). In forests
where rotation lengths are longer, the net accumulation of nutrients

declines after canopy closure (Miller, 1984; Meier et al., 1985). At this time,

61

maximum wood production occurs through the retranslocation of nutrients
from older tissue. The importance of retranslocated nutrients in fast-
growing plantations from roots is not well characterized (Nambiar and
Fife, 1991). Horwath et al. (1992) found only small differences in nitrogen
concentration of dead and live roots in hybrid poplar. Nambiar (1987) found
no signiﬁcant evidence of nutrient retranslocation from roots of Pinus
radiata. Similarly, woody heathland plants demonstrated no root nutrient
retranslocation (Aerts, 1990). In contrast, Meier et al. (1985) suggested that
the internal redistribution of N and P from ﬁne roots of various age Abies
amabilis stands signiﬁcantly met the requirements for the production of
ﬁne roots. The high nutrient requirements of plantation forestry may alter
soil N processes through changes in rotation length, intensity of culture,
season of harvest and loss of nutrient rich biomass through harvest. It is
also important to realize that changes in shoot/root to maximize production
may alter litter input to soil. The change in litter replenishment can
signiﬁcantly alter processes conserving the labile or active N and soil
organic matter pools.

The change in litter input will alter the quality of plant residues, a
major source of energy and nutrients for heterotrophic microorganisms in
forest ecosystems. The subsistence of soil organisms on residues will alter
concurrent immobilization, mineralization and stabilization reactions that
maintain nutrient cycling and conserve soil organic matter (V oroney et al. ,
1989). Therefore, the understanding of ﬁne root production, mortality and
turnover is essential to understanding the affects of forest management
practices on soil quality and long term nutrient cycling.

In the present study, I investigated the role of root and leaf litter in

nutrient cycling and soil organic matter dynamics. The introduction of

62

dual tracers, 14C and 15N, into trees at different times in the growing
season encompassed different patterns of C and N allocation. The objective
was to determine the fate of C and N within the tree and compare the
potential nutrient return and soil organic matter production from the two
different litters. Root turnover and production were estimated using pool
dilution of labeled 140 roots with unlabeled C. Measurement of 14C and
15N ﬂux into soil and microbial biomass provided a means to determine the
turnover rate of the two litter types. The fate of C and N in the tree was also
determined. This report describes the long term fate (1-2 years) of dual
labeled trees and litter in an intensively-cultured short-rotation hybrid

poplar plantation.

MATERIALS AND METHODS

A short rotation-intensively cultured hybrid poplar plantation was
established on a 0.2 hectare plot at the Kellogg Biological Station, LTER site,
Michigan State University, in southwest Michigan. Cuttings of Populus
euramericana cv. Eugenei, 10 cm in length, were planted on a 2 by 1 m
spacing in the spring of 1989. The Kalamazoo soil series is a ﬁne loamy,
mixed, mesic, Typic Hapudalf. The site was prepared by moldbroad
plowing and disking a ﬁeld previously used for agricultural crops. During
the establishment year, two sets of 8 trees consisting of 4 trees in adjacent
rows were selected for uniformity of growth characteristics. An
undisturbed 1 m3 soil block was trenched around each tree to 1 m depth
(Horwath, Chapter 3). Plywood partitions were inserted into the trenches

and the vertical faces of the soil pedon were wrapped with vinyl sheeting

63

before backﬁlling the slots. The partition served both as a gas tight seal for

root respiration studies and for the quantitative recovery of tracers.

15N Injection

The two sets of trees were stem injected with 15N, one set July 13,
1990 and the other set September 1, 1990. Each tree received 100 mg of 98
Atom % enriched (15NH4)2SO4 in an artiﬁcial sap solution (Horwath,
Chapter 2). Leaves were collected daily from the time of labeling and until
the ﬁnish of leaf fall in autumn. Upon collection leaves were placed in a
60°C forced air drying oven. Leaves from each tree were pooled for

subsequent analysis.

140 Labeling

Trees were exposed to 14C02 for an entire photoperiod on July 19,
1990 (July labeling) and September 5, 1990 (September labeling). Ambient
levels of 14CO2 (290-350 ppm) and temperature were maintained during the
assimilation period (Horwath, Chapter 3). The level of C02 was monitored
by an Infra Red Gas Analyzer and controlled by a computer operated 14CO2
generator. In July, a total of 3.04 x 108 Bq for an average 3.79 x 107 Bq tree'1
and speciﬁc activity of 4215 Bq mg“1 CO2-C was assimilated in the labeling
chamber. During the September labeling, 2.85 x 108 Bq was added to the
chamber for an average 3.56 x 107 Bq tree"1 and speciﬁc activity of 4300 Bq
mg‘1 C02-C.

Sampling Procedure
Three trees of eight were randomly selected and destructively

sampled two weeks after each labeling. In November, an additional three

64

trees were sampled from the July (November 2, 1990) and the September
labelings (November 4, 1990). The remaining two trees from each set were
sampled one year later on July 25, 1991 and September 5, 1991. The
sampling included the entire tree located above and within the 1 m3 soil
block. The sampling of leaves, branches and stem was described previously
(Chapter 3). The soil block was excavated by depth (0-25 cm, 25-60 cm, 60-
100 cm) and sieved through an 8 mm screen to recover coarse roots.
Subsamples of soil from each depth were hydronuematically elutriated to
recover and estimate ﬁne root biomass. The washed roots were stored in
80% ethanol.

The washed roots were separated from organic residues by hand and
the length determined using a modiﬁed Newman's line intercept method
(Newman, 1966). The weight of the washed roots from each soil sample
was determined after drying at 80°C for 24 hours. The root length and the
oven dry weight of < 0.5 mm roots was used to develop a regression equation
that estimated the relationship between root length and weight [(weight of
washed roots < 0.5 mm, mg) = 0.984 (root length, cm) + 0.076, R2=0.97,
standard error of estimate = 6.5%)]. The equation was used to derive the
weight of roots (< 0.5 mm) from trees sampled 1 year after the dual
labelings. Roots taken from the ﬁeld sieved soil were hand sorted according
to size classes; < 0.5 mm, 0.5-1 mm, 1-3 mm, 3-10 mm, > 10 mm and

cutting.

Plant Tissue and Soil Analysis

Leaf area was determined using a video imaging system (Delta-T
meter, Decagon Devices, Pullman, WA). Plant material was dried at 80°C
for 48-96 hours. Plant tissue was ground in a Wiley mill to pass a 60 mesh

65

screen. Plant and soil samples were combusted in a biological sample
converter (Roboprep, Europa Scientiﬁc, Crewe, England) to yield N2 and
CO2. The gases were analyzed on a continuous-ﬂow dual isotope ratio
mass spectrometer (ANCA-MS Europa Scientiﬁc, Crewe, England) for N,
C, 15N. Reference standards consisted of natural abundance 0.3663 atom %
(15NH4)2SO4 and 44% C, as cellulose (Whatman no. 1 ﬁlter paper, Harris
and Paul, 1989). The CO2 from the combusted plant tissue was trapped in a
C02 absorbing scintillation cocktail (Harvey Instrument Co. Hillsdale,
N.J., USA) and the 14C determined on a liquid scintillation spectrometer
(Packard Instrument Co., Downers Grove, IL). Radiolabeled glucose
containing 120 Bq 5 ul'1 was pipetted onto the cellulose ﬁlter paper and
dried at 80° C to determine the efﬁciency of 1‘er02 trapping.

Soil 14C was determined using a dichromate digestion and trapping
of CO2. Soil (1 g, oven dry) was digested with 1g KzCr203 and 12 mL of
H2SO4: H3PO4 (3:2, v/v) in a modiﬁed 25 X 200 mm culture tube. A 15 x 45
mm vial containing 3 mL of C02 trapping scintillation cocktail was
inserted and rested on a restriction placed into the glass wall of the culture
tube (Smith et al., 1989). The tube was promptly capped (Poly seal cap,
Fisher Scientiﬁc, Fairlawn, N.J,) and placed on a digestion block at 120°C
for 2 hours. Following digestion, the tubes were allowed to stand overnight.
The vials were capped and enclosed in 10 x 22 mm glass scintillation vial

and the 14C activity determined on a liquid scintillation spectrometer.

Microbial Biomass Determinations
Soil microbial biomass was estimated following each sampling of all
plots. Composite soil samples from each tree plot were resieved through a 4

mm screen. Soil moisture was determined gravimetrically following

%

drying at 105°C for 24 hours. Initial inorganic N levels were determined on
2 M KCl (5:1, v/w) soil (25 g, wet weight) extracts. Inorganic N was
analyzed on an auto-ﬂow analyzer (Lachet Chemicals Co., Mequin, WI).
Soil (50 g) was fumigated with ethanol-free chloroform in an evacuated
desiccator for 18-24 hours (Horwath and Paul, 1992). After the removal of
the chloroform, the soil was brought to 55% of water holding capacity and
placed into 1 L Mason jars. A vial containing 5 mL of 0.6 M NaOH was
placed into the mason jar to consume respired C02. The jars were closed
and allowed to incubate for 10 days at 25°C. An unfumigated set of controls
was incubated under the same conditions. After 10 days, the soil was
extracted for inorganic N as described above. Half the contents of the CO2
trap were titrated to a phenolthalein endpoint (pH 7) to determine respired
CO 2. The remaining NaOH was combined with 10 mL of scintillation
cocktail (Safety solve, Research products Int. Corp., Mount Prospect, IL)
and 14C activity determined on a liquid scintillation counter.

Microbial biomass C was calculated according to Voroney and Paul
(1984) using the following equation,

Biomass C = Cf/ 0.41
where, Cf indicates the amount of C02 respired from the fumigated soil
and 0.41 is the efﬁciency of the microbial biomass C determination. The
amount of 14C within the microbial biomass was determined by dividing
the 14C released upon fumigation incubation by 0.41 after correction for
background. Microbial biomass N was calculated by subtracting the initial
soil NH4+ level from the fumigated N ﬂush and correcting for efficiency
using a variable efﬁciency factor (kn) (V oroney and Paul, 1984).
kn = (186* (Cf/ Nf) -0.879

and

67

Biomass N = (Nf— Ni) / kn
Nf is the amount of NH4+ mineralized in the fumigated soil, Ni initial soil
NH4+ level and kn is the efﬁciency of the microbial biomass N

determination.

Litter Exchange Experiment
Leaf litter from the two labelings was collected from each tree until
late autumn when all leaves had abscised. Litter from each tree was
analyzed separately to complete whole tree 14C and 15N budgets and then
composited by labeling. The litter was placed on plots that were trenched to
a 25 cm depth and lined with plywood and Visqueen plastic. Each of these
plots contained a tree planted at the same time as the labeled trees
simulating the same design as those used in the dual labeling procedures.
Unlabeled leaf litter was placed on plots with labeled trees to complete the
litter exchange experiment. The amount of litter collected varied from each
labeling due to differences in the collection period. The amount of leaf litter
placed on the plots receiving September dual labeled leaves was less than
plots with July labeled leaves. The September plots contained 268 g and
July plots contained 601 g of leaf litter. These values were averages of the
total amount of litter that fell from the different trees labeled in July and
September. The litter on each plot was covered with plastic netting (garden
netting, 1.8 cm mesh) at a height of approximately 2 cm by attaching the
netting to the 15 cm tall plywood plot lining.
At approximately 6 month intervals a 50 by 50 cm area was sampled
from each plot. Remaining litter was collected from the soil surface by
carefully removing petioles and lamella structures. The soil was excavated

in two depths, 0-10 cm and 10-25 cm. The soil was sieved through a 6 mm

68

sieve, homogenized in a cement mixer and a subsample taken. The litter
material was dried at 60°C for 24 hours, coarsely ground in a Waring
blender and ball milled (Harris and Paul, 1989). The litter, soil and
microbial biomass were analyzed for 14C and 15N as described previously.

Expression of Results

Data are expressed as means with standard errors of the mean on
replicate components of the tree-soil system. During the ﬁrst year of
labeling and harvest, each mean is composed of 3 replicates (n = 3). The
harvests done one year later were composed of 2 replicates (n = 2). The ﬁrst
two harvests of the litter exchange experiment consist of 3 replicates (n=3)
for September labeled leaves and 4 replicates (n=4) for July labeled leaves.
The last harvest consisted of 2 replicates (n=2) for each litter. The 14C
results are expressed as speciﬁc activity (SA, Bq mg‘1 C), labeled C (total Bq
found in a component/SA of added 14COZ) and % of 14C applied. The 15N
results are expressed as atom percent 15N excess (atom percent 15N of
component - natural abundance, 0.3663%), labeled N (atom percent excess x
% N x sample wt) and % of 15N applied. Standard errors of the mean are
shown to demonstrate the reproducibility of dual isotopic labeling in the
ﬁeld.

RESULTS

Tree Characteristics
The trees grew rapidly, doubling in height every growing season. By
the end of the third summer, the trees were approximately 5 m tall (Table
4.1). Leaf area determined on July harvested trees remained constant

Table 4.1. Height and leaf area measurements taken from samplings done
during the growing season from trees labeled July 19, 1990 and
September 5, 1990. Standard error of the mean shown in parentheses.

 

 

Year
Labeling 1 2 3
July labeled trees
Height (current 159.3 (10.7) 151.0 (5.2) 207.5 (27.5)
increment) (cm)
Total height (cm) 159.3 (10.7) 310.3 (11.9) 517.8 (30.0)
Leaf area NA 6.1(0.6) 6.0 (1.4)
(m'2 tree'l)
September labeled trees
Height (current 153.3 (5.7) 183.7 (32.8) 172.0 (28.0)
increment) (cm)
Total height (cm) 153.3 (5.7) 337.0 (33.3) 509.0 (43.5)
Leaf area NA 3.1(0.6) 1.0 (0.3)

(m‘2 tree'l)

70

through the second and third year at approximately 6 m2 tree'l. Leaf area
from September labeled trees was reduced from leaf diseases such as
Melampsora rust and Marsonnina leaf spot . The dry weight of trees dual
labeled in July, 1990 and sampled through July, 1991 increased two fold
(Table 4.2). The September, 1990 dual labeled trees had less leaf biomass
than July labeled trees as a result of leaf drop in both 1990 and 1991 (Table
4.3). Despite the leaf mass loss in the September dual labeled trees, these
trees were still useful for the study of tracer dynamics and budgets. The
trees assimilated 1‘er according to leaf area and tree biomass thus

achieving similar concentrations and turnover rates of tracers.

Distribution of C and N in the Tree-Soil System

Figure 4.1 shows the total C and N contained within the tree and soil
microcosm of July labeled trees sampled August 2, 1990 and November 2,
1990. The soil contained 6.9 kg of C. Tree C represents 11% and the soil 74%
of the total C in the tree-soil system. The remaining C in the system was
composed of leaf litter, microbial biomass and soluble soil C. The amount of
C in the microbial biomass was twice the amount found in ﬁne roots (< 0.5
mm). Leaf litter C was approximately twice microbial biomass C and four
times standing ﬁne root biomass (< 0.5 mm) C in November. Fine roots
contain less than 1% and leaf litter 3.4% of soil C. The root system nearly
doubles in C content from July to November.

The soil within the 1 m3 microcosm contained 608 g of N (Figure 4.1).
The tree comprises 3% of the total N in the system. Leaves contain more
than 50% of total tree N during the growing season. Fine roots comprised
5% of tree N during the growing season and dormancy. The N level in

woody shoots and coarse roots (> 0.5 mm) doubles from July to November.

71

Table 4.2. The dry weight (g) of tree components from July 19, 1990 dual
labeled trees sampled over the following year. Standard error of the
mean shown in parentheses.

 

 

— Statehouse
2-Ati-90 2-Nov-90 25-Jul-91
stem 781 (122) 1053 (106) 1468 (59)
branches 432 (96) 534 (77) 702 (168)
leaves 528 (52) 619 (22) 597 (190)
total 1742 (270) m6 (203) 2766 (81)
roots 0-25 cm
< 0.5 mm 88 (8) 70(8) 114(4)
0.5-1 mm 4(1) 10(2) 21 (1)
1-3 mm 11 (1) 23 (1) 16(3)
3-10 mm 53 (20) a) (3) a) (31)
> 10 mm 82 (42) 188 (32) 3% (192)
cutting % (18) 183 (11) 423 NA
total 336 (82) 554 (28) 1095 (225)
roots 25-60 cm
< 0.5 mm 51(4) 45 (5) 63 (4)
0.5-1 mm 7(2) 9(1) 2(1)
1-3 mm 18(2) 31 (5) 14(0)
3-10 mm 55 (21) SB (15) 76 (18)
> 10 mm $ (11) 114 (46) 104 NA
total 155 (34) 2% (24) KB (75)
roots 60-100 cm
< 0.5 mm 46 (11) 51(1) 48 (6)
0.5-1 mm 5(1) 9(1) 11 (0)
1-3 mm 2) (5) 35 (5) 22 (2)
3-10 mm $ (13) 81 (11) 59 (14)
> 10 mm 3 NA Q (7) 27 NA
total 108 (28) 205 (20) 154 (24)
Root total 599 (93) 1054 (42) 1456 (238)
Tree total 2341 (370) 3%0 (239) 4223 (45)

 

72

Table 4.3. The dry weight (g) of tree components from September 5, 1990
dual labeled trees sampled over the following year. Standard error of the
mean shown in parentheses.

 

 

20-Sep-90 4-Nov-90 5-Sep-91
stem 770 (125) 654 (56) 1650 (255)
branches 519 (28) 375 (67) 470 (229)
leaves 303 (14) %1 (10) 106 (37)
total 1592 (158) 1310 (122) 22% (522)
roots 0-25 cm
< 0.5 mm 107 (11) 49 (9) 74(2)
0.5-1 mm 10(2) 7(1) 8(2)
1-3 mm 17(3) 13(1) 12(2)
3-10 mm 65 (22) Q (4) 40 (29)
> 10 mm 163 (38) 77 (24) 1% NA
cutting 191 (10) 133 (18) 233 (53)
total 552 (16) 348 (32) 431 (151)
roots 25-60 cm
< 0.5 mm 55 (8) :5 (3) 48(5)
0.5-1 mm 6(0) 5 (0) 8(1)
1-3 mm 16(2) 15(3) 19 (7)
3-10 mm 82 (11) 99 (16) 72 (2)
> 10 mm % (19) 71 (6) 37 (17)
total 255 (7) 2% (14) 184 (17)
roots 60-100 cm
< 0.5 mm 51 (8) 49 (5) 40 (5)
0.5-1 mm 8 (0) 7(1) 7 (1)
1-3 mm m (3) w (2) 11 (1)
3-10 mm 84(6) 8) (13) 48 (26)
> 10 mm 14(5) 9(3) 4 NA
total 177 (13) 170 (10) 108 (33)
Root total 984 (21) 744 (36) 723 (156)
Tree total 2576 (182) $54 (168) 2949 (723)

 

Leaves leaves
213 . 12.8

  

 

 

 

Branches Branches l
184 2. 5
(208) (4.5)

 

 

 

 

 

 

 

 

 

   

 

 

 

 

 

 

 

 

 

Total l :12“ I ,_. .
Fine roots roots ., :f . " a l'.'- :33: Flne roots
6 I ' - :: < 0.5 m
< 022711111 218 E29"; ' .ﬁ ‘1. . . 4.1 1 3
(359) - ‘1 8.4 . -
(38) 2. '. .‘ ( l i (0.8)
i Microbial ‘ {5.4.1: ii; :0 .'.' .. u, .: ."-‘: '.,-::i:-‘:l"v Microbial
biomass C _-':-' 2.32..- - .',' '21. '. H‘ _;"' "9 ._ biomass N
103 .51.}: .2. ,- .. .1?" 19.4

Soluble 1 Total Total

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

soil norganlc ‘
76

p I 3.3

 

 

 

 

 

 

 

Figure 4.1. The distribution of C and N (g) in the Populus-soil system for
July 19, 1990 labeled trees sampled August 2, 1990. The values in

parentheses represent the amount of C and N for the November 2, 1990
sampling.

74

The apparent retranslocation of N from senescing leaves represents a third
of total leaf N or 5.1 g on a mass basis. The level of microbial biomass N
was equivalent to total tree N in the second year of growth. This indicates
the importance of the microbial biomass as a source-sink for N and other
nutrients in this system. Inorganic N (NO3' and NH4") represents 15% of
both tree and microbial N. Complete tree C and N concentrations for all

labelings can be found in Appendix A (Tables A.1-A.4).

Distribution of 14C and 15N in the Tree

During mid-growing season, the shoot was actively growing and
contained greater than 70% of the tree biomass and 14C (Figure 4.2).
Leaves retained greater than 50% of the injected 15N 19 days following the
dual labeling procedure. Roots contained approximately 10% of the 15N and
14C during July. The majority of the recovered 14C was in the stem. The
distribution of 14C closely resembles the weight distribution of the tree in
July. The distribution of 14C represents the allocation of net primary
production. The shoot was the strongest sink for C and N during mid-
growing season.

The September dual labeling illustrates the basipetal movement of
both C and N during autumn (Figure 4.2). The shoot received 80% and
roots 20% of the injected 15N. The major N sink in autumn was the woody
shoot, especially branches. The root system contained approximately 45%
of the assimilated 14C 14 days after labeling. The root system received the
majority of net primary production in September. The shift in C and N
allocation from July to September exempliﬁes the phenological growth
pattern of this hybrid poplar genotype.

 

 

 

 

 

 

 

 

   

 

 

 

 

 

A. Julyhbel
m __ El leaves stem 8 roots > 1 mm
: branches [:1 roots < 1 mm

i

*3
s s
'5
g 60
g \ \ \ \ \ \ \ \
“E 40

.n

m —0—
a? :\ V
0 F k l I
weight
B. Sqrtelnberlabel
1% __ leaves E] stem El roots > 1 mm
i branches [:1 roots < 1 mm

E 10° : ;.;.;.;.;.;.;.;.;.

.1:

f"-

E

c:

O

‘3

:9

g V
‘8

A

 

 

 

 

 

 

  

14C

 

weight

Figure 4.2. Relative distribution of weight (g), 15N (mg) and 140 (mg)
within trees labeled on July 19, 1990 (A) and September 5, 1990 (B).

76

Speciﬁc Activity of 14C

Tables 4.4 and 4.5 present the speciﬁc activity, Bq mg“1 C, of the
different components of the tree-soil system. In July, the speciﬁc activity of
the shoot components were twice that of the root system. The increased 14C
activity demonstrates the sink strength for C of the shoot during mid-
growing season. The speciﬁc activity of roots was dependent on root
diameter. Small diameter roots contained less 14C than coarse roots or the
cutting during mid-growing season. The speciﬁc activity of roots (< 3 mm)
declines with increasing soil depth. The decline indicated lower sink
strength for C with increasing soil depth. Coarse roots (> 3 mm) maintain
similar speciﬁc activities at all soil depths. Microbial biomass C speciﬁc
activity was low compared to tree components. The assay for microbial
biomass encompassed the entire microbial population diluting any
rhizosphere interactions.

The November sampling of the July labeled trees shows dilution of
speciﬁc activity in all tree components and no change in the microbial
biomass. The 14C dilution in the woody shoot did not occur entirely from an
increase in unlabeled biomass. The dilution of 14C was also a result of
retranslocation and respiration of the assimilated labeled C. The dilution of
14C activity in roots (> 3 m) can be accounted for through the addition of
unlabeled biomass. The 14C dilution of roots (< 3 mm) was dependent on
soil depth. The constant biomass of these ﬁne roots from July to November
indicates either turnover of biomass and or, labile C.

One year following the July labeling, the speciﬁc activity of the shoot
and roots (3-10 mm) continued to decline. This dilution did not occur from
the addition of unlabeled biomass. This illustrates that C was still being

retranslocated or used in maintenance respiration one year after

Table 4.4. The speciﬁc activity (Bq mg'1 C) for the different tree components

sampled during the year following 14C labeling on July 19, 1990.
Standard error of the mean shown in parentheses.

 

 

 

Samulinulatc
2-Aui90 2-Nov-90 25-Jul-91
stem 35.6 (2.5) 23.4 (3.3) 12.3 (0.7)
branches 36.6 (4.8) 21.0 (3.5) 10.0 (0.3)
leaves 42.8 (0.7) 25.9 (1.7) 0.3 (0.1)
root 0-25 cm
< 0.5 mm 16.3 (1.7) 9.5 (1.2) 9.8 (2.7)
.5—1 mm 18.7 (2.0) 9.1 (1.8) 12.2 (2.7)
1-3 mm 18.8 (3.0) 7.9 (1.5) 6.7 (2.5)
3-10 mm 22.0 (5.6) 8.6 (0.2) 5.7 (1.4)
> 10 mm 22.7 (6.2) 11.1 (0.7) 5.6 (0.1)
cutting 20.3 (3.9) 10.7 (0.5) 9.2 NA
root 25-60 cm
< 0.5 mm 11.6 (1.6) 7.6 (0.6) 10.5 (0.7)
.5-1 mm 13.0 (2.8) 8.2 (0.4) 3.7 (2.5)
1-3 mm 15.9 (4.5) 8.0 (0.7) 9.1(0.9)
3-10 mm 19.7 (5.8) 6.3 (0.6) 6.0 (1.4)
> 10 mm 21.3 (8.8) 7.3 (0.4) 4.8 NA
root 60—100 cm
< 0.5 mm 7.9 (2.6) 6.7 (1.7) 6.9 (1.3)
.5-1 mm 8.8 (3.0) 7.2 (1.0) 8.2 (0.9)
1-3 mm 14.5 (3.7) 6.5 (1.2) 7.2 (0.5)
3-10 mm 21.4 (5.2) 6.7 (0.9) 5.1 (0.1)
> 10 mm 11.5 N A 7.5 (0.9) 4.0 NA
Emma] Biomass C 1.3 (0.2) 1.4 (0.2) 1.0 (0.2)

 

 

Table 4.5. The speciﬁc activity (Bq mg1 C) for the different tree components

sampled during the year following 14C labeling on September 5, 1990.
Standard error of the mean shown in parentheses.

 

 

SamplinLdate
20-Sep-90 4-Nov-90 5-Sep-91
stem 17.3 (1.9) 13.6 (1.5) 4.8 (0.5)
branches 12.4 (1.8) 10.9 (0.5) 4.2 (0.4)
leaves 50.8 (4.6) 49.3 (3.5) 0.5 (0.2)
root 0-25 cm
< 0.5 mm 35.7 (7.7) 27.4 (2.1) 21.2 (0.2)
.5-1 mm 38.7 (6.0) 36.1(1.8) 27.7 (3.8)
1-3 mm 37.8 (12.0) 32.2 (5.6) 24.8 (1.8)
3-10 mm 41.6 (8.4) 30.5 (1.6) 19.6 (5.0)
> 10 mm 35.1 (5.5) 19.0 (1.9) 9.0 NA
cutting 14.7 (2.8) 6.8 (1.7) 8.6 (2.4)
root 25-60 cm
< 0.5 mm 33.3 (3.5) 32.4 (4.3) 29.9 (3.8)
.5-1 mm 40.7 (2.9) 38.4 (6.0) 33.8 (4.9)
1-3 mm 38.7 (2.3) 36.3 (1.4) 29.8 (3.1)
3- 10 mm 34.3 (7.6) 29.3 (4.7) 27.4 (6.6)
> 10 mm 41.1 (6.1) 24.5 (5.7) 19.0 (2.9)
root 60-100 cm
< 0.5 mm 16.1 (3.2) 18.7 (2.0) 30.0 (3.7)
.5-1 mm 25.7 (6.6) 22.0 (1.2) 26.3 (0.9)
1-3 mm 20.8 (3.6) 22.3 (5.1) 18.4 (3.9)
3-10 mm 42.2 (8.2) 34.7 (3.7) 11.5 (3.5)
> 10 mm 59.3 NA 36.2 (4.4) 15.5 NA
ILIicrobial Biomass C 4.5 (0.5) 2.6 (0.2) 1.1(0.1)

 

79

assimilation. The 14C dilution in roots > 10 m can be accounted for
through an increase in biomass. Some shoot 14C may have also been
translocated to the root system. Roots (< 3 mm) maintain the same levels
speciﬁc activity from dormancy in November to one year following the
labeling. The constant ﬁne root 14C activity indicated minimal pool
turnover or synthesis of new roots from labeled reserves.

In the September labeling, the speciﬁc activity of roots exceeded that
of the woody shoot (Table 4.5). The cutting behaved similarly to the woody
shoot in 14C activity. Roots in the 0-60 cm soil proﬁle have become
uniformly labeled with 14C regardless of diameter. The microbial biomass
became signiﬁcantly labeled, containing 3.5 times more 14C activity than in
the July labeling. The increase in all below-ground components of the tree-
soil system was typical of this clone and exempliﬁes the basipetal
movement of C during autumn, some of which moves directly into
microbial biomass.

The dilution of 140 activity in shoot tissue was small from September
to November compared to the July labeling 14C dynamics. The speciﬁc
activity of leaves and leaf litter remain unchanged. The lack of 14C dilution
in most of the shoot-root system illustrates that C storage was ending just
after budset. The dilution of 14C still occurred in coarse roots (> 3 mm) and
ﬁne roots (< 0.5 mm) in the 0-25 cm soil proﬁle. The dilution of the 14C
activity in ﬁne roots was similar to results of the July labeling and may
indicate ﬁne root turnover during the latter part of the growing season and
the beginning of dormancy.

One year following the September, 1990 labeling, dilution of 14C had
occurred in the woody shoot and coarse roots. Leaves regain only a minor

portion of the previous years assimilated 14C. Little dilution of the 14C

8)

activity of ﬁne roots (< 3 mm) occurs. This difference between July labeled
and September labeled trees show the potential importance of late season
reserves in the production of ﬁne roots. The 14C activity of the microbial
biomass probably declines from the lack of root C input. The dilution of 14C
activity in coarse roots and lack of dilution in ﬁne roots may indicate
movement of reserve C to sustain the ﬁne root system during the next
growing season. Both the July and September labeling exemplify that ﬁne
root turnover was probably occurring during late autumn and early

dormancy immediately after leaf fall.

Distribution of 15N Activity

The largest atom % 15N excess was found in the shoot and coarse
roots (> 10 mm) from the stem injection of 15N in July, 1990 (Table 4.6).
Leaves and coarse roots contained similar concentrations of 15N. The atom
% 15N excess of roots (< 10 mm) was dependent upon root diameter
corroborating data obtained with 14C. Small diameter roots contained the
lowest level of 15N, typically ranging from 10 and 20% of the 15N excess
found in large roots (> 10 mm). The concentration of 15N within root size
classes was not affected by depth of the soil proﬁle as it was with 14C
activity. This indicates that C and N may be utilized via different
mechanisms since the sink strength of the two tracers was different within
the same root size classiﬁcation.

Subsequent sampling of the July, 1990 15N injection revealed a
decrease in the atom % in shoot and coarse roots (> 3 mm). Leaves of the
subsequent growing season contained signiﬁcant 15N, having an atom %
excess of 0.1%. The decline in atom percent of the stem, branches and

coarse roots was probably associated with uptake of unlabeled soil N and

81

Table 4.6. Atom percent 15N excess (atom % - natural abundance) is shown

for the different tree components sampled the year following 15N
injection on July 14, 1990. Standard error of the mean shown in

parentheses.

 

 

2-Aug-9o 2-Nov-90 25-Jul-91
stem 0.58 (0.07) 0.30 (0.04) 0.18 (0.00)
branches 0.63 (0.08) 0.35 (0.04) 0.13 (0.01)
leaves 0.46 (0.05) 0.37 (0.01) 0.10 (0.02)
roots 0-25 cm
< 0.5 mm 0.06 (0.01) 0.09 (0.02) 0.09 (0.00)
.5-1 mm 0.11 (0.01) 0.11 (0.02) 0.11 (0.01)
1-3 mm 0.18 (0.03) 0.16 (0.04) 0.20 (0.07)
3-10 mm 0.26 (0.03) 0.20 (0.03) 0.14 (0.03)
> 10 mm 0.31(0.04) 0.21(0.03) 0.14 (0.02)
cutting 0.48 (0.06) 0.21 (0.01) 0.16 NA
roots 25-60 cm
< 0.5 mm 0.07 (0.01) 0.09 (0.02) 0.08 (0.00)
.5-1 mm 0.10 (0.01) 0.12 (0.02) 0.09 (0.03)
1-3 mm 0.17 (0.04) 0.16 (0.03) 0.12 (0.01)
3-10 mm 0.25 (0.06) 0.18 (0.04) 0.13 (0.01)
> 10 mm 0.40 (0.12) 0.20 (0.04) 0.13 NA
roots 60-100 cm
< 0.5 mm 0.05 (0.02) 0.07 (0.02) 0.08 (0.03)
.5-1 mm 0.07 (0.02) 0.11 (0.02) 0.11 (0.03)
1-3 mm 0.16 (0.03) 0.14 (0.02) 0.12 (0.02)
3-10 mm 0.23 (0.01) 0.16 (0.03) 0.11 (0.01)
> 10 mm 0.43 NA 0.14 (0.00) 0.11 NA

 

Table 4.7. Atom percent 15N excess (atom % - natural abundance) for the

different tree components sampled during the year following 15N
injection on August 31, 1990. Standard error of the mean shown in

 

 

parentheses.
SamplinLdate
20-Sep-90 4-Nov-90 5-Sep-91
stem 0.70 (0.06) 0.68 (0.06) 0.26 (0.01)
branches 0.78 (0.12) 0.91 (0.07) 0.29 (0.08)
leaves 0.22 (0.04) 0.22 (0.04) 0.21 (0.05)
roots 0-25 cm
< 0.5 mm 0.12 (0.02) 0.09 (0.01) 0.12 (0.03)
.5-1 mm 0.16 (0.01) 0.14 (0.01) 0.16 (0.05)
1-3 mm 0.26 (0.03) 0.21(0.03) 0.21(0.07)
3-10 mm 0.30 (0.03) 0.27 (0.03) 0.28 (0.09)
> 10 mm 0.29 (0.02) 0.29 (0.02) 0.20 NA
cutting 0.31 (0.03) 0.26 (0.05) 0.25 (0.08)
roots 25-60 cm
< 0.5 mm 0.13 (0.01) 0.12 (0.01) 0.15 (0.04)
.5-1 mm 0.17 (0.01) 0.17 (0.03) 0.20 (0.05)
1-3 mm 0.23 (0.00) 0.22 (0.03) 0.24 (0.07)
3-10 mm 0.22 (0.02) 0.25 (0.04) 0.31(0.11)
> 10 mm 0.24 (0.03) 0.25 (0.02) 0.31(0.11)
roots 60-100 cm
< 0.5 mm 0.08 (0.02) 0.09 (0.01) 0.14 (0.03)
.5-1 mm 0.13 (0.02) 0.12 (0.01) 0.15 (0.03)
1-3 mm 0.16 (0.02) 0.18 (0.03) 0.16 (0.01)
3-10 mm 0.23 (0.05) 0.25 (0.02) 0.14 (0.01)
> 10 mm 0.26 (0.05) 0.21(0.03) 0.18 NA

 

83

retranslocation of 15N to new leaves. The decrease of atom % 15N excess in
the woody shoot and coarse roots may have also maintained ﬁne roots. The
atom % 15N excess of roots (< 3 mm) remained constant into the next
growing season. The constant level of 15N in ﬁne roots suggested that
turnover of this biomass pool was minimal or that stored 15N was used to
construct new roots. These observations were in agreement with the 14C
results, mentioned earlier and implies the two tracers were behaving
similarly over time.

In the September N injection, the largest atom % 15N excess
occurred in the woody shoot with branches containing 0.78% and stem
0.65% (Table 4.7). The remaining tissues of the tree contain 0.31% or less
atom % 15N excess. The atom % excess. of roots was again dependent on
diameter as it was in the July 15N injection. Roots (> 1 mm) in the 0-25 cm
soil proﬁle contained slightly more atom % 15N than roots at lower soil
depths. The subsequent November sampling of the September injected trees
showed slight dilution of labeled N throughout the tree. An increase of
atom % 15N excess occurred only in branches from 0.78 to 0.91% and was
probably a result of the retranslocation of N from leaves before senescence.

No signiﬁcant changes in the atom % excess of roots had occurred by
September, 1991, one year following the 15N injection. The shoot tissue 15N
enrichment decreased dramatically during the third growing season. New
leaves acquired 15N from stored N. The constant 15N enrichment of roots
corroborated with the July, 1990 15N injection results (Table 4.6). These
results combined with the 14C activity data (Tables 4.4 and 4.5) suggest that
either minimal turnover or reliance on labeled reserves maintained the
ﬁne root system from dormancy to the next growing season. Nitrogen

taken from the soil to support the third season leaf growth does not interact

84

with the existing N within the root system. The contrasting N results may
provide evidence for the persistence of ﬁne roots from the second to third

growing season.

The Fate of Labeled C and N

The recovery of 14C and 15N from the July and September labelings
declined during the following year (Table 4.8 and 4.9). The loss of the
applied 14C one year after labeling was 63% in July and 73% in September
labeled trees. Similarly, the loss of injected 15N was 28.2% in July and 45%
in September labeled trees. A large portion of both tracers was lost with the
fall of leaves during autumn. The majority of the 14C loss from the trees,
excluding loss from leaf and root turnover, was associated with respiratory
activity. Stem, branches and roots (> 0.5 mm) from both labelings lost a
signiﬁcant amount of 14C over the year. Fine roots (< 0.5 mm) from both
sets of labeled trees lost about 50% of their 1‘er from the time of labeling to
the November sampling indicating rapid turnover of 14C. A small increase
of 14C in ﬁne roots one year following both labelings indicates that new
growth of ﬁne roots was dependent on reserve C. Little 1‘er was found in
new leaves, less than 0.2% of the applied, indicating new leaf growth does
not rely heavily on stored reserves or that the reserves were mainly used to
initiate the formation of new leaves.

The fate of 15N in the shoot and roots of these hybrid poplars was
similar to 14C (Tables 4.10-4.11). Major movement of the injected 15N to
other areas occurred from leaf litter, woody shoot and coarse roots
amounting to over 50% of the applied for both labelings. New leaf growth
the year following injection contained 10.3% in July labeled trees and 4.2%

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89

in September injected trees. The recovery of 15N in September injected trees
was low due to early leaf drop. Fine roots (< 0.5 mm) maintained constant
levels of 15N exhibiting similar trends as with 14C. The loss of 15N from
the woody shoot and coarse roots was difﬁcult to explain since N was
thought to be tightly conserved. Turnover of the fine root system can not
explain this disappearance since the levels of 140 do not increase in the soil
(Tables 4.8- 4.9). However, C and N may not necessarily be linked. For
example, N may preferentially lost through amino acid and protein

exudation from roots.

Dynamics of 140 and 15N in Leaf Litter

The turnover of leaf litter was determined on plots containing
unlabeled trees. This was done to compare the amount of C and N input to
soil from the different tree litters. The decomposition of leaf litter 140
proceeded rapidly regardless of when the leaves were labeled (Table 4.12).
At 613 days, 29% of the July and 8% of the September leaf litter 1‘er
remained in the litter. Soil 14C accumulated rapidly to 248 mg of labeled C
or 17.1% of the 14C originally present in July labeled leaves. In the
September labeled litter, the amount of 14C in soil declines from 212 mg at
168 days to 166 mg at 613 days or 13% of that originally present in the litter.
The amount of 140 found in the microbial biomass was always less than 2%
of what was originally present for both leaf and root derived C.

The dynamics of leaf litter 15N differs from that of 140. Movement of
15N from leaf litter does not occur until 168 days for September and 613 days
for July labeled litter. This difference can be attributed to the C:N ratio, the
July labeled litter has a C:N ratio 2.5 times that of September labeled leaves.

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92

than the September litter. The July labeled litter mass was more typical of
the amount of litter that would fall and decompose under the rotation
conditions. Microbial biomass and soil 15N begin to accumulate at 168
days. The lower C:N ratio litter (September litter) releases 15N to the soil
faster and earlier than the higher C:N ratio litter. The microbial biomass
does not accumulate more than 5% of the applied 15N. By day 613, 61% of
July litter and 22% of the September litter 15N remained in the litter.

DISCUSSION

Carbon and N Allocation

The late season transport of photosynthate to stem and roots sinks for
storage is characteristic of hybrid poplars (Isebrands and Nelson, 1983).
Our data demonstrate the shift in C allocation from the acropetal (up) to
basipetal (down) orientation from July to September. The largest increase
in biomass C occurred in the stem and coarse roots. Fine roots (< 0.5 mm)
maintain a consistent standing biomass C of 5% of net primary production
from mid-growing season to the onset of dormancy. This low proportion of
net primary production in standing ﬁne roots is typical of plantation forests
(Santantonio and Grace, 1987 ; Pregitzer et al. , 1990).

In August, the above-ground tree components contained 82% of total
tree N. Leaves contained 60% of the total tree N. The major site of N
storage was in the woody shoot and coarse roots. Pregitzer et al. (1990)
found similar seasonal N distributions in one year-old hybrid poplars. The
two-year-old trees represented 3% of total N found in the plant-soil system.
Updegraﬁ‘ et al. (1990) found a similar above-ground-soil N balance in three-
year-old hybrid poplar plantations. These data exemplify the importance of

93

N distribution at centers of meristematic shoot activity during the growing
season and at storage sites in woody shoot and coarse roots during
dormancy.

In this study, microbial biomass N represented a large labile pool of
N which was equivalent to total tree N. Microbial biomass N turnover is
replenished by litter input and acts to conserve soil nutrients for plant
uptake. The major standing crop of litter N was found in leaf litter and
amounted to 50% of the N found in the microbial biomass. The standing
crop of ﬁne root N represented 5% of microbial biomass N. Signiﬁcant ﬁne
root turnover must occur to be equivalent to leaf litter N. However, turnover
of root biomass may not be the only means that N is returned to soil from
roots. Root exudation and mycorrhizal turnover may contribute to N input

to soil, but have yet to be studied carefully (Fahey, 1992).

The Fate of 1‘er

The speciﬁc activity is deﬁned as the ratio of labeled C to unlabeled C.
Any change in the ratio of the two C isotopes indicates a change in pool size
either through addition of unlabeled C or loss of labeled C. Caldwell and
Camp (1974) used the ratio of 1‘ltC/IZC in structural residue to evaluate
below-ground production and turnover in cool desert communities.
Milchunas et al., (1985) found this relationship to be invalid in plants when
stored reserves containing 14C were used to construct new tissue. After
estimating the amount of 14C used to construct new tissue, the researchers
found good agreement with actual production estimates. They suggested
that sampling for 14C dilution in the ﬁeld should be performed the year

after labeling to avoid problems with retranslocation.

 

 

 

94

The largest dilution of 14C activity occurred in stem and branches of
both labelings in the following year. A comparable loss of 14C occurred in
roots (> 10 mm) from the September labeling. The loss of 14C in roots (> 10
mm) from the September labeling as compared to no loss in July labeled
roots was probably associated with photosynthate storage. In July, the
actively growing tree was depositing C into structural residues, while in
September, non-structural C was being stored. Therefore, C assimilated in
September was more labile and subject to retranslocation and respiratory
loss. Loss of 14C from ﬁne roots (< 0.5 mm) occurred from labeling to the
second harvest in November for both labelings. No signiﬁcant change in
ﬁne root mass occurred during this period. Most of the dilution in 1‘er/ 12C
occurred in the 0-25 cm soil proﬁle and indicated that the production of ﬁne
roots or turnover of labile C accounted for approximately 50% of the
standing crop. The ﬁne root pool 14C/120 ratio remained unchanged from
dormancy into the next growing season. The lack of the dilution of 1‘er/12C
during this period indicated minimal root production, turnover and or the
synthesis of new roots from labeled reserves. Milchunas et al., (1992)
suggested that the movement of labile C may also have been used to support
new root growth in a short grass steppe. The use of reserve C in both ﬁne
root maintenance and construction has been demonstrated in seedlings of
Pseudotuga menziesii (Mirlu.) Franco (van den Dressche, 1985).

Horwath (Chapter 5) found that the amount of stored nonstructural
reserves (in glucose equivalents) in the root system was equivalent to the
standing crop of ﬁne roots. The reserves had double the 1‘ltC/12C ratio than
the ﬁne root residue C. Chung and Barnes (1977) estimated that it would
take 1.5 g glucose to synthesize 1 g of Pinus taeda roots. Assuming the

construction costs of roots between species are similar, the amount of root

%

reserves contained in these hybrid poplars was sufﬁcient to replace the ﬁne
root system 1.3 times with the addition of current photosynthate to dilute the
higher speciﬁc activity reserves. The cost of root construction does not
include maintenance activity of the root system during dormancy. The
reserves in coarse roots were not exhausted during the sampling period.
This would reduce the estimates of ﬁne root production unless coarse root C
was replenished from retranslocated reserves from the shoot.

The movement of C from the woody shoot to roots during and after the
dormant season would explain the low loss of labeled C from the root
system. Most studies on reserves in fruit trees have shown acropetal
movement of stored C to synthesize new leaves (Hansen, 1967; Quinlan,
1969; Kandiah, 1979). Lockwood and Sparks (1978) demonstrated the
acropetal movement of labeled reserves to synthesize leaves followed by a
basipetal shift back to the root system after leaves were fully expanded. In
this study, only 0.2% of the applied 14C was recovered in new leaves the
following year. The amount of labeled C lost from the woody shoot was
equivalent or less than the amount in the root system, and thus, would not
change the magnitude of the estimate of ﬁne root production from reserves

and current photosynthate.

The Fate of 15N

The activity and disposition of 15N within the tree was similar to the
fate of 14C. The stem injected 15N composed no more than 1% of tissue N in
both July and September labelings (Horwath, Chapter 3). The atom % 15N
excess varied among different tree components. Nommik (1966) found
similar differences in atom % 15N excess in various organs and in organs

of different ages from fertilizer 15N in Pinus silvestris L. and Picea abies

%

karst. This 15N distribution was attributed to the preferential loading karma“
active tissue, such as meristem regions. The majority of the injected 15N in
both July and September remained in the shoot 20 days after injection. In
September, the woody shoot receives the majority of 15N. The root system in
September accumulates twice as much 15N as compared to 'mid growing
season in July. However, the root system never receives more that 20% of
the injected 15N in July or September. Nommik et al. (1983) had also
demonstrated lower recovery of soil applied 15N in roots than shoots of
Pinus sylvestris L. 2 years after application.

The temporal shift in N allocation was exhibited by the distribution of
isotopically labeled N. During July, in mid-growing season, the majority of
N was found in leaves and was being used to support growing meristematic it
regions in the shoot (Dickson, 1989). During autumn, N storage begins in
the woody shoot and roots? Movement of 15N to the root system from the
shoot was observed in July trees. No movement of 15N occurred after the
September injection indicating that the storage of N occurs at or before -
budset. The constant atom % 15N excess of dormant roots and woody shoots
from the September labeled trees indicates no uptake of N occurred after
budset. During the September labeling, leaf drop was proceeding rapidly.
Weinbaum et al. (1978) found that 15NO3' uptake in non-bearing prune
trees was correlated to leaf retention. The authors attributed the presence
of leaves in supplying labile photosynthate for the energy requiring process
of NO3' uptake. Similarly, maximum uptake of 15N03' in Malus occurred
in August and the least in October after leaf fall (Hill-Cottingham and
Lloyd-Jones, 1975).

The loss of 15N occurred from both July and September trees during
the following year. Knowles (1975) reviewing 15N studies in forest

97

ecosystems found that 23-104% of 15N applied to the soil was recovered from
the plant-soil system. Recovery of 15N was dependent on the length of time
after application. The loss of 15N in this study from the tree-soil system
was 35% in July and 43% in the September injections, including the
amount recovered in leaf litter. The application of 15N-urea to Malus leaves
resulted in similar losses (Hill-Cottingham and Lloyd-Jones, 1975). The
average accounted for loss of 15N from the urea application was 32% during
the following growing season. Some of the 15N loss from this system may
have occurred through leaf leaching on to adjacent plots along the canopy
drip zone. Chapin and Moilanen (1991) estimated loss of N from leaching
by autumn rains in Betula papyrifera to be 25% of leaf N. Gaseous loss of N
from leaves has also been suspected (Luxmore et al., 1981).

The signiﬁcance of N turnover through the root system seems
negligible since the standing crop of ﬁne roots contained less than 2% of the
injected 15N. No signiﬁcant dilution of 15N occurs in the ﬁne root pool from
November into the following growing season for both July and September
injections. The production of new roots from higher atom % 15N enriched
pools of N in coarse roots must be considered as it was with 14C. The
tracers 14C and 15N behaved similarly and there is no reason to believe they
would be utilized differently in the construction of ﬁne roots. Clark (1977)
observed that the 15N content of roots in a short grass prairie remained
constant over a ﬁve year period. He attributed the constant level of 15N in
roots to efﬁcient recycling and uptake of mineralized N from the
decomposition of above-ground tissue. However, in our system the leaf
litter was collected and replaced with unlabeled litter, eliminating 15N
recycling through leaf decomposition. The dilution of atom % 15N excess

occurred only in July labeled coarse roots and in the shoots from both

%

injections. Apparently, the uptake of N to dilute these pools does not
interact with N contained in ﬁne roots. The stability of the dual tracers in
the ﬁne root pool may suggest that this component is long-lived in hybrid
poplars.

The Turnover of 14C and 15N

The intent of litter residue decomposition studies is to determine the
amount of C and nutrients entering the active soil organic matter fraction
and microbial biomass (Paul and Juma, 1981). The amount of C and N
entering this labile fraction has been studied extensively with isotopically
labeled residues in agronomic systems (Shields and Paul, 197 3; Jenkinson
and Rayner, 1977; Ladd et al., 1981; Voroney et al., 1989). These studies
have used unconﬁned litter to determine the processes involved in
conserving the labile active soil fraction and associated nutrients. These
processes have been described as a sum of events occurring in two phases;
the initial rapid mineralization of substrate and incorporation into
microorganisms (Shield et al., 1973; Amato et al., 1984), followed by a
slower mineralization of microbial products and resistant plant materials
(Jenkinson and Rayner, 1977 ; Voroney et al., 1989). Researchers studying
these processes in forest systems describe the above process as a short
period of nutrient accumulation and a period of slow release from
decomposing litter conﬁned in bags (Melillo and Aber, 1982; Berg, 1988).

The 1‘er pool dilution technique to estimate below-ground production
and turnover was found to work well on recalcitrant biomass components,
such as lignocellulose (Caldwell and Camp, 1974). Milchunas and
Laurenroth (1992) using this technique found anomalous root production

and turnover estimates in short grass steppe and suggested that non-

99

uniform labeling and movement of labeled C were the cause. Estimating
and correcting for the synthesis of new roots from reserves, root turnover
was calculated to be 8 years in these perennial grasslands. Conversely,
estimates of root production were zero in 2 of 4 years using the method of
changes in standing crop. This slow rate of below-ground production was
realistic since the added 14C was only sufﬁcient to tag biomass pools, and
therefore, the absolute amount was limiting in the production of new
biomass. The slow loss of 14C from root system of trees has been observed
in previous studies (Hansen, 1967; Schier, 1970; Lockwood and Sparks,
1978). Unfortunately, these tree labeling experiments have used potted
plants and their comparison to natural systems is inappropriate.

In this study, the 14C activity of structural residue of ﬁne roots does
not change over the year sampling period for both labelings (Horwath,
Chapter 5). The approach of Caldwell and Camp (1974) would indicate no
turnover of the ﬁne root pool. However, our study shows an increase in soil
14C and decrease in 14C activity of the labile ﬁne root pool. The possibility
that the labile pool was turning over through root exudation and
mycorrhizal maintenance must be considered. Vesicular-arbuscular
mycorrhizal infection rates were observed to be 50% of the ﬁne root surface
in the 0—25 cm soil proﬁle (Horwath, personal observation). Most of the 14C
dilution of ﬁne roots occurred in the 0-25 cm soil proﬁle where mycorrhizal
infection was the greatest. The largest addition of 14C to soil occurred at
the same time 14*0 dilution occurred in the ﬁne root pool. No 1‘er dilution
occurred from November to the one year sampling for both labelings and no
signiﬁcant increase in soil 1‘er was detected.

The stability of microbial material in soil has been studied extensively

and has been suggested to be important in conserving soil organic matter

100
(Shields and Paul, 1973; Voroney et al., 1989). The turnover of microbial

material from mycorrhizal maintenance or microbial utilization of root
exudates are possible sources of soil 14C. The turnover of 14C stabilized into
the soil active organic matter from the root labeled plots amounts to 1-3% of
the 14C applied from both labelings. Though no signiﬁcant ﬁne root
standing crop production or turnover could be demonstrated with the
traditional 14C pool dilution techniques, signiﬁcant amounts of labeled C
turnover occurred through synthesis of new roots from reserves, exudation
and or mycorrhizal turnover. This C input was playing an active role in
maintaining soil organic matter.

The decomposition of the isotopically labeled leaf litter proceeded at a
rapid rate. The loss of 14C from September labeled leaves occurred more
rapidly than from July labeled leaves. This discrepancy can be attributed to
the labeling of the leaf constituents. During July, the leaf growth was
occurring, therefore, uniformly labeling the separate leaf constituents.
Conversely, in September the leaves were mature and primarily
synthesizing carbohydrates for export. The C/N of the September leaves
was lower explaining the more rapid decomposition (Melillo and Aber,
1982). The difference in litter quality within the same stand and year has
been observed in Populus tremuloides to affect rates of decomposition
(Taylor and Parkinson, 1988).

The fate of 15N differs substantially from that of 14C in the dual
labeled leaf litters. Signiﬁcant 15N movement from decomposing leaf litter
does not occur until 328 days for September and 613 days for July labeled
leaves. The lower C/N September leaves released labeled N at a faster rate
the higher C/N July leaves. The release of 15N in July leaves occurs when
the C/N of the decomposing litter reaches a C/N of 13, similar to the

 

101

September litter. However, in both litters the accumulation of 15N in soil
and microbial biomass occurs at 168 days. The amounts of 15N in soil,
microbial biomass and litter exceeded the amount applied at day 168 for
September labeled leaves. This error was partially explained in that the
litter was lain uniformly on to a 1 m2 area by distributing a known quantity
on the soil surface. Subsequent samplings of one quarter of each plot may
have been subject to error by non-uniform litter distribution across the plot.
The high standard errors of the mean for soil 15N indicate that the
discrepancy lies in the 15N recovered from soil. Conversely, the low
amount of error and 15N found in the microbial biomass does indicate
movement of 15N into soil. During the second year of decomposition at 613
days 40-80% of litter 15N has transferred to soil. Since the amount of N in
litter was 10 times the amount found in the standing crop of ﬁne roots, the
return of N to soil for future uptake was dominated by leaf litter N in this
system during stand initiation.

The amount of 14C entering the soil was 13-17% of the initial litter
content. The 1‘er speciﬁc activity of the microbial biomass under
decomposing leaf liter was approximately 10 times that of microorganisms
utilizing root derived labeled C (data not shown). Yet the amount of labeled
C stabilizing in soil was similar between the two litters. The distribution of
leaf 14C in soil was mainly in the surface soil (0-10 cm) while root C input
occurred over the entire soil proﬁle (0-100 cm). These differences in C input
to soil demonstrate the role of these litters in the spatial conservation and
formation of soil organic matter. This would imply that the litters also have
a spatial role in developing soil quality.

102
CONCLUSION

The amount of C entering the soil was similar for both leaves and
roots. The low estimates of root production and turnover by 14C pool
dilution and potential synthesis from reserves could not account for this C
input into soil. The C entering the soil from the labeled root system may
have come from sloughing, exudates and or, mycorrhizal turnover.
Nitrogen cycling through the root system to soil could not be demonstrated.
The constant atom % 15N and labeled N in the root system implies minimal
turnover through root biomass loss. Soil microbial biomass obtains more
substrate from leaf litter than it does from root C based on 14C enrichment
of microbial C. This may indicate that more stabilized organic matter will
form from leaf litter derived C than from root C. Both litters play an active
spatial role in conserving the soil organic matter with leaf litter primarily
enriching surface soil with C and N. In this system, it is important to
retain leaf litter on site to conserve available soil N to satisfy these fast-
growing nutrient-demanding trees during the initial phases of stand

generation.

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Vogt, K. A., C. C. Grier, S. T. Gower, D. G. Sprugel, and D. J. Vogt. 1986.
Overestimation of Net Root Production: A Real or Imaginary
Problem. Ecology. 67:577-579.

Voroney, R. P. and E. A. Paul. 1984. Determination of Kc and Kn in situ for
the calibration of the chloroform fumigation-incubation method. Soil
Biol. Biochem. 16: 9-14.

Voroney, R. P., E. A. Paul, and D. W. Anderson. 1989. Decomposition of
wheat straw and stabilization of microbial products. Can. J. Soil Sci.
69:63-77.

Wardlaw, I. F. 1990. The control of carbon partitioning in plants (Tansley
Review No. 27). New Phytol. 116:341-381.

Warembourg, F. R., and E. A. Paul. 1977. Seasonal transfers of assimilated

14C in grassland: Plant production and turnover, soil and plant
respiration. Soil Biol. Biochem. 9:295-301.

Weinbaum, S. A., M. L. Merwin, and T. T. Muraoka. 1978. Seasonal
variation in nitrate uptake efﬁciency and distribution of absorbed
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103:516-519

Wollum, A. G. and C. B. Davey. 1975. Nitrogen accumulation,
transformation and transport in forest soils. In: Forest soils and land
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4th North American Soils Conf. Les Presses de l'Universite Laval,
Quebec.

Chapter 5

THE DYNAMICS OF ASSIMILATED 14C02 IN THE CHEMICAL
FRACTIONS OF THE SHOOTS AND ROOTS OF POPULUS
PLANTATION S

ABSTRACT

The allocation of carbon to the chemical fractions of perennating
tissue is characterized by a series of competing centers of growth and
storage inﬂuenced by seasonal growth patterns. This research was
conducted to distinguish between the utilization of current and reserve
photosynthate for the long-term maintenance of the shoot and root system of
hybrid poplars. Trees were pulsed labeled with 14002 in the ﬁeld during
the growing season of their second year of growth to encompass seasonal
growth patterns. Tree tissue was fractionated into nonstructural (lipids,
sugars, protein and starch) and structural cell wall (cellulose and lignin)
components. During the mid-growing season, between 0.5-2.7% of the
assimilated 14C was in sugars. In autumn, up to 4.6% of the radiolabeled
C was in the sugar fraction. Less 1‘er was found in structural components
late in the growing season. The distribution of labeled chemical fractions
was similar to the distribution of the unlabeled fractions indicating that
assimilated 14C was following normal translocation pathways. In both the
mid-growing season and autumn labeling, the speciﬁc activity of the sugar,
starch and residue fractions of ﬁne roots changed little from dormancy to
one year after the radiolabeling procedure. The lack of decline of 14C
activity in the ﬁne root cell wall residue indicated a lack of ﬁne root
turnover or the utilization of reserves to maintain ﬁne root during the next

growing season.

110

111
INTRODUCTION

The allocation of carbon to perennating tree tissues is characterized
by the distribution of photosynthate between competing sinks. Normally,
leaves are considered sources of assimilate production, but when
immature, leaves both produce and import photosynthate (Dickson, 1989).
Mature leaves export photosynthate to competing centers of growth and
storage. Carbon ﬂow early in the growing season is predominately in the
acropetal direction in perennial plants supporting the growth of the shoot
(Donnelly, 1974; Isebrands and Nelson, 1983; Dickson, 1989). The amount of
current photosynthate used for shoot growth will differ among species
according to shoot type and growth pattern. Heterophyllous species, such
as hybrid poplar, require more photosynthate for shoot growth because of
continual growth and longer growing seasons (Kozlowski and Keller, 1966).
As the season progresses, lower leaves of hybrid poplars export C
basipetally to the stem and root system (Isebrands and Nelson, 1983). Late
in the season large quantities of photosynthate move to the root system,
bypassing branch and stem sinks. This seasonal distribution of
photosynthate is common among many tree species with varying shoot and
growth patterns. The basipetally exported assimilates are stored as
carbohydrates, lipids and nitrogenous compounds (Dickson, 1989). These
reserve C compounds are used for cold hardening of tissues, respiratory
maintenance and for the initiation of growth the following season
(Kramer and Kozlowski, 1979). The importance of reserve material can
not be underestimated since the availability of these substances will

determine the success of the tree during the next season.

 

112

The seasonal variation of storage compounds in trees has been the
object of many studies (Ziegler, 1964; Kozlowski and Keller, 1966; Glerum,
1980; Loescher et al., 1990). Deciduous trees must transpose seasons when
no photosynthesis can occur and are, therefore, reliant on stored reserves
from the previous season for the maintenance of life and resumption of
growth in the spring (Priestly, 1970; Krammer and Kozlowski, 1979).
Gymnosperms, on the other hand, utilize both current and stored
photosynthate to maintain life and growth throughout the year (Glerum,
1980). The highest concentration of reserves is usually found in roots while
the major quantity is found in the above-ground portion of the tree as a
result of the accumulation of biomass in the stem (Krammer and
Kozlowski, 1979). The use of reserves in respiration during dormancy, in
leaf initiation and expansion, in cambial growth of woody tissue and root
growth are not well characterized for most trees (Loescher et al., 1990).

Patterns of carbohydrate, fat and protein distribution and turnover
directly inﬂuence phenological expression and plasticity as a result of
competition, abiotic interactions and cultural practices. However, it is
extremely difﬁcult to distinguish current and reserve photosynthate in
order to resolve their origin, transformation and utilization. The
radiolabeling of assimilates can distinguish between utilization of current
and reserve photosynthate. Few radiolabeling studies have been done
under ﬁeld conditions to characterize the temporal fate of radiolabeled
photosynthetic products (Dickson, 1989; Loescher et al., 1990).

To characterize the fate of reserves in trees I radiolabeled hybrid
paplars in their second season of growth. The radiolabeling procedure was
done under ambient conditions of 002 and temperature on replicate trees

for one photoperiod. The labelings were done to encompass two distinct

113

physiological growth patterns of rapid shoot elongation and the autumn
basipetal movement of photosynthate. Sampling of the labeled whole trees
was done for a period of one year. The intent of this research was to
characterize the fate of radiolabeled chemical fractions, especially of
nonstructural and structural carbohydrates and residue. I hoped to
discern the origin and utilization of assimilates and to distinguish the
usage of current from reserve in the maintenance of ﬁne roots. This
chapter describes the radiolabeling of rapidly-growing, intensively cultured
hybrid poplars and the fate of radiolabeled photoassimilates over time.

MATERIAL AND METHODS

Plant Material

Cuttings of Populus euramericana cv. Eugenei, 10 cm in length,
were planted on a 2 by 1 m spacing in the spring of 1989. The site was
located at the Kellogg Biological Station, LTER site, Michigan State
University, southwest Michigan. The Kalamazoo soil series is classiﬁed as
a ﬁne loamy, mixed, mesic, Typic Hapudalf. Eight different trees were
exposed to 14C02 in a large Plexiglas chamber on July 19 and September 5,
1990. The radiolabeling procedure was described previously (Chapter 3).
Ambient 14C02 concentration (290-350 ppm) and temperature were
maintained for the entire photoperiod. The level of 14C02 was maintained
with an Infra Red Gas Analyzer that was interfaced to a computer operated
14C02 generator. The chamber air temperature was controlled by forcing
the enclosed air though a water-cooled heat-exchanger. In July, 3.04x108
Bq of 14C02-C with a speciﬁc activity of 4215 Bq mg“1 C02-C for an average
of 3.79x107 Bq tree'1 were added to the labeling chamber. During the

114

September labeling, 2.85x108 Bq of 14C02-C with a speciﬁc activity of 4300 Bq
mg‘1 C02-C for an average of 3.56x107 Bq tree -1 were assimilated.

Sampling Procedure

Three randomly selected trees were sampled two weeks after each
labeling. An additional three trees from each labeling were sampled in
November, 1990 and the remaining two trees were sampled one year latter.
The sampling included all above- and below-ground tree material located
within a 1 m3 soil block. Leaves, branches and stem were separated by
height and age (Chapter 3). The soil block was excavated by depth (0-25 cm,
25-60 cm and 60-100 cm) and the soil from each depth was thoroughly mixed
and subsampled to recover roots (< 0.5 mm) by hydronuematic elutriation
(Smucker et al., 1982). Roots were sorted by hand into the following size
classes; < 0.5 mm, 0.5-1 mm, 1-3 mm, >3 mm, >10 mm and cutting. Plant
material was dried at 80°C for 48-96 hours and was ground in a Wiley mill
to pass a 60 mesh screen. Tissue samples (leaves, branches, stem and all
root classes) were composited by the proportion of weight found in all

sources of that particular tissue.

Analysis of Chemical Fractions

Plant tissue was fractionated into chloroform solubles (lipids, cutin,
suberin and pigments), sugars, protein, starch and structural residue
(Figure 5.1). Homogenized plant material (25-30 mg) was weighed in
triplicate into 50 mL centrifuge tubes (Sardstedt , Newton, NC). The tissue
was extracted [modiﬁed Bligh and Dyer (1959)] at room temperature with 4
mL of methanol: chloroform: water (12:5:3, vzv). After the addition of the

solvent, the sample was sonicated for 45 seconds in a water bath, allowed to

115

 

Plant tissue 25-30 mg
Homogenize in 4 mL CH30H:CHCI3:H2O (12:5:3)
Centrifuge 1300 g 10 min
Repeat 3 times
Supernatant ' l ’ Pellet
Add 5 mL CHC13 and 4 mL H20 Pronase digestion _>Protein
Seperate phases
ll
1 l l
CHgOH/HgO CHCl3
Amylase digestion _> Starch
1 Fat, cutin, suberin, etc.
Ion exchange Column
l Residue
Sugars

Figure 5.1. Diagram of the tissue fractionation protocol.

116
stand for 10 minutes, and centrifuged (1300 g, 0°C, 10 minutes) for a total of

four extractions. The supematants were combined into another centrifuge
tube. The pellet was set aside for subsequent analysis. The supernatant
was combined with 5 mL of CHCl3 and 4 mL water, vortexed, and then
centrifuged (1300 g, 0°C, 5 minutes). The chloroform and methanol-water

phase (top) were separated and analyzed separately.

Analysis of the Chloroform phase

The chloroform phase was transferred to a dried (24 hours at 45°C)
pretared scintillation vial and evaporated for 24 hours under a fume hood.
The vial was then dried overnight at 45°C. The vial was reweighed to
determine chloroform soluble residue. The residue was digested with 1 mL
of methylbenzonium hydroxide (Sigma, St. Louis, M0) by wetting the sides
of the vial, capping and incubating at 50°C for 24 hours. Hydrogen peroxide
(100 uL) was added to highly colored samples (i.e. leaves) to remove color.
Acetic acid (0.5 mL) was added to the digested residue to quench
chemiluminescence. Finally, 10 mL of scintillation cocktail (Research
Products International Corp., Mount Prospect, IL) were added, and the
samples counted in a liquid scintillation spectrometer (Packard
Instrument Co., Downers Grove, IL). The CHCl3 soluble residue was
assumed to be 75% C by weight for 14C calculations (calculated from
Robertson, 1980).

Analysis for Sugars
Neutral sugars were separated from amino acids, organic acids and
sugar phosphates by eluting the methanol-water phase through tandem

cationic and anionic exchange columns. The columns were prepared from

117

cation exchange resin (DOWEX 50W; 50x8-400; 200-400 mesh, Sigma, St.
Louis, MO) and anion exchange resin (DOWEX 1; 1x8-400; 200-400 mesh,
Sigma, St. Louis, MO) washed in an equal volume of acetone overnight on a
stir plate. The acetone was decanted and the resin washed in an equal
volume of methanol for 8 hours. The methanol was decanted and the resin
washed 3 times with puriﬁed water. The resin was then covered with 2
volumes of 1.0 M HCL and slowly heated on stir plate to 60°C and repeated.
The resins were then washed with water to a neutral pH. The cationic
exchange resin was washed two times in 80% ethanol and stored under
80% ethanol at room temperature. The anionic exchange resin was
washed with 2 volumes of 1.0 M NaOH and then drawn through a #1
Whatman ﬁlter with a vacuum and sidearm Erlenmeyer ﬂask. This
procedure was repeated with fresh NaOH for a total of 20 washes. The
anion exchange resin was then washed to a neutral pH with water and
then washed twice with two volumes of 1.0 M Formic acid. The anion
exchange resin was washed to a neutral pH with water, washed twice with
80% ethanol and stored under 80% ethanol.

Ion exchange columns were prepared from Pyrex tubing (15 cm long,
7 mm I.D.) for the cation exchange resin and 4 mL pasteur pipette (Fisher,
Fairlawn, NJ) for the anion exchange resin. The tips of the columns were
plugged with glass wool and loaded with a slurry of resin under 80%
ethanol to a bed length of 4 cm. Columns were washed several times with
80% ethanol to a neutral pH and connected with Tygon tubing (cation over
anion). A syringe needle (18 gauge) was inserted downwards into the
Tygon tubing to avoid an air lock forming between the two columns. The
tandem columns were placed with the tip of the pasteur pipette into a 50 mL

volumetric ﬂask. The methanol-water phase from the initial extraction

118

was applied to the column and eluted with 80% ethanol until the volumetric
ﬂask was nearly full. The volume of the ﬂask was brought to 50 mL with
80% ethanol and the elutent analyzed for 14C and total sugars.

Total sugars were measured colormetrically using the phenol-
sulfuric acid technique (Dubois et al., 1956). Glucose standards were
prepared by drying dextrose (J. T. Baker Inc., Phillipsburg, NJ) at 80°C and
preparing a standard series in methanol-water eluted through ion
exchange columns. The elutent (5 mL) was transferred to a scintillation
vial and dried at 50°C. Water (1 mL) was added to hydrate the dried sugar
and the 1‘er activity was determined in a liquid scintillation spectrometer
after the addition of 10 mL of scintillation cocktail. The sugars were
assumed to be 40% C by weight for 14C calculations.

Analysis for Proteins

Protein was hydrolyzed and separated from the plant tissue by
proteolytic digestion (Dickson, 1979). The pellet remaining after the
methanol-chloroform-water extraction was incubated with 2 mL 0.4%
Pronase (Calbiochem, La Jolla, CA) in 0.05 M Tris pH 7.4 and 0.1 mL 0.01
M sodium azide to prevent microbial contamination. The pellet-Pronase
mixture was incubated at 30°C for 48 hours, with occasionally swirling.
The pellet was then centrifuged (1300 g, 0°C, 10 minutes) and the
supernatant drawn off with a pasteur pipette. The pellet was re-extracted
with 2 mL water three more times. The supematants were combined,
freeze-dried and rehydrated with 1 mL of water. An aliquot (0.25 mL) was
combined with 5 mL of scintillation cocktail and 14C activity determined on
a liquid scintillation spectrometer. Total protein was determined using a

ninhydrin technique (Lee and Takahashi, 1966). For each set of samples,

119

1% albumin bovine (Sigma, St Louis, MO) was hydrolyzed and used to set
up a series of standards. Protein was assumed to be 51% C by weight for
140 calculations.

Analysis for Starch

Starch was hydrolyzed to glucose by enzymatic digestion with B-
amylase (Dickson, 1979). The d-amylase enzyme Clarase 5000 (Miles
Laboratory, Elkhart, IN) was dialyzed to remove contaminates. The
dialysis tube (Spectrum Medical Industries Inc., Los Angeles, CA),
molecular weight cutoff of 12,000-14,000, was prepared by autoclaving in 2%
sodium bicarbonate and 1 mM EDTA for 10 minutes. A 0.5 % Clarase 5000
solution was dialyzed in water for 48 hours at 4°C, changing the water
frequently. The volume of the enzyme solution was determined and
combined with sodium acetate to achieve a 0.1 M (adjusted to pH 5.5 with
acetic acid) enzyme-buffer solution. The pellet remaining after the protein
hydrolysis was boiled with 1 mL 0.1 M sodium acetate buffer to gelatinize
the starch. After the pellet was brought to room temperature, 1 mL of
Clarase 5000 buffer and 0.1 mL 0.01 mM sodium azide were added to each
sample. The samples were incubated for 48 hours at 50°C, swirling the
samples occasionally. Starch in the samples was determined with
peroxidase-glucose oxidase-o-dianisidine dihydrochloride reagent (Sigma
Chemical Co., St. Louis, MO). A glucose standard series was developed
from a standard glucose solution, 1 mg mL'1 (Sigma Chemical Co., St.
Louis, MO). Starch was assumed to be 40% C by weight for 14C

calculations.

12)

Analysis of the Structural Residues

The pellet remaining after the starch hydrolysis was transferred into
a dried (50°C for 24 hours) and pretared glass vial with 100% ethanol. The
samples were dried at 45°C and reweighed. Total C was determined on a
Gas Chromatograph-Mass Spectrometer (Europa Scientiﬁc, Crewe,
England) using 44% C (as cellulose) Whatman no. 1 ﬁlter paper as a
reference standard (Harris and Paul, 1989). Radiolabeled C from the GC-
MS exhaust was collected in C02 absorbing scintillation cocktail (Harvey
Instrument Co., Hillsdale, NJ) and analyzed on a Liquid Scintillation

Spectrometer.

RESULTS

Distribution of Chemical Fractions.

The concentration of biochemical fractions varied seasonally and
among tree components. The concentration of sugars was 4-13% (dry
weight) in July and September in above-ground tissue and 1-6% in the root
system (Tables 5.1-5.2). Leaves had the highest concentrations of sugar
ranging from 10-13% and was no doubt due to the fact that leaves were sites
of sugar synthesis. The levels of sugar were 6% in branches and 4.1-4.8%
in coarse roots (> 3 mm) in diameter during the growing season. During
dormancy in November, the concentration of sugar in roots (> 3 mm)
increased to 7-10% of the dry weight. Regardless of the season, the
concentration of sugars in roots was proportional to root diameter and
probably related to the amount of ray parenchyma cells available for
storage. Leaf litter contained a signiﬁcant amount of sugar, 3% by weight.

The remaining sugar in leaves was probably a result of translocation

121

 

 

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constraints from the unloading and loading of phloem cells with
photosynthate during autumnal photosynthesis.

The concentration of starch in roots (> 1 mm) always exceeded starch
levels found in the above-ground tissue (table 5.1-5.2). In July, the
concentration of starch in ﬁne roots (< 0.5 mm) was 0.5% and large roots
contained up to 12.8%. Levels of starch increased in ﬁne roots to 2-3% and
up to 19% for large roots during September and November. The above-
ground tissue contained between 1-5% starch from July to November. Leaf
litter contained 1% starch. The concentration of starch in roots was
proportional to diameter and was similar to the distribution of sugar in
roots. The cutting always contained less starch and sugars than large
structural roots and more closely resembled carbohydrate concentrations of
the stem tissue.

Protein levels generally comprised 1-5% of tissue throughout the
growing and dormant seasons. The protein concentration of leaves was
4.6% in July and 7.2% in September. The concentration of leaf litter protein
changed little from that of green leaves. Root protein levels increased from
about 1% in July to 5% in November. Coarse roots contained more
hydrolyzable protein than ﬁner roots and were similar to protein levels
found in the stem. Protein content was proportional to root diameter except
that roots (> 10 mm) and cutting contained less protein. The protein
determination using Pronase is not effective in removing cell wall bound
protein. The protein determined is indicative of cellular proteins not
associated with the cell wall. There is no reason to suspect any changes in
the concentration of cell wall structural protein in the woody shoot and
roots throughout the season, consequently, the increase in tissue protein

was probably associated with the accumulation of storage protein.

124

The chloroform-soluble fraction is composed of lipids, cutin, suberin,
waxes and glycerated substances, such as phospholipids and glycolipids.
This fraction is an integral component of cells involved in photosynthesis,
storage and can itself be a form of C storage. The chloroform fraction
varied in all tissues during the growing and dormant season. In July, this
ﬁ‘action composed approximately 6% of branch and stem tissue and 8-9% of
leaves and roots. An increase in the chloroform fraction in roots (< 10 mm)
occurred in September. Conversely, this fraction decreased in roots (< 10
mm) from July to November. An increase of chloroform soluble material
occurred in branches from September to November. The increases found in
these fraction can be associated with a storage function, but many of the
constituents of this fraction are also involved in other cellular functions,
such as cold acclimation.

The residue fraction is composed of transformed sugars (cellulose,
hemicellulose and pectin), lignin and cell wall protein. The trees were
composed of 50-88% residue throughout the growing and dormant seasons.
The branches and stem contained the greatest concentration of residue by
weight. Leaves generally contained 50% residue, while roots (3-10 mm)
contained similar or less residue than leaves during November. Roots
contain a higher proportion of ray parenchyma cells resulting in less
residue than found in stem tissue. The percentage of storage cells in roots
(3-10 mm) increased as food reserves accumulate during the dormant
period. There was a general decline in residue content of woody tissue

during dormancy.

125

Distribution of 14C Within Chemical Fractions

The data in Tables 5.3 and 5.4 show the amount of 14C (%) contained
within the C component of each biochemical fraction. The data exempliﬁes
the relative sink strength of storage, metabolic and structural pools of the
different tissues over the growing season. In July, 1-1.5% of the
radiolabeled C was found in the sugar pool throughout the tree. The similar
levels of 14C in the different tree components shows the uniformity of
labeled C distribution. The quantity of 14C derived from the labeled
assimilates was inversely proportional to root diameter with the ﬁne roots
(< 1 mm) containing the highest amount of 140 two weeks after the
labeling. Labeled C in the starch pools ranged from 0.5-2.7% with leaves
having the highest proportion and the cutting the least. The C derived from
labeled C in root starch was slightly higher than in the sugar pool and was
also inversely distributed according to root diameter. The increase of 14C in
the composition of the starch pool indicates that equilibrium distribution of
14C was shifting to starch from sugar as a result of new unlabeled C
diluting the sugar pools. This indicated that even after two weeks following
exposure to 140 the radiolabeled photosynthate in nonstructural
carbohydrates was still equilibrating. Root respiration studies in Chapter 3
indicated that a minimal amount of 14C was being respired after two weeks
from the root system. Therefore, the sugar pool was in some way
compartmentalized from current photosynthate since root respiration had
been mostly diluted with unlabeled C. The compartmentalization of
current assimilates from reserve sugar suggested that current
photosynthate was the source of C for root respiration during mid growing

8688011.

1%

In autumn, sugars in roots (<10 m) contained approximately 4% of
the labeled C. Coarse roots (> 10 mm) and cutting contained approximately
2.5% of the 14C. The above-ground woody tissue contained 1-1.5% labeled C
and leaves possessed 2.1%. Root starch contained 1.4-1.9% 14C and,
therefore, contained less 14C than the sugar pools in contrast to July
results two weeks after labeling. The woody shoot tissue contained 0.7%
14*C in starch and leaves contained 3.7%. The 1‘er activity of the starch pool
was now proportional to root diameter, differing from the July results,
while the sugar pool showed no relationship to root size. The labeled sugar
pool was not contributing signiﬁcantly to root respiration two weeks after
labeling. Again, this indicated that the labeled sugar pool was separate
from current photosynthate in supplying the substrate for root respiration.

In November, the 14C comprising the sugar and starch pools from
both labelings had decreased due to utilization and dilution with unlabeled
C. The lone exception was the root starch pools from trees labeled in
September. These showed little change except for a decline in roots (> 10
mm) and the cutting. The starch deposited in roots (< 10 mm) in September
had not accumulated any additional unlabeled C (Table 5.4). This implied
that the storage of photosynthate in the root starch pool had been completed
in early September before photosynthesis had ceased. The 14C in the root
sugar pool was diluted from approximately 4% in September to 1% in
November. The dilution of the root sugar pool occurred both from unlabeled
C and utilization since the concentration and mass of sugar in roots
declined only slightly. The utilization of the root sugar pool may have been
indicative of the transition from current to storage photosynthate for root

maintenance as dormancy approached.

 

 

 

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129

The 14C content of the protein pools was similar to the sugar pools
following the labeling periods and November sampling (Table 5.3-5.4). In
July, root and leaf 1‘lC-protein accounted for 1-1.9% of the protein C and
was distributed in the roots proportional to diameter. In the above-ground
tissue, protein was signiﬁcantly labeled in branches (2.6%) and less in the
stem (0.5%). The increased 1‘er content of root protein in September
corroborated the basipetal flow of C during autumnal photosynthesis.
Labeled C in the protein of roots amounted to 1.9-4.3% of the C content in
September. The woody shoot protein C, including the cutting, contained
approximately 1% 140 and leaf protein C contained 2.1%. Large dilutions of
14C occurred after each labeling and was exempliﬁed by lower values for C
derived from 14C in all tissue except litter, which remained relatively
unchanged. This indicates that the protein pool turns over rapidly during
the growing season through dormancy.

The labeled C in the chloroform soluble fraction, though lower than
any other fraction followed a distinctly different pattern than either the
nonstructural carbohydrates or protein. Generally, root chloroform soluble
material increased in 14C content from summer to autumn. In roots (> 10
mm) and above-ground tissue, the percentage of 14C declines from July and
September to November. The decline was most likely due to dilution of the C
component of these tissues. The increase in the percentage of 140 in the
chloroform soluble C in the roots (< 10 mm) was probably due to cambial
activity for diameter growth, storage and winter hardening requirements.
The 14C needed to enrich these pools was no doubt taken from reserves
which were signiﬁcantly more labeled.

The disposition of 14C in the residue C pool followed closely the
seasonal ﬂow of C to active sinks of growth and storage. During July, the

13)

14C content in the residue of above-ground tissue was 20 to 50% greater
than the root system (Table 5.3). The above-ground preferentially utilized
photosynthates to support shoot synthesis. As autumn approached, the
roots became active sinks for photosynthates with residue C comprising 2-5
times more 140 than residue in branch or stem tissue. The residue pools
became diluted with unlabeled C from the addition of new biomass from
July to November. The dilution of the residue pool during the September
labeling was less due to storage of photosynthate taking precedent over the
synthesis of cell wall components. The 14C content of residue was lower
than nonstructural carbohydrates and protein indicating that the
assimilated 140 had not equilibrated between structural, metabolic and
storage pools two weeks following exposure in summer or autumn. This
lack of equilibrium indicated that storage of reserve material in roots had
already begun during mid growing season. This trend became more
evident in autumn when starch and sugar pools contained 2-3 times more
14C than the residue. A similar trend was evident for the above-ground

components in September.

Redistribution of 14C Over Time

The nonstructural carbohydrates, protein and chloroform soluble
pools of roots (< 0.5 mm) contained signiﬁcant amounts of 140 one year
latter (Tables 5.5 and 5.6). The residue C of ﬁne roots contained a similar
amount of 14C to that found in November of the previous year. The biomass
of ﬁne roots increased slightly during the next growing season (Figure 5.6).
The 14C composition of the ﬁne root residue may have been sustained

through the utilization of labeled reserves. It was not possible to determine

131

 

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133

the metabolic state of these roots, therefore, the measurement of 14C activity
of this root class encompassed both live and dead roots. The residue pool of
ﬁne roots had similar amounts of 14C indicating the importance of reserves

in the both synthesis maintenance.

Distribution of 14C in Chemical Fractions

Figures 5.2-5.4 represent the distribution of sugar, starch and
residue within the tree and the 140 contained within each fraction. In
July, the bulk of tree sugar was located in the shoot. The 14C-labeled sugar
was distributed according to the distribution of sugar within the tree. This
indicated that the above-ground tissue was a strong sink for labile
photosynthates as well as recently ﬁxed C during mid-growing season. The
distribution of 14C-sugar became diluted in roots (> 1 mm) and stem from
; the utilization and addition of unlabeled 0 (Figures 5.2a-5.2b). The
September labeling showed that the predominant ﬂow of C was basipetal.
By November, some equilibration between the root system and above-ground
tissue had occurred with leaf litter retaining proportionately more 14C in
the sugar pool.

The distribution of starch and 14C-starch was always highest in the
root system (Figures 5.3a-5.3b). The distribution of starch and 14C starch
between shoot and root was relatively evenly distributed for the July labeling
and November sampling (Figure 5.3a). The location of labeled and
unlabeled starch was mainly in the root system in September to November
(Figure 5.3b). Again, leaves and leaf litter contained proportionately more
labeled starch and was similar to leaf 1‘itC-sugar distribution.

134

Figure 5.23. The distribution of total sugars and labeled 14*C--sugars among
the shoot and root components. The July labeling with subsequent
sampling August 2, 1990 and November 2, 1990 show the acropetal
translocation tendency of sugars synthesized in July during mid-
growing season. Standard error of the mean shown as line bars.

 

   
   

 

 

 

 

 

 

 

Figure 5.2a
August 2, 1990
60
3 ar
50 _ 12g
C-sugar
40 _.
e2 a) —
m _. ..................................
10 _. ............................. _
0 a _
leaves branches stem root root
< 1 m > 1 mm
November 2, 1990
so

 

 

 

 

 

 

leaves branches stem root root

136

Figure 5.2b. The distribution of total sugars and labeled 140- sugars among
the shoot and root components. The September labeling with subsequent
sampling September 20, 1990 and November 4, 1990 show the basipetal
translocation of C assimilated in September during autumn for this
hybrid poplar clone. Standard error of the mean shown as line bars.

 

 

 

 

 

 

 

 

leaves branches stem root root
< 1 mm > 1 mm

November 4, 1990

 

 

 

 

 

 

 

leaves branches stem root root

138

Figure 5.3a. The distribution of total starch and labeled 1‘lC-starch among
the shoot and root components. The July labeling and subsequent
sampling August 2, 1990 and November 2, 1990 show the tendency of
this clone to translocate C to the root system for storage in July during
mid-growing season. The distribution of labeled starch increases in the
root system as dormancy is induced. Standard error of the mean shown
as line bars.

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

Figure 5.33
August 2, 1990
8)
70
I Stﬂl‘Ch 14C-starch I
(i)
50
a“ 40
leaves branches stem root root
< 1 m > 1 mm
November 2, 1990
so
70 s 14 I
I starch % C-starch
a)
50
8 40

 

 

 

 

 

 

leaves branches stem root root
1

138

Figure 5.3a. The distribution of total starch and labeled 14=C-starch among
the shoot and root components. The July labeling and subsequent
sampling August 2, 1990 and November 2, 1990 show the tendency of
this clone to translocate C to the root system for storage in July during
mid-growing season. The distribution of labeled starch increases in the
root system as dormancy is induced. Standard error of the mean shown
as line bars.

 

 

 

 

 

 

 

 

 

 

 

 

 

 

Figure 5.3a
August 2, 1990
so
70
I starch 14C-starch
(i)
50
6‘ 40
[MM
sequtrl
161301 “ leaves branches stem root root
rdllﬂﬂl < 1 m > 1 mm
sinl!
ishm
November 2, 1990
8° I
70
I starch E 14C-starch
5Q

 

 

 

 

 

 

leaves branches stem root root
< 1 m > 1 mm

140

Figure 5.3b. The distribution of total starch and labeled 14C-starch among
the shoot and root components for trees labeled September 5, 1990. The
subsequent sampling done on September 20, 1990 and November 4, 1990
show the basipetal translocation and loading of the coarse root system
with starch and labeled C assimilated during autumn. Standard error
of the mean shown as line bars.

arch am
i, 1990. h
nber 4, 195“.
root systl
mdm‘d em

leaves

leaves

141

Sepember 20, 1990

branches

stern root
< linrn
November 4, 1990

branches

stern

root
< ltnni

 

root
> 11nni

 

root
> linnn

142

Approximately 60-80% of the residue was located in the woody shoot
(Figure 5.4a-5.4 b). Root residue increased through the dormant season.
Labeled C assimilated in July had a similar distribution to residue
distribution with leaves containing proportionately more and roots less.
The trend continued through November with the exception that the stem
accumulates pr0portionately more 14C and the leaf litter contained less.
Carbon assimilated in July exhibits acropetal movement and stabilization
in above-ground tissue. The photosynthate assimilated in September was
distributed mainly to the stem and root system. The increase in 14C in root
residue in September and November (Figure 5.4b) clearly demonstrated
active ﬁne root growth after new leaf expansion has stopped. Branches
retained proportionately less 14C while leaves and litter retained an
increased level. The distribution of 14C within residue changes little from
September to November (Figure 5.4b).

Figure 5.5a-5.5b represents the distribution of 14'0 in the different
chemical fractions expressed as a percentage of recovered from the total
tree. The residue pool always contained the largest amount of 14C
compared to other fractions. In the trees labeled in July, the labile fractions
contained no greater than 20% of the recovered 14C, except in litter. The
accumulation of 14C in starch remained constant in roots (> 1 mm) (Figure
5.5a). In September the labile fraction of roots (> 1 mm) and branches
contained up to 40% of the recovered 14C (Figure 5.5b). The accumulation of
14C in the labile fraction remained constant in branches and roots (> 1 mm)
in November from the September labeling (Figure 5.5b). A signiﬁcant
amount of 1‘er accumulated in the chloroform soluble fraction of branches
and stem from the September labeled trees. The recovery of 14C in root

starch remained constant from September to November. The distribution

143

Figure 5.4a. The distribution of total residue and labeled 1‘ltC-residue
among the shoot and root components of trees labeled July 19, 1990. The
subsequent sampling done on August 2, 1990 and November 2, 1990 show
that the shoot residue comprises a signiﬁcant proportion of the tree.
Standard error of the mean shown as line bars.

Figure 5.4a

a?
I-residie
990.
99051101
the tr!
e

144

 

 

 

 

 

leaves branches

 

 

 

stem root

 

 

 

I residue

 

1 4C—residue

 

 

 

 

leaves branches

 

 

stem root

 

145

Figure 5.4b. The distribution of total residue and labeled 1‘lC-residue
among the shoot and root components of trees labeled September 5, 1990.
The subsequent sampling done on September 20, 1990 and November 4,
1990 show that the root residue is both increasing in mass and receiving
more labeled C at this time during the season. Standard error of the
mean shown as line bars.

146

Figure 5.4b

September 20, 1990

 

 

I residue 14C-residue

 

50

 

 

 

 

 

leaves branches stem root root

November 4, 1990

 

I residue 14C-residue
50

 

 

 

 

 

 

147

Figure 5.5a. The distribution of 140 within the chemical fractions of the
trees labeled July 19,1990. The subsequent samplings done on August 2,
1990 and November 2, 1990 reveal that trees labeled during the growing
season synthesize a signiﬁcant amount of the residue fraction as a
result of shoot and root expansion.

 

 

 

 

 

 

149

Figure 5.5b. The distribution of 14C within the chemical fractions of the
trees labeled September 5,1990. The subsequent samplings done on
September 19, 1990 and November 4, 1990 show that trees labeled after

budset synthesize less of the residue fraction and allocate more
photosynthates to labile fractions.

Figure 5.5b

 

_/z

 

 

 

 

    

 

_| _____

 

of

58

151

of 1"*C in the residue pool changed little regardless of date labeled or
sampled (Figure 5.5a-5.b).

Fate of Reserves

Figure 5.6a and b show the speciﬁc activity (Bq mg'1 C) of
nonstructural carbohydrates and the residue fraction of ﬁne roots (< 0.5
mm) one year following the radiolabeling procedures. The speciﬁc activity
of the residue fraction changed little over the year period despite a. slight
increase in the mass of ﬁne roots from November. The 14C activity of the
nonstructural carbohydrates generally declined during the year from the
utilization and accumulation of new reserves. This implied that the
synthesis of ﬁne roots was derived from labeled reserves. Since the reserves
had twice the speciﬁc activity of the residue in November, it was possible
that the addition of unlabeled photosynthate sustained the activity of the ﬁne
root residue at previous year levels. This implied ﬁne root growth depended
on signiﬁcant stored photosynthate accumulated during the previous year.
Since the ﬁne roots themselves do not contain sufﬁcient reserves to increase
their mass, movement of reserves from roots (> 0.5 mm) and possibly the
above-ground tissue must have occurred. The amount of starch reserves in
the root system in November was greater than the increase in mass of ﬁne
root pool the following season indicating this scenario may be realistic.

Considering the total reserves contained within the tree, the constant

speciﬁc activity of the ﬁne roots after one year seems plausible.

 

152

Figure 5.6a. The amount of ﬁne roots and the total nonstructural
carbohydrates (sugars and starch) in the root system for the July
labeling and subsequent harvests (upper graph). The speciﬁc activities
of sugar, starch and residue fraction of ﬁne roots (< 0.5 mm) for the year
following labeling (lower graph). Standard error of the mean shown as
line bars.

July

Figure 5.6a

roots < 0.5 m (g)

______
______

 

.n gum.
mmmmiw
cccc

 

 

154

Figure 5.6b. The amount of ﬁne roots and the total nonstructural
carbohydrates (sugars and starch) in the root system for the September
labeling and subsequent harvests (upper graph). The speciﬁc act1v1t1es
of sugar, starch and residue fraction of ﬁne roots (< 0.5 mm) for the year
following labeling (lower graph). Standard error of the mean shown as
line bars.

Figure 5.6b

roots < 0.5 m (g)

sugar (g)

I starch (g)

_ _ _ _
_ _ _ _ _ _ J

 

 

 

156
DISCUSSION

This research showed the similarity between 120 and 1‘er
assimilation during normal tree growth. The amount of storage reserves,
including sugar, starch and protein, increased from July to November in
most tissue, especially in coarse roots. The amount of radiolabeled
photosynthate in roots increased and surpassed the concentration in woody
shoot tissue, verifying that the below-ground tissue had become the
dominant sink following budset. In the July labeling, the amount of 140 in
carbohydrate and protein pools typically ranged between 1-2%. The sugar
and protein pools were comprised of less 14C than the starch pool showing
that these pools were being diluted with unlabeled C. In contrast, the root
sugar pool in September remained more than twice as enriched as the
starch pool indicating that the accumulation of sugar was occurring at a
rate exceeding its conversion to starch. Regardless of when the 14C
labeling occurred, the protein and carbohydrate pools retained labeled
photosynthate into dormancy.

Few studies have followed the fate of 14C-photosynthates into labile
and insoluble pools in tree tissue on a seasonal basis and most do not
represent ﬁeld data. The results of laboratory and glasshouse studies in
conifers (Schier, 1970; Glerum and Balatinecz, 1979) and hardwoods
(Kandiah, 1979; McLaughlin et al., 1980) indicate that the fate of
assimilated C from earlier in the growing season is dynamic, never
achieving equilibrium during the season of assimilation. Our data

conﬁrms that the 14C is located in both labile and structural pools of ﬁne

roots in the following growing season.

 

S}:
the
sin];

157

The storage of carbohydrates in the root system begins before
diameter growth of the coarse roots (Dickson, 1989). Diameter growth of
roots follows that of the shoot growth, generally as a wave of cambial
activity from developing buds (Kramer and Kozlowski, 1979). Deposits of
starch in the ray and parenchyma cells of expanding roots begins in mid
summer (Wargo, 1976) and is illustrated by our data (Tables 5.1-5.4). This
early starch storage is not hydrolyzed to support cambial activity (Dickson,
1989). This infers that current photosynthate is used to support growth and
respiratory activity of the root system. The ﬁne root pool may also act as
storage for carbohydrates (Nguyen et al., 1990). The increase in
carbohydrate storage, especially starch, in ﬁne roots is evident both in
increasing concentration and 14C activity. The results in Figures 5.2-5.4
indicate the importance of storage by exemplifying the correlation between
the distribution of 140, storage pools and structural residue C. These
results are in contrast to the Marshel and Waring (1985) hypotheses which
predict that starch deposition occurs during the initiation of ﬁne (roots < 2
mm) in Douglas ﬁr (Pseudotsuga menziesii). van den Dressche (1987),
using Douglas ﬁr seedlings, found that current photosynthate was
important to new root growth, but also found root growth dependent on the
supply of storage reserves. Clearly, the relative importance of current and
storage photosynthate needs further research to adequately deﬁne the
potential for and limits to growth (Glerum, 1980; Dickson, 1989).

The storage of C is also accompanied by the temporary storage of N
which is necessary for the perennial growth of trees. The uptake,
synthesis, distribution and recycling of nitrogen compounds is similar to
the fate of photosynthates in that allocation is dependent upon competing

sinks. As the growing season passes, the N requirements of the shoot

158

diminishes when the synthesis of photosynthetic enzymes ceases (Titus and
Kang, 1982). During leaf senescence, some leaf protein is hydrolyzed to
amino acids and transported back into the woody tissue of the shoot and
roots. An increase in the concentration of root protein was distinguishable
in November (Tables 5.1-5.2). The salvage of leaf protein nitrogen will
depend on a number of factors, including the level of tree N, biotic
inﬂuences such as infectious leaf diseases, and abiotic factors such as frost
damage. The percentage of N retranslocated from leaves of Populus is
generally between 60-80% (Baker and Blackmon, 1977). The present data
indicate that from 6-25% of protein N was salvaged from leaves based on
concentration. The low translocation of leaf N may be attributable to the
nitrogen-rich site which was formerly under alfalfa cultivation and may
explain the lack protein decrease in leaf litter.

Storage of N in trees can be either in the form of amino acids or
proteins. The storage of amino acids increases in response to N
fertilization in Malus (Tromp, 1983) and in Pinus slyvestris (Nasholm and
Ericsson, 1990). Sauter et al. (1988) revealed the presence of protein bodies
in ray cells of Populus during the dormant season. The occurrence of these
protein storing vacuoles has subsequently been determined in other
deciduous and conifer species (Sauter et al., 1989). The characterization of
these proteins has shown them to be high in arginine and other basic
amino acids (Titus and Kang, 1982; Tromp, 1983). Our data reveals this
trend showing an increase in protein content of the woody shoot and roots
during dormancy. The occurrence of storage protein in trees is a subject

receiving little attention and more research is needed to understand the

accumulation and utilization in response to fertilization and stress

(Dickson, 1989).

 

159

Trees have been classiﬁed according to the predominant form of
storage during dormancy as "fat" or "starch" trees. Ziegler (1964) noted
that many conifers and diffuse porous trees were "fat" trees and many ring
porous trees were "starch" trees. This distinction applies mainly to the
stem and not the root system, which generally stores starch regardless of
species. The extraction of plant tissue with organic solvents removes
considerable material besides fat. The crude extraction represents less
than 10% of the tissue (Krammer and Kozlowski, 1979). Our values for
methanol/chloroform extracts range from 5% in roots to 13.7% in leaf litter
tissue. Chung and Barnes (1977) using a benzene/ethyl alcohol extraction
of needle and axes of Pinus taeda found that the tissue contained 4.7% lipids
and 15-20% phenolics. An extraction of Quercus alba tissue, including
roots, determined that chloroform soluble material composed 1.3-8.7% of
the dry weight (McLaughlin et al., 1980). Nelson and Dickson (1981) found
that stems of Populus deltoides contained 1-3% triglycerides (components of
lipids) in dormancy induced one-year-old plants. These literature values
are difficult to compare since no standard methodology exists for cross-
comparative purposes.

Roots (< 10 mm) contained signiﬁcantly more chloroform extractable
material in September (Table 5.2). At this time the roots receive more
photosynthate due to the basipetal ﬂow of C. The transported sugars were
undergoing conversion to starch for long term storage. Sauter (1988) has
documented the formation of vesicular and dilated ER-cisternae during
sugar-starch interconversions in Populus stems stored at 0°C. The
formation of this vesicular material would require the accumulation of
membrane lipids. During dormancy induction, sugars are accumulating

and undergoing conversion into starch and may explain the increase in the

1H)

chloroform-soluble fraction in roots labeled in September. The
accumulation of lipids in response to root diameter growth and for the
synthesis of storage cells are additional reasons for lipid accumulation in
roots.

The distribution of recovered 14’0 in the chloroform-soluble fraction
reaches a maximum of 27% in branch tissue in November from the
September labeling. This agrees with Ziegler's (1964) interpretation of a
diffuse porous being a fat storing tree. However, roots (> 10 mm) have an
equivalent amount of chloroform soluble material. Perhaps, since the
lipids in these pools were not deﬁned, the comparison of these fractions is
inappropriate. Therefore, the comparison to other research may also have
no meaning. However, since this class of compounds are susceptible to
nonpolar organic solvents the comparison is noteworthy. Glerum (1980)
found recoveries of 14C in the lipid pool of Pinus banksina leaves of greater
than 35%, in roots 8-18% and woody shoot tissue displaying intermediate
values. The distribution of 14'0 in ether soluble material of Pinus resinosa
trees was generally between 2.1-7.4% in the woody tissue and 16% in
needles (Schier, 1970). The lipid fraction contains both signiﬁcant mass
and photosynthate, but is rarely studied in terms of a storage component in
trees.

The residue component is a complex mixture of polysaccharides and
lignin. This component is structural in nature and its synthesis is related
to the initiation and expansion of shoot and root tissue. The amount of 14C
recovered in the residue fraction decreases as the growing season passes.
In September, the relationship between the allocation of 120 and 14C to the
mass of shoot residue does not correlate as it did in July. The increase in

the amount of 14C distributed to roots (> 1 mm) is probably the result of

161

diameter growth. The diameter growth of the root system is typical of this
clone in autumn and also provides tissue for the storage of photosynthate
(Pregitzer et al., 1990). The retention of leaves in hybrid poplar after budset
in autumn is an important factor contributing to stem and root growth as
well as the formation of reserves (Nelson et al., 1982; Michael et al., 1988;
Pregitzer et al., 1990).

Glerum and Balatinecz (1979) described a similar distribution of 140
in the residue component of Pinus banksina seedlings from July to early
September. However, as the season progressed into November, less than
10% of the 14C was recovered in residue, with the majority now present as
sugars and starch. Dickson and Nelson (1982) also found that the amount
of 14C recovered from the residue declined in 1-year-old Populus deltoides
stems as dormancy was induced under short day length regimes in growth
chamber conditions. These researchers found between 20-30% of the 14C in
the residue fraction of budset plants 48 hours after exposing the them to
14002. The lower recovery of 14C in the residue fraction of Populus
deltoides as compared to our results is probably due to sampling 48 hours
after 140 exposure. Our sampling was done 336 hours after 14C exposure
allowing more time for equilibration and incorporation of label into the
residue fraction.

The incorporation of photosynthate into the residue fraction may
involve both synthesis of structural and storage components. It has been
suggested that hemicellulose is a form of storage photosynthate (Glerum,
1980; Bonicel et al., 1987 ). Meyer and Splittstoesser (1971) found
hemicellulose to be the primary form of storage in seedlings of Taxus. This
could explain the high incorporation of 14C into residue in September after

budset, especially in leaf tissue. After budset, no initiation or expansion of

162

the leaf canopy occurs. Yet, a surprising amount of 14C is recovered from
the residue fraction of leaves (Figure 5.4). However, hemicellulose is a
complex polysaccharide and has strong intermolecular bonds with
lignocellulose leading physiologists to believe these carbohydrates are
structural in origin (Krammer and Kozlowski, 1979). More likely, is the
retention of C in leaves in response to wounding inﬂicted by the various leaf
diseases. The higher retention of 1"rC-photosynthate in ozone damaged
leaves is an example of such a phenomenon (McLaughlin et al., 1982). The
increase of 14C in the residue of roots may also be associated with the
increase of recalcitrant polysaccharides, such as chitin, from mycorrhizal
symbiosis.

The importance of storage photosynthate is exempliﬁed in the lack of
decline in the speciﬁc activity of ﬁne roots (< 0.5 mm) one year following the
exposure to 14002 in both July and September of the previous year. The
reserves accumulated in mid growing season (July label) and autumn
(September label) played an active role in the initiation of new ﬁne roots the
following season. Other researchers have shown the importance of
autumnal photosynthate storage in the growth of new shoots of apple trees
(Quinlan, 1969; Kandiah, 1979). Lockwood and Sparks (1978) found that 78%
of the 14C in roots in November was present in roots after shoot elongation
had ceased in pecan (Carya illinoensis). Similarly, Hansen (1967) found
69% of the 14C present in apple roots in September to November of the
following year. The same or increased recovery of 14C was found in roots of
Pinus resinosa labeled in July and October up to 10 months after exposure
(Schier, 1970). Generally, these studies have observed C storage in
relationship to shoot and fruiting dynamics, neglecting their role in the

maintenance and growth of roots.

163

The speciﬁc activity of the reserves, sugar and starch, in the ﬁne
roots was approximately twice that of the residue pool in November
regardless of when 14C02 exposure occurred (Figures 5.6a-b). The speciﬁc
activity of these storage pools increased with increasing root diameter.
Therefore, the utilization of these pools to initiate new ﬁne root growth
required that they be diluted with an equivalent amount of unlabeled C to
maintain the speciﬁc activity of the ﬁne root pool from the previous year.
The source of the unlabeled C is current assimilates and indicates that the
growth of ﬁne roots is dependent on reserve and current photosynthate.
The rate of reserve utilization and its subsequent dilution in the growth of
ﬁne roots was not distinguishable since the root system was composited
when harvested. It is possible that depending on demography and ontogeny
of ﬁne roots the utilization of reserve material was not uniform. Any
growth of roots before shoot elongation would have been entirely dependent
on reserves and would produce roots of the same speciﬁc activity as the
reserve material. It is evident from our data that growth of ﬁne roots is
dependent on reserves and current photosynthate. The speciﬁc activity of
reserves remaining are beginning to equilibrate with structural residues

one year after exposure to 14C.

CONCLUSION

This research has presented evidence that the synthesis and
turnover of the root system is inﬂuenced by tree activities occurring over
more than one season. During the growing season, current assimilates
are utilized for respiration, growth and storage of reserve material from a

whole-tree perspective. In autumn, the shoot stops expanding and

164

assimilates are both stored and translocated at an increasing rate to the
root system. The dormant period is characterized by the consumption of
reserve material to maintain life and some growth of the root system before
shoot expansion the following growing season. During the next growing
season, reserve and current photosynthate are used in shoot and root
system expansion. The persistence of reserve material and undeﬁnable
dilution of cell wall tissue in the ﬁne root system exempliﬁes the
importance of storage reserves in these hybrid poplars.

In conclusion, the phenological plasticity of the shoot and root system
will be directly inﬂuenced by the amount and utilization of reserve material
stored in current and subsequent growing seasons. The rate of
accumulation of reserve material will be highly sensitive to factors such as
competition, abiotic interactions and cultural practices. It is evident that
the success of these hybrids in producing increased biomass at an
accelerated rate will depend on the ability to sequester reserves in more
than one growing season. The storage and utilization of reserves, is
therefore, an extremely important process that inﬂuences the success of
this and other species in managed and unmanaged ecosystems.
Understanding patterns of mobilization and utilization of reserve material
is necessary to fully understand the consequences of cultural practices in

ecosystems designed with decreasing diversity and increasing production.

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the Stem of Populus x canadensis robusta. Plant Physiol. 132:608-612.

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Biochemical Results on the Localization and Distribution of Storage
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1%

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Recycling in Apple Trees. Horticultural reviews. 4: 204-246.

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Chapter 6

MICROBIAL BIOMASS AND DYNAMICS OF C AND N
IN POPUL US PLANTATIONS

ABSTRACT

Soil microbial activity and organic matter dynamics are central to
carbon cycles and nutrient ﬂuxes in ecosystems. The contribution of leaf
and root production and turnover to the soil microbial biomass and soil
organic matter (SOM) maintenance are vaguely understood. This study
was designed to determine the role of leaf and root turnover in the
dynamics of carbon and nitrogen cycling through the microbial biomass
and SOM. Two-year-old hybrid poplars were labeled with 14C and 15N in
the ﬁeld at different times of the growing season to characterize the ﬂux of
C and N in the tree-soil system. Labeled leaf litter was placed on unlabeled
plots to determine the relative contribution of leaf and root turnover.
Microbial biomass methodologies of direct microscopy, chloroform-
fumigation incubation (CFI) and chloroform fumigation-extraction (CFE)
were used to measure microbial biomass. The comparisons of the
microbial biomass methods showed that the techniques of CFI and CFE
correlated well against direct microscopy, but gave consistently higher
values. The subtraction of a partial control from CFI gave values similar to
direct microscopy in surface and subsurface soils. The comparison of the
14C activity of microbial biomass C from CFI and CFE indicated these
methods may be assaying different pools of microbial C. Long-term

incubation of soil the year following labeling was used to study the

170

171

dynamics of C and N mineralization. Mathematical analysis of the
mineralization curves provided decomposition rates, kinetic parameters
and pool sizes of labile and more stable soil C pools. Kinetic analysis of
long-term soil incubations indicated a greater contribution of C and N to
soil organic fractions from leaf litter decomposition in the early stages of
the rotation. The mean residence time of 14C in surface and subsurface
soil ﬁom leaf and root turnover was 14-64 days for labile organic soil C and

2-16 years for stabilized C.

172
INTRODUCTION

Soil organic matter dynamics (SOM) are intrinsically related to
carbon (C) cycles and nutrient ﬂuxes in managed and unmanaged
ecosystems. These in turn are inﬂuenced by cultural-management
practices, the magnitude, quality and timing of residue inputs as well as
soil physical and chemical characteristics (McGill and Cole, 1981; Paul,
1984). Various components of SOM act as temporary energy and nutrient
reservoirs. The determination of the pool size and the rates of release of
nutrients from these pools is important in determining the nutrient cycles
of ecosystems and the fertilizer requirements of managed systems.

Attempts to utilize a chemical or physical extraction technique to
predict the amount of N that would be released during a crop's growth have
not been very successful (Keeney, 1982; Juma and Paul, 1984). The
exception can be found in dry climates where soils accumulate nitrates due
to the lack of leaching over autumn, spring, or fallow periods. The SOM
pool that provides nutrients required for plant growth over a reasonably
short term is usually referred to as the "active fractions". The size and
kinetics of active fractions can be determined quantitatively with tracer
analysis (15N, Jansson, 1958; 14C, Coleman and Fry, 1992). These
components are thought to be comprised primarily of partially decomposed
plant residues, microbial biomass, and recent microbial byproducts. The
size of these fractions have been determined by long-term incubation
combined with mathematical analysis of the accumulated inorganic C and
N (Standford and Smith, 1972; Boyle and Paul, 1989; Blet-Charaudeau et al.,
1990).

Ann—.—

173

The measure of nitrogen (N) availability and cycling provides a
tangible determination of the sustainability and production potential of
ecosystems. Nitrogen availability controls the amount and substrate
chemistry of plant litter (Aber and Melillo, 1982; Vitousek, 1982;
McClaugherty et al., 1985). Many methods of assessing N availability via
incubation have been developed to determine fertilizer need and understand
productivity at the stand level. These include: soil incubation at increased
temperature (37°C) and periodic leaching (Standford and Smith, 1972);
statically incubated soils with sampling to determine N accumulation
(Vitousek et al., 1982); buried polyethylene bags (Zak and Pregitzer, 1990);
soil cores encased in ion exchange resin (Di Stefano and Gohlz, 1986); ion
exchange resin in soil (Binkley and Matson, 1983); and anaerobic soil
incubation (Powers, 1980). These methods describe differences in N
availability between sites, but fare poorly when the methods are compared
indicating that N availability is being qualitatively assessed (Hart and
Firestone, 1989; Powers, 1990).

A number of studies have shown that the size of the microbial
biomass itself appears to be well correlated with the size of the active
fractions (Anderson and Domsch, 1989; Smith and Paul, 1990). In addition,
the net magnitude of microbial mediated processes involved in the
immobilization-mineralization reactions are reasonably well understood
(Paul and Clark, 1989). The gross magnitude of nutrient ﬂux is not well
characterized. There have been a number of studies showing that the use
of tracers combined with long-term incubations can be used to determine
the ﬂuxes involved in N mineralization, nitriﬁcation and immobilization of

N (Robertson and Vitousek, 1981; Vitousek et al., 1982).

"-.a

174

Nutrient dynamics in forests are difﬁcult to study due to different
growth rates of the plant components and their large size, making the use
and interpretation of tracer research difﬁcult. This is complicated further
by fertilization effects caused from increases in carbon dioxide and
deposition of atmospheric N and other nutrients (Kauppi et al., 1992). The
recent interest in short-rotation forestry, such as Christmas tree farming
and production of biomass for pulp and energy industries has led to a
requirement for information on the N needs of these systems. Short-
rotation forestry provides an amenable system to study (Horwath et al.,
1992). A number of reasonably sized, uniform and non-competing trees can
be labeled with 140-15N , (Chapter 3; Smith and Paul, 1988).

In the present study, I examined the role of root and leaf litter C and
N in the production of microbial biomass and active organic matter pools.
Long-term soil incubations measured the occurrence and stabilization of
14C and 15N in soil organic fractions derived from the labeled tree litters.
The tracer content of the microbial biomass and SOM fractions was
measured by analytical assays. Kinetic analysis of long-term soil
incubation data provided information on pool sizes and ﬂuxes. The
uncertainty associated with microbial biomass assays has led to the
development of a variety of techniques to determine microbial C, N and
tracer content. To establish which assay was suitable to determine C and N
content of the soil microbial biomass I compared the results of direct
microscope counts, chloroform fumigation incubation (CFI) and
chloroform fumigation extraction (CFE). This comparison is followed by a
discussion of the results of long-term soil incubation data and kinetics of C

mineralization.

175
MATERIAL AND METHODS

Plant 14C and 15N Labeling

A hybrid poplar plantation (Populus americana cv. Eugenei) was
established in conjunction with the LTER studies at the Kellogg Biological
Station, Michigan State University, Hickory Corners, Michigan. Site,
plantation characteristics and 15N and 14C labeling procedures are
described in Chapter 3. Table 6.1 describes some of the important plant and
soil characteristics of this system. Two sets of eight trees labeled during the
1990 growing season were sampled the following year. July labeled trees
were sampled August 2, 1990 (day 14), November 2, 1990 ( day 107) and July
25, 1991 (day 372). September labeled trees were sampled September 19, 1990
(day 14), November 4, 1990 (day 60) and September 5, 1991 (day 366). Labeled
leaf litter collected during autumn was placed over plots containing
unlabeled trees on December 21, 1990 (Chapter 4). The litter exchange was
done to compare the dynamics of 140 and 15N in soil and microbial biomass
from leaf and root derived C and N. The leaf litter exchange plots were
sampled during the following year on June 6 (day 168) and November 14,
1991 (day 328).

The soil sampling and preparation has been described previously
(Chapter 3). The root labeled soil was sampled to a depth of 100 cm at
intervals of 0-25 cm, 25-60 cm and 60-100 cm. The plots containing labeled
leaf litter were sampled to a depth of 25 cm at 0-10 cm and 10-25 cm
intervals. Soil moisture was determined gravimetrically after drying at
105°C for 24 hours. Soil 14C activity was determined using a
sulfuric/phosphoric acid digestion in modiﬁed culture tubes (Smith et al.,
1989).

 

176

Table 6.1. Characteristics of the plant-soil system studied.

 

Wopulus americana cv. Eugenei
Spacing 2 m x 1 m

Management Short-rotation Intensive-culture

Soil type Kalamazoo ﬁne, loamy, mixed, mesic,

Typic Hapudalf

 

Soil depth
Mots % C % N
0-25 cm 1.06 0.080
25-60 cm 0.37 0.033
60-100 cm 0.27 0.025
lanthanum
0-10 cm 1.10 0.101

10-25 cm 1.01 0.092

 

177

Microbial C and N Determinations

Microbial biomass determinations were conducted three times
during each long-term incubation using CFI ( Horwath and Paul, 1992).
Microbial C determinations by CFI were compared to direct microscopy and
CFE on the ﬁrst two samplings of the labeled tree plots and ﬁrst sampling of
the litter exchange plots. Microbial C determined by CFI was calculated
without the use of a control, Kc=0.41 (V oroney and Paul 1984). The Kc for
the CFE determination was 0.35 (V oroney et al., 1992). Microbial N was
calculated based on the ratio of the fumigated ﬂush of C to N, Kn (V oroney
and Paul, 1984). CFI data were also recalculated using a partial control as
determined by Horton, Horwath and Paul (Personal communication).

Direct microscopic counts of acridine-orange-stained bacteria and
fungi were done on the ﬁrst two samplings of the July and September
labeled trees. Soil (10 g) was homogenized with 200 mL of 0.2 M Tris (pH 8)
in a Waring blender for 60 seconds. Serial dilutions of 1 mL homogenate to
9 mL Tris buffer were done to obtain 0.1 mg soil mL'1 for bacteria and 0.5
mg soil mL'1 for fungi. The ﬁnal dilutions were stained with 0.5 mL
acridine orange (0.1% w/v in water) for 10 minutes (Faegri et al., 1977). The
dilutions were preserved by adding 0.1 mL formaldehyde. The entire ﬁnal
dilution was drawn through a pre-wetted (1 mL 0.1% tween in water) 0.2
pm black Nucleopore membrane ﬁlter on a vacuum side arm ﬂask. The
ﬁlter was mounted on a microscope slide with a drop of immersion oil and
cover slip. Graticule ﬁelds were counted randomly until at least 300
bacteria were counted per slide. Fungal hyphae length was determined
with a line intersect method using an ocular grid (Hanssen et al., 1974). A
mean volume of 0.12 m3 for bacteria (Bakken and Olsen, 1983) and a 2.3

um diameter for fungi (Dave Harris, personal communication) were used

_J.—.—'_'

178

in the calculation of microbial biomass. Carbon content of bacteria and
fungi was calculated using values for speciﬁc gravity of (1.1), solids content
of (0.3) and 45% C (Paul and Clark, 1989).

Additionally, microbial biomass C was determined by the CFE
method (Vance et al., 1987). Fumigated and unfumigated soils (25 g wet
weight) were extracted with 0.5 M K2SO4 (5:1 v/v) for 1 hour. Soluble C was
determined on extracts using a modiﬁed persulfate digestion (Horwath and
Paul, 1992). The soil extract (15 mL) was digested with 1 g persulfate and 1
mL 0.05 M H2804 in a modiﬁed culture tube containing a vial with 1 mL 0.1
M NaOH. The culture tubes were sealed with Poly seal caps and digested at
120°C for 2 hours. The contents of the vial were titrated to a phenolthalien
endpoint with 0.1 M HCl after the addition of an equivalent amount of BaC12
to determine total C. The contents of the titrated base traps were rinsed into
scintillation vials, 10 mL of scintillation cocktail added and 14C activity
determined on a liquid scintillation spectrometer (Packard Instrument Co.,

Downers Grove, IL).

Long-Term Soil Incubations

The soil was weighed (60 g ﬁeld moist, approximately 52 g dry
weight) into urine specimen cups and placed into Mason jars for long-term
COz mineralization studies. A vial containing 5 mL of 0.6 M NaOH was
placed with the soil and the Mason jar sealed. The soil was incubated at
25°C for apprmdmately 150 days. The vials were changed every 15-20 days
with less frequent sampling as the incubations proceeded past 75 days. The
contents of the vials were divided in two to determine mineralized 002-0
and 14C activity. Mineralized 002 was determined by titrating the NaOH
with 1.5 M HCl to a phenolthalien endpoint (pH 7) after the addition of an

179
equivalent amount of BaClz (Horwath and Paul, 1992). The remainder of
the vial contents was combined with 10 mL of scintillation cocktail
(Research Products International Corp., Mount Prospect, IL.), sufficient
water to gel the mixture and the 14C activity determined.

Soil (350 g, oven dry weight) for long-term N mineralization and
microbial biomass determinations was placed into Mason jars with 1 mm
diameter holes punched into the lids. Soil moisture was maintained
gravimetrically during the incubation period. Mineralized N was
determined throughout the long-term incubation at approximately 0, 10, 20,
40, 50, 60, 70, 90, 120 or 150 days. Soil (25 g) was removed from the Mason
jars and extracted with 2 N KCl (5:1, v/v) for 0.5 hour. Inorganic N (NO3'
and NH4+) was determined colorimetrically using a ﬂow-injection
autoanalyzer (Lachet Chemicals Co., Mequin, WI). The diffusion
procedure (Brooks et al., 1989) was used to determine the 15N content of
inorganic soil N and microbial biomass N. The 15N analysis was done on a

Gas chromatograph-Mass Spectrometer (Europa Scientiﬁc, Crewe,

England).

Calculations

Standard deviation and error of the mean are shown to demonstrate
the reproducibility of ﬁeld and laboratory techniques. Potential C
mineralization (C1) was calculated using a mixed order and modiﬁed
double exponential equations. The mixed order equation (1) incorporates a
zero and ﬁrst order component describing the cumulative C mineralization

curves from the long-term soil incubations (Blet-Charaudeau et al. , 1990):

Ct: C1*(1-exp(-k1*t) + k2*t (1)

180

where Ct is the total amount of COz-C evolved at time t; k1, kg and Cl are
constants expressing, respectively, the ﬁrst order rate constant, the zero
order rate constant and potentially mineralizable C. A modiﬁed double
exponential equation (2) was adapted from (Bonde and Rosswall, 1987):

Ct=C 1*(1-6XP(-kl*t))+((Csoil/ 2)-C 1)*(1-exP(-k2*t)) (2)

where Csoil is the total organic C in soil. In equation 2, the above assumes
three soil C pools Cl, 02 and C3. The potentially mineralizable Cl pool is
described by the ﬁrst order equation. 03 was estimated from carbon dating
as being 50% of the total soil C (Csoil/Z) with a turnover time sufﬁciently
long it does not affect the amount of C02 evolved during the long-term
incubations (Martel and Paul, 1974), therefore, the more stable soil C pool is
deﬁned as C2=Csoi1/2-Cl. The parameters were developed for the
cumulative C mineralization curves using a nonlinear model estimation
procedure with Quasi-Newton minimization methods (Systat: Statistics,
Evanston, IL). The software computes mean square errors, asymptotic
standard errors and correlation by estimating the Hessian (second

derivative) matrix.

RESULTS

Microbial Biomass
Direct microscopy is a traditional method to determine soil microbial
biomass C and N and is used as standard to calibrate other methods

(Horwath and Paul, 1992). The advantage of direct microscopy to determine

 

181

microbial biomass is the ability to differentiate fungal and bacterial
populations. Fungal hyphal lengths were 109 to 146 m g'1 (72.9-98.1 llg C g'
1 soil) in the surface soil (0-25 cm) (Table 6.2). They dropped by a factor of
2.9 to 3.7 (26-42 ug C g-l soil) at the 60-100 cm depth. Bacterial numbers
ranged from 1.26-1.6 x 109 cells g-l (22.5-28.7 pg C g1 soil) of surface soil.
The surface to subsurface bacterial number ratio was 4.0 to 8.0, thus the
subsurface soil contained more fungal C than bacterial C indicating the
importance of fungi in subsurface soil. Microscopy, however, does not
measure the turnover of the fungal hyphae.

Soil microbial C determined by CFE for surface soil was 112-147 pg C
g‘1 for the ﬁrst two sampling of tree plots in 1990 (Table 6.3a). The CFI
method gave slightly higher values of 143-219 ug C g'1 soil for the same
sampling and an additional ﬁnal sampling of the tree plots the following
year (Table 6.4a). Direct counts, CFI and CFE all showed similar changes
with time, therefore, the difference in the biomass for the surface soils are
considered to be meaningful measurement of seasonal differences.
Microbial biomass C values were lower in subsurface soil for both
fumigation methods corroborating the similar trend found for direct
microscopy.

The microbial biomass C values for the ﬁrst sampling of the leaf litter
exchange plot were approximately 350 pg C g‘1 in the surface soil (0-10 cm),
as determined by both CFI and CFE methods (Tables 6.3b and 6.4b). The
high values for microbial C subsided for the second sampling of the leaf
litter exchange experiment determined by CFI. The subsurface soil (10-25
cm) gave estimates of microbial C more similar to the surface soil of the

labeled tree plots. The apparent increase in microbial biomass was

182

 

 

 

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183

Table 6.3a. Microbial biomass C determined by the CFE method for the ﬁrst

and second samplings of trees labeled with 14C and 15N July 19,1990 and
September 5,1990. Standard error of the mean shown in parentheses.

   

 

 

 

Uncorrectd Corrected
Soil Extractable biomass C Biomass C Bq mg-l
depth soil C ”g C g-l “g C g-l biomass C
(cm) FEE-1 soil soil
Wm
Sampled August 2, 1990
0—25 50.4 (1.0) 51.5 (4.6) 147.1 (13.2) 1.0 (0.2)
25100 57.1 (3.9) 30.1 (8.2) 86.0 (23.4) 1.1 (0.4)
60-100 37.9 (2.5) 29.5 (4.3) 84.2 (12.4) 0.7 (0.1)
Sampled November 2, 1990
0-25 65.0 (3.6) 56.4 (8.0) 161.2 (23.0) 1.2 (0.1)
25-100 80.2 (4.3) 15.0 (1.1) 42.9 (3.2) 1.5 (0.2)
60-100 49.3 (4.6) 9.2 (3.1) 26.3 (8.8) 1.9 (0.4)
W
Sampled September 20, 1990
0-25 63.2 (1.1) 39.4 (4.2) 112.5 (12.1) 2.0 (0.1)
25-100 60.7 (2.7) 21.7 (4.2) 62.1 (12.1) 1.0 (0.1)
60-100 45.2 (2.5) 11.4 (1.8) 32.6 (5.3) 2.0 (0.6)
Sampled November 4, 1990
0-25 59.9 (2.6) 52.2 (2.4) 149.1 (6.7) 1.7 (0.2)
25-100 74.9 (4.4) 21.4 (4.4) 61.1 (12.6) 1.3 (0.9)

60-100 52.7 (0.7) 11.3 (0.3) 32.4 (0.8) 2.2 (0.3)

 

“E

184

Table 6.3b. Microbial C determined by the CFE method for the ﬁrst

sampling, June 6, 1991, of the litter exchange experiment. Standard
error of the mean shown in parentheses.

 

 

Uncorrected Corrected
Soil Extractable biomass C Biomass C
depth soil C pg C g-l pg C g-l Bq mg'l
(cm) Fig-1 soil soil biomass C
Leaf litter exchange, started
December 21,1990
September litter
0-10 29.7 (2.1) 123.4 (5.3) 352.5 (15.0) 4.1 (0.3)
10-25 28.8 (2.0) 91.6 (4.1) 261.7 (11.8) 0.5 (0.1)
July litter
0-10
10-25 36.6 (1.1) 122.3 (2.8) 349.3 (8.0) 1.8 (0.1)
36.0 (3.2) 80.7 (6.0) 230.6 (17.0) 0.2 (0.0)

 

.w' __ "—411.“

f .

185

Table 6.4a. Microbial biomass C, 14C and N determined by the CFI method
for all samplings of trees labeled on July 19,1990 and September 5,1990.
Standard error of the mean shown in parentheses.

 

Soil Fumigate Corrected Bq mg-l Biomass N C/N
d
depth ﬂush biomass C biomass C ”g g-1
(cm) pg c g-1 ”g g-1 soil
soil soil
W
Sampled August 2, 1990

0-25 58.6 (1.9) 143.0 (4.5) 1.9 (0.1) 25.7 (0.7) 5.6
25-60 29.1 (9.6) 70.9 (23.3) 1.1 (0.3) 10.7 (3.2) 6.6
60-100 20.7 (5.7) 50.6 (13.9) 0.8 (0.4) 7.5 (1.8) 6.8

 

Sampled November 2, 1990

0-25 76.3 (3.2) 186.1 (7.7) 1.1 (0.2) 32.1 (1.2) 5.8
25-60 11.1(3.8) 27.2 (9.3) 2.0 (1.0) 4.4 (1.4) 6.2
60-100 3.1 (3.2) 7.6 (7.7) 4.2 (2.5) 1.4 (1.3) 5.5

Sampled July 25, 1990

0-25 87.5 (4.6) 213.3 (11.3) 0.5 (0.1) 34.9 (0.1) 6.1
25-60 26.7 (6.8) 65.0 (16.6) 0.5 (0.1) 9.7 (3.8) 6.7
60-100 10.8 (3.2) 26.4 (7.7) 0.7 (0.2) 4.0 (1.7) 6.6
W
Sampled September 20, 1990

0-25 73.1 (0.6) 178.3 (1.4) 3.2 (0.3) 31.6 (0.4) 5.6
25-60 27.8 (1.4) 67.9 (3.4) 3.8 (0.6) 10.6 (1.0) 6.4
60-100 17.8 (1.2) 43.3 (3.0) 2.9 (0.3) 5.9 (0.5) 7.4

Sampled November 4, 1990

0—25 77.8 (6.6) 189.8 (16.2) 2.5 (0.2) 33.0 (4.4) 5.8

25-60 32.7 (2.6) 79.7 (6.2) 2.2 (0.2) 11.1 (0.6) 7.2

60100 20.6 (1.6) 50.2 (3.9) 2.3 (0.2) 6.9 (0.9) 7.3
Sampled September 5, 1990

0-25 89.8 (2.3) 219.0 (5.7) 0.7 (0.1) 37.6 (1.6) 5.8

25-60 28.7 (0.1) 70.0 (0.4) 0.5 (0.1) 9.6 (0.6) 7.3

60-100 15.5 (1.4) 37.8 (3.5) 0.8 (0.2) 5.8 (1.1) 6.5

 

186

 

 

 

 

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187

perplexing, since an equivalent amount of unlabeled leaf litter was placed
on the labeled tree plots.

The speciﬁc activity of the biomass C is similar at all depths for the
labeled tree plots across time indicating a similar relation between roots
and microorganisms and that the recently incorporated C is behaving
similarly with time at all soil depths (Tables 6.3a and 6.4a). The lower 14C
levels in the subsurface soil is a result of less root biomass. Approximately
50% of both the ﬁne root biomass and microbial biomass was located below
25 cm (Chapter 4). The 14C activity of the microbial biomass from the leaf
litter exchange experiment plots was concentrated in the surface soil (0-10
cm) (Table 6.3b and 6.4b). The higher 14C concentration in the surface soil
of the litter treatment was a fundamental difference between the input of C
to soil from leaf litter (leaf derived C) and root turnover (root derived C).

The microbial biomass of the tree plots contained more 140 activity 14
days after labeling in the September labeling than in the July labeling
(Tables 6.3a, b and 6.4a, b). The higher amount of 14C found in the biomass
during September corresponds to the increase in the translocation of C to
the root system for this Populus clone during autumn (Chapters 4 and 5).
The biomass 14C in the September labeled trees subsided to levels similar to
July labeled trees one year later. The drop in 14:0 activity in microbial C
over time in the tree plots shows the turnover of the microbial biomass and
the utilization of new unlabeled substrate from the soil. The speciﬁc activity
of the ﬁne roots had not dropped signiﬁcantly during this time. This trend
is not exhibited in the leaf litter labeled plots indicating that the labeled C
input is being sustained by the continued movement from the aboveground
leaf litter to the soil beneath. This demonstrates an additional fundamental

difference in the role of C soil input between leaf and root derived C.

188

Microbial biomass N values in surface soil ranged from 26-36 pg N g1
soil in tree plots (Tables 6.4a, b). The elevated amount of microbial N (27-
62.6 pg N g1) found in the ﬁrst sampling of the leaf litter exchange plots is
similar to the microbial C results for these plots. These higher values are
partly explained by dividing the surface soil in to 0-10 and 10-25 cm depth
increments. However, this does not explain all of the increase which could
be attributable to the spatial proximity of the leaf litter to 0-10 cm soil. Table
4a shows that there appears to be a tendency of the microbial C:N ratio to
widen with depth. Since the microscopic counts showed a greater
proportion of fungi with depth and since ﬁmgi have wider C:N ratios, the
C:N ratio as determined by the CFI should widen. This again veriﬁes the
accuracy of the techniques employed.

The comparisons of the fumigation methods to direct microscopy for
microbial biomass are shown in Figures 6.1a-c. The comparison of CFI
and CFE show high correlation across time and soil depth. This indicates
that the different constants and variables used to calculate microbial
biomass are both meaningful and applicable, especially with soil depth.
Similarly, the comparison between CFI and CFE produce a slope of 1 and
R2 of 0.98 (Figure 6.1d). The similarity in microbial C values between the to
fumigation methods is remarkable since the CFE methods subtracts a
control. The biomass 14C activity determined by the two methods are not
statistically different (Figure 6.1e), although CFE had a tendency to be
somewhat lower. This indicates that the two fumigation methods maybe
sampling two somewhat different C pools.

The results of all methods to determine microbial biomass were
normalized to CFI values to facilitate their comparison (Table 6.5). The
data include a computation that subtracts a partial control from the CFI

 

 

189

 

 

 

 

 

 

 

 

 

 

 

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The comparison of the fumigation and direct microscopy

methods of microbial biomass determination are shown in a-c. Figures
d and e compare the biomass C and speciﬁc activity of biomass C from

the fumigation methods.

190

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191

values. The partial control was determined by the addition of an internal
standard to the fumigated and unfumigated soil. The internal standard
(substrate) was uniformly labeled 14C-straw. The value of the partial
control was calculated by determining the difference in substrate utilization
between unfumigated and fumigated samples and deriving the formula:

Biomass C=O.229+0.0882(fumC/conc)+O.O969(fumC/conc)2-
0.016(fumC/conc)3

where, fumC is the fumigated ﬂush after 10 days of incubation and cone is
the amount of C mineralized in 10 days in an unfumigated soil (Horton,
Horwath and Paul, personnel communication). The corrected CFI value is
remarkably similar to the results of direct counting (Figure 6.1c and Table
6.5).

Long-Term Incubations

Long-term incubations were done on the soil from the second and
third samplings of the labeled trees and the ﬁrst two samplings of the leaf
litter exchange plots. The object of the long-term soil incubations was to
determine the amount of recently assimilated tracer 140 and 15N entering
the different fractions of the SOM from root and leaf litter derived C and to
determine the contribution of the microbial biomass to the mineralizable C
and N pools. The data were also useful to calculate rate constants and pool
sizes for C mineralization. Nitrogen mineralization was represented by a
straight line, and therefore, cumulative N mineralization could not be

described using ﬁrst order kinetics.

Soil C

The accumulation of mineralized C (ug COz-C g'l) declined as the
long-term incubations progressed for both tree and leaf litter plots (Figure
6.2a and 6.3a, Appendix B.1-B.10). Carbon mineralization values for
surface soil from the tree plots were 315-708 ug C g'1 soil. Carbon
mineralization values for subsurface soils dropped to one sixth of this
value. The decline in the accumulation rate of C is associated with the
mineralization of a labile C soil pool and the subsidence of a possible
disturbance effects associated with sampling. The C mineralization curves
for the labeled tree plots remained essentially similar over time, especially
in the surface soil (Figure 6.2a). The C mineralization from the leaf litter
exchange plots were higher than the tree plots and decreased from the ﬁrst
to second sampling (Figure 6.3a). This was similar to the results of the
microbial biomass determination for tree plots. The level subsides to values
resembling the tree labeled plots for the second sampling and long-term
incubation and was similar to the results of the microbial biomass
determinations.

The mineralization of 14C followed patterns similar to that of the
native C for both tree and leaf litter plots (Figures 6.2b and 6.3b). The
amount of 14C mineralized declined in the second long-term incubations
(soils sampled at greater than 300 days in the ﬁeld) for all plots. The decline
in speciﬁc activity of the mineralized C from both the leaf litter and tree
plots is less pronounced for the long-term incubations from soils sampled
between 328-372 days (Figures 6.4a-d) showing that ﬁeld microbial activity
had removed much of the available 14C. This pattern of decline is
inﬂuenced by the date of 1‘er labeling. The 14'0 data for the September

labeled trees showed greater translocation below-ground.

 

 

 

 

 

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ﬁrst (60 days) and second (366 days) long-term incubation of September
labeled trees. The graph lines are simulated values calculated using
equation 2. Standard error of the mean shown as line bars.

 

 

 

 

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Figure 6.3. The C (Fig. 6.3a)and 14 C (Fig. 6.3b) mineralization data for the
ﬁrst (168 days) and second (328 days) long-term incubation of September
litter exchange. The graph lines are simulated values calculated using
equation 2. Standard error of the mean shown as line bars.

-1.

A Leaf litter
' September label

 

 

C?
N
o 3— 4) ‘
H0 i .41 + ¢¢\ \
'uo -- - - .........
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a?
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Days incubated

Figure 6.4. The decline in speciﬁc activity of mineralized C during the
long-term incubation of soils labeled through leaf and root derived C.
Figure 6.4a and 6.4b are from leaf litter plots and 6.4c and 6.4d are from
plots from the whole tree labeling. Standard error of the mean shown as

line bars.

1%

The differences in the speciﬁc activity of root and leaf derived C indicate
that the disposition of assimilated 14C is depended on growing season and
phenology as discussed in Chapter 5. The consistency in slope of speciﬁc
activity in soil incubated for 300 days from the ﬁeld shows that this material
had become equilibrated with constituents of SOM fractions.

N Dynamics

N mineralization occurred at a constant rate for all the long-term soil
incubations (Figure 6.5,6.6; Appendix B.11-B.15). The N was mineralized at
a constant rate of 0.426 ug N g‘1 (1‘1 soil for surface soil of all plots even
though C mineralization data had shown differences between root and leaf
litter. Subsurface soils released 0.08 pg N g'1 d'1. The lack of ﬁrst order
kinetics suggests a large potentially mineralizable N pool. The recently
incorporated 15N from leaf litter decomposition also displayed a linear rate
of mineralization during the long-term soil incubations (Figure 6.5). The
similar response of native N and recently incorporated N suggests that the
potentially mineralizable N pool is being utilized similarly. However, the
mineralization of unlabeled and labeled material display different slopes or
rates of accumulation. These rate differences may be attributable to the
spatial differentiation of more recently incorporated N. The turnover of 15N
in from root labeled soil was not detectable in the incubation. The N content
of the roots, especially ﬁne roots, was not large enough to enrich soil N
pools through turnover (Chapter 4). Similarly, labeled N was not detected
below 10 cm in leaf litter labeled soil. The lack of N movement past 10 cm
soil depth is probably due to efﬁcient microbial immobilization and tree

Uptake. Since some of the 14C moved to 10-25 cm and 15N did not, one

 

 

 

 

 

 

A.
100 __ June 6, 1991 _1 100
80 —- t . - so
- —c+ -NO—10cmSept.litter ¢w
:=: ' - ./' . 0?:
8 6O __ --.- -N0-10chulylitter /. ,/- _ m _.
'1' ’ " ' 2?
m _ _ .
z 40 _ ’8’. 8 4o «3‘
no —— _ ' ‘
1 : 9, (8" "B“ISNO-IOcmSepLIitter é:
. /. ./- —I- - 15N 0.10 chuly litter
20 __ ./ ' - Z)
'9.“ :-*—-e—=—‘i:.:::f—:—.:L;:::—:ﬁ
O ' I I I I I I 0
0 25 50 75 100 125 150 175
Day
B.
100 a, November 14, 1991 — 100
80 ~5~ ~ 80
E — @- -N0-10cmSepL litter ./-+
2: r -..-.N0-10chulylitter ,, K. 0%
8 60 “L— /. ' — a) 3
5° F (I) /- ‘ a?
z 40 - ,0 ' 40 .'..
go _I__ . _
. . a? a
t f " “B"‘5NO-10c1nScpt. litter
20 __ , r —I- - ”NO-mommy ﬁtter _ 21)
h C - —-—— - 7:15
i,'. 7'1"-"— :--:"_‘.g':—’ .........
0 ' I I I I I I 0
O 25 50 75 100 125 150 175
Day

Figure 6.5. Cumulative N and 15N mineralization curves from long-term
soil incubations of surface soil (0-10 cm) from the litter exchange
experiment. Figure 6.5a represents the ﬁrst sampling on June 6, 1991
and ﬁgure 6.5b the sampling done on November 14, 1991 of both
1September and July litter. Standard error of the mean shown as line

ars.

 

 

A .
100 ‘ 0 Sept. litter sampr at 168 days
[3 July litter sampled at 168 days
30 _ 0 Sept. litter sampled at 328 days
,8 I July litter sampled at 328 days
'1‘ 60 -4
00
Z. I EB a
On 40 — 8
Z 3 a '
20 - s 8 m
it a g
0 I I I I I
0 40 80 120 160 200
Days incubated
B.
75 — 0 July 0—10 cm 0' Sept. 0—10 cm
[:1 July 25—60 cm I Sept. 25-60 cm
60 ﬂ 0 July 60-100 cm 0 Sept. 60-100 cm
'§
,_. _ (D
50 45 (1) (D
2.
g 30 a 0 ¢ +
22 O
15

 

 

1 Q . (1) O
’11! a aw s ’ W a ‘9
0 I I I I I
0 40 80 120 160 200
Days incubated

Figure 6.6. Nitrogen mineralization data for all the subsurface (10-25 cm)
samplings of the leaf litter exchange (Fig 6.6a) and all soil depths of the
ﬁrst long-term incubation of July and September labeled tree plots (Fig.
6.6b). Soil depth and dates are indicated. Standard error of the mean
shown as line bars.

199

would speculate that the 14C movement was by leaching of high C material.
Soil fauna movements should have moved equal concentrations of both 14C

and 15N through the soil proﬁle.

Microbial Biomass C and N

The microbial biomass C measurements at day zero of the long-term
incubations corroborate the ﬁeld measurements discussed earlier. On
incubation, the biomass C dropped in the surface soil samples (Tables 6.6-
6.9) by approximately one third. Subsurface samples of the July label did
not show a similar drop, but those from the September label did (Table 6.7).
The leaf litter experiments again showed the increase in biomass C (Table
6.8) and the incubations of the later samplings showed that the biomass
was at similar levels to the tree plots.

Biomass N again showed somewhat similar results to that of
biomass C (Tables 6.9-6.11). The microbial biomass N from subsurface soils
of the July labeled trees increased upon incubation in the lab. Conversely,
the subsurface soil from the September labeled trees dropped in biomass N
during lab incubations. The microbial biomass N from the leaf litter plots
behaved similarly to the surface soils of both July and September labeled
trees.

The speciﬁc activity of the microbial biomass N (Tables 6.11a, b) was
similar to that of biomass C, dropping during the long-term incubation
(Tables 6.6-6.8). The drop in biomass C is an artifact of the extended
incubations and is attributable to the lack of soil C input. The drop in
speciﬁc activity of the biomass C shows turnover and increased dependence

on nonlabeled soil C pools.

Table 6.6. Microbial biomass C, speciﬁc activity and percent of applied 14C
from long term soil incubations of tree plots labeled July 19, 1990.
Standard error of the mean shown in parentheses.

 

Soil Days Biomass C Bq mg-l % of
depth in Days pg C/0.41 biomass C applied
(cm) field incubated 5:1 soil {1 soil 140*
0-25 0 186.1 (7.7) 1.1 (0.22) 1.62 (0.19)
107 51 152.5 (13.5) 0.5 (0.11) 0.55 (0.15)
124 137.3 (9.4) 0.3 (0.04) 0.37 (0.05)
0 213.3 (19.5) 0.5 (0.11) 0.84 (0.02)
372 55 195.8 (3.5) 0.4 (0.00) 0.67 (0.11)
149 139.7 (2.2) 0.4 (0.09) 0.48 (0.03)
25-60 0 27.2 (9.3) 2.0 (0.96) 0.62 (0.11)
107 51 29.0 (11.3) 0.7 (0.42) 0.24 (0.08)
124 36.9 (6.7) 0.4 (0.17) 0.19 (0.06)
0 65.0 (28.7) 0.5 (0.23) 0.34 (0.05)
372 55 44.5 (16.6) 0.4 (0.19) 0.23 (0.00)
149 52.6 (2.9) 0.3 (0.17) 0.19 (0.10)
60100 0 7.6 (7.7) 4.2 (2.46) 0.31(0.11)
107 51 14.1 (8.6) 0.7 (0.20) 0.11 (0.07)
124 18.0 (8.0) 0.4 (0.08) 0.11 (0.06)
0 23.3 (9.1) 0.8 (0.32) 0.21 (0.04)
372 55 22.6 (5.6) 1.2 (0.86) 0.13 (0.03)
149 25.4 (2.1) 0.3 (0.03) 0.08 (0.01)

 

* Based on the amount of 14'0 found in the root system and soil from the ﬁrst sampling.

201

Table 6.7. Microbial biomass C, speciﬁc activity and percent of applied 14C
from long term soil incubations of tree plots labeled September 5, 1990.
Standard error of the mean shown in parentheses.

 

 

 

Soil Days Biomass C Bq mg-l % of -=
depth in Days ug C/0.41 biomass C applied
(cm) field incubated '1 soil 5‘1 soil 14C*
0-25 0 189.8 (28.1) 2.5 (0.38) 1.16 (0.12)
60 51 118.8 (8.6) 1.4 (0.14) 0.41 (0.05)
124 110.0 (10.1) 1.1 (0.19) 0.31 (0.05)
0 219.0 (9.9) 0.7 (0.12) 0.57 (0.28)
366 61 162.5 (29.5) 0.6 (0.00) 0.26 (0.06)
151 104.8 (1.3) 0.6 (0.13) 0.19 (0.05)
25-60 0 79.7 (10.8) 2.2 (0.29) 0.66 (0.07)
60 51 43.2 (6.1) 2.5 (0.42) 0.37 (0.06)
124 47.1 (15.8) 1.9 (0.89) 0.31 (0.03)
0 70.0 (0.6) 0.5 (0.10) 0.17 (0.03)
366 61 62.2 (9.4) 0.4 (0.10) 0.12 (0.01)
151 36.0 (3.9) 0.6 (0.08) 0.09 (0.01)
60100 0 50.2 (6.8) 2.3 (0.38) 0.49 (0.02)
60 51 16.8 (1.1) 5.4 (1.27) 0.30 (0.02)
124 20.7 (4.6) 3.3 (0.69) 0.27 (0.02)
0 37.8 (6.0) 0.8 (0.27) 0.13 (0.01)
366 61 35.1 (3.5) 0.5 (0.07) 0.08 (0.03)
151 12.0 (4.7) 1.8 (0.63) 0.07 (0.00)

 

* Based on the amount of 14C found in the root system and soil from the ﬁrst sampling.

202

Table 6.8. Microbial biomass C, speciﬁc activity and percent of applied 14C
from long term soil incubations of the litter exchange plots. Standard
error of the mean shown in parentheses.

 

Soil Days Biomass C Bq mg-l % of
depth in Days ug C/0.41 biomass C applied
(cm) field incubated 5'1 soil £1 soil 140*
W
0 346.3 (17.0) 4.70 (0.78) 3.68 (0.66)
0-10 168 70 186.1 (19.8) 4.98 (0.52) 2.08 (0.42)
143 195.4 (6.1) 3.04 (0.13) 1.34 (0.18)
0 219.3 (42.2) 4.42 (0.98) 2.02 (0.22)
328 55 186.5 (15.2) 3.61 (0.18) 1.47 (0.29)
124 199.1 (7.0) 2.44 (0.29) 1.10 (0.25)
0 248.9 (10.3) 0.59 (0.32) 0.41 (0.15)
10-25 168 70 149.8 (62.5) 0.52 (0.19) 0.18 (0.08)
143 160.7 (12.1) 0.38 (0.21) 0.17 (0.07)
0 163.7 (32.1) 1.06 (0.82) 0.33 (0.10)
328 55 146.9 (22.8) 0.55 (0.19) 0.22 (0.05)
124 154.5 (39.0) 0.46 (0.18) 0.22 (0.12)
Jubdittenﬂghange
0 355.9 (31.8) 2.34 (0.37) 1.82 (0.47)
0-10 168 70 211.4 (42.3) 2.43 (0.77) 1.03 (0.11)
143 204.7 (37.1) 1.32 (0.06) 0.62 (0.10)
0 261.3 (80.6) 2.80 (1.46) 1.40 (0.14)
328 $ 189.5 (18.4) 2.33 (0.55) 1.00 (0.13)
124 219.5 (17.9) 1.32 (0.18) 0.71 (0.10)
0 216.4 (20.3) 0.37 (0.21) 0.24 (0.16)
10-25 168 70 148.8 (18.3) 0.16 (0.09) 0.06 (0.03)
143 156.4 (11.2) 0.16 (0.05) 0.08 (0.02)
0 146.3 (35.3) 0.68 (0.44) 0.20 (0.06)
328 $ 125.9 (34.4) 0.47 (0.15) 0.16 (0.01)
124 165.2 (16.7) 0.28 (0.04) 0.13 (0.02)

 

* Based on the amount of 1‘er found in the leaf litter.

Table 6.9. Microbial biomass N from the long-term incubations of soil from
the tree plots labeled July 19, 1990. Standard error of the mean shown in
parentheses.

   

 

Sol ___ "My? _ ' 2 _ ﬂ . (Bloass F

depth in Days #8 N/kn*
(cm) ﬁeld incubated 5.1 soil

0-25 0 32.1 (1.2)

107 51 26.6 (2.0)

124 24.2 (1.5)

0 34.9 (0.1)

372 $ 34.7 (1.1)

149 24.9 (0.0)

25-60 0 4.4 (1.4)

107 51 5.3 (0.2)

124 6.2 (1.0)

0 9.7 (3.8)

372 55 7.9 (2.7)

149 8.7 (0.7)

60-100 0 1.4 (1.3)

107 51 2.3 (1.6)

124 2.9 (1.1)

0 3.6 (1.1)

372 55 3.9 (0.9)

149 4.2 (0.1)

 

* Kn=0.148*(fumigated COz ﬂush/fumigated NH4+ flush)+0.39

Table 6.10. Microbial biomass N from the long-term incubations of soil
from the tree plots labeled September 5, 1990. Standard error of the
mean shown in parentheses.

 

Soil Days Biomass N

depth in Days ug N/kn*
(cm) ﬁeld incubated 5-1 soil

0.25 0 33.0 (4.4)

60 51 21.2 (1.9)

124 19.7 (2.0)

0 37.6 (1.6)

366 55 29.1 (4.5)

149 19.0 (0.4)

25—60 0 11.1 (0.6)

60 51 7.0 (1.4)

124 8.1 (2.6)

0 9.6 (0.6)

366 55 10.3 (1.4)

149 5.6 (0.8)

60-100 0 6.9 (0.9)

60 51 2.9 (0.3)

124 3.5 (0.6)

0 5.8 (1.1)

366 55 5.8 (0.7)

149 2.0 (0.8)

 

* Kn=0.148*(fumigated C02 flush/fumigated NH4+ flush)+0.39

Table 6.11a. Microbial biomass N, 15N and percent of applied 15N from soil
incubations of September litter plots. Standard error of the mean in
parentheses.

 

Soil Days Biomass N Biomass 15N % of
depth in Days ug N/kn“ ng 15NH4+-N applied
(cm) field incubated 5-1 soil g'l soil 15N§
WWW
0-10 0 60.3 (3.0) 9.1 (1.9) 8.2 (1.2)
168 70 34.4 (3.1) 4.2 (1.2) 3.7 (0.8)
143 33.9 (0.9) 2.8 (0.4) 2.2 (0.4)
0 40.7 (6.6) 4.8 (2.1) 4.3 (1.8)
3% 55 34.0 (2.7) 3.6 (1.5) 3.3 (1.6)
124 34.5 (0.8) 3.3 (1.5) 2.9 (1.4)
10-25 0 42.9 (1.9)
168 70 27.4 (10.3)
143 28.3 (1.9)
0 30.0 (5.2)
3% 55 26.7 (3.7)
124 27.4 (6.5)

 

* Kn=0.148*(fumigated 002 ﬂush/fumigated NH4+ ﬂush)+0.39
§ Based on the amount of 15N in the litter.

Table 6.11b. Microbial biomass N, 15N and percent of applied 15N from soil
incubations of July litter plots. Standard error of the mean in
parentheses.

 

 

Soil Days Biomass N Biomass 15N % of
depth in Days 11g N/kn* ng 15NH4+-N applied
(cm) field incubated £4 soil 5'1 soil 15N§
lubdittauxchange
0-10 0 62.6 (5.5) 8.5 (0.5) 4.4 (0.3)
1% 70 39.0 (6.9) 4.4 (1.6) 2.3 (0.8)
143 35.8 (5.8) 2.9 (1.0) 1.5 (0.5)
0 42.0 (6.5) 6.9 (2.2) 4.0 (1.2)
3% 55 35.1(3.0) 3.7 (2.3) 2.1(1.3)
124 38.2 (3.1) 4.1(2.6) 2.1(1.3)
10-25 0 37.6 (3.5)
1% 70 27.4 (3.0)
143 27.4 (2.0)
0 27.2 (5.7)
3% 55 23.3 (5.7)
124 28.6 (2.7)

 

* Kn=0.148*(fumigated C02 ﬂush/fumigated NH4+ ﬂush)+0.39
§ Based on the amount of 15N in the litter.

N7

The changes in the leaf litter experiment (Tables 6.11a, b) showed a
four fold drop in biomass 15N for the long-term incubations of soil sampled
at 168 days. The soil sampled at 328 days showed lower amounts of 15N in
the biomass, but only a 30% drop during the lab incubation showing that
similar results are found during ﬁeld and laboratory exposure. The
microbial biomass accounted for 8.2% of the 15N at 168 days and only 4.3%
of the biomass N at 328 days in the September litter. The July litter showed
a similar drop in 15N (ng g'1 soil), but did not show a similar drop when
expressed as a percent of applied. The lack in the decrease in the amount of
15N applied in the July litter is in most part attributable to the larger
amount of labeled N in the leaf litter lain on the plots. The larger amount of
15N found in the July leaf litter was a result of the increased sink strength
for N of leaves during mid-growing season (Chapter 3 and 4).

The increase in mineralized 15N corresponds to a decrease in
microbial 15N in the leaf litter plots (Figure 6.7). A similar decline in
microbial biomass 14C is associated with the accumulation of mineralized
14C (Figure 6.8). This demonstrates the importance of the microbial
biomass as a component of the labile C and N fraction of the SOM. The
microbial biomass N pool may also represent the portion of the labile N
fraction that mineralizes at a different rate. The loss of N from the
microbial biomass amounts to approximately 50% of the total N mineralized
in the 0-25 cm soil during the long-term soil incubations. About 25% of the
C mineralized can be accounted for from the loss of microbial biomass C

during the long-term incubations.

15 +

 

 

—9— Biomass l5N ,
.‘é
7w
nz
a”
l
O 50 100 150
Day
um ’ I ’ ’ i
E ’ ’ . ,
3?

 

100 150

 

Figure 6.7. The amount of mineralized 15N and microbial biomass 15N for
the surface soil of the September labeled litter. Figure A represents the
ﬁrst sampling on June 6, 1991 and ﬁgure B the sampling done on
November 14, 1991. Standard error of the mean shown as line bars.

Bq mg'l COz-C

 

 

0 25 50 75 100 125 150 175

7 —e—Mhmhzod”c
' 'B' ' Biomass ”C

Bq mg'l COz-C
M

W
I
I
I
I
I
I

N

 

O 25 50 75 1% 125 150

Figure 6.8. Speciﬁc activity of mineralized C and microbial biomass C for
the surface soil of the September labeled litter. Figure A represents the
ﬁrst sampling on June 6, 1991 and ﬁgure B the November 14, 1991
sampling. Standard error of the mean shown as line bars.

210

Potentially Mineralizable C

The models to describe C mineralization produced good ﬁts (Figures
2b and 3b), consequently, no difference between equations 1 and 2 were
observed. Equation 2 was used to develop all of the rate parameters and pool
sizes of SOM fractions. The C mineralization during the ﬁrst 50 days of the
long-term soil incubations followed ﬁrst order kinetics. The amount of C
mineralized during this period was approximately 50% of the total C
mineralized. Zero and ﬁrst order kinetics described the C mineralization
during the latter part of the long-term incubations equally well. The
mineralization of C at soil depths below 25 cm was substantially lower than
the surface soil. This indicates that SOM pools are smaller in subsurface
soils.

First order rate constants for labile unlabeled C (C1) lie between
0.014-0.036 d'1 for surface soil above 25 cm for all plots using equation 2
(Table 612-6. 14a). The ﬁrst order rate constants for the C1 pool below 25 cm
are variable and ranged from 0.01-0.09 d'l. The variability is associated
with the low amount of C mineralized in subsurface soil and the possible
greater effects of disturbance from sampling. Rate parameters for the
more stable C pool (C2) in surface soil ranged from 0.0001-0.0005 d'l. The
mean residence time (MRT) for the C1 pool was 24-74 days for surface soil
and 11-54 days for subsurface soils for all plots. The Cl pool amounted to 1-
3% of the SOM. The MRT of the Cz pool was variable and lies between 6 to
23 years for all soil depths. The C2 pool was 47-49% of total C in SOM as
estimated by subtracting out the Cl and considering the remaining soil C
(50%) to be recalcitrant and not contributing to mineralized C during the

incubations.

211

Table 6.12. Parameters developed from the modiﬁed exponential equation
applied to the 002 mineralization data from long-term incubations of
soil from the sampling of labeled tree plots. MRT is mean residence

 

 

time.

Soil Days C1 C2
depth in K1 K2 C1 02 1.1g C g-1 pg C g-l
(cm) ﬁeld d-1 d-1 MRT d-l MRT y-l soil soil

lubhlabelsdm

0-25 107 0.04 0.0004 26.9 7.6 112.3 5187.7
372 0.04 0.0003 23.7 8.8 105.8 5194.2

2560 107 0.09 0.0002 11.2 13.7 25.9 1824.1

60-100 107 0.09 0.0003 11.2 9.8 29.1 1320.9

WM

025 60 0.01 0.0002 73.7 17.1 237.7 5062.3
366 0.03 0.0003 33.2 8.8 85.0 5215.0

2560 60 0.04 0.0002 26.5 14.4 39.7 1810.3
366 - - - - - -

60-100 60 0.02 0.0001 54.3 22.8 45.5 1304.5
366 - - - - - -

 

212

Table 6.13. Amount of C found in the soil microbial biomass of the trees
labeled in July and September 1990 and C mineralized at 150 (C150).

Csoil is total soil C. Biomass C is C in the soil biomass.

 

Soil C150 Biomass C/ Biomass C/
depth pg C g‘1 C l/Csoil C150 Csoil
(cm) Day soil % % %
Wes
0-25 107 384.61 1.06 48.4 1.8
372 341.63 1.00 62.4 2.0
25-60 107 79.80 0. 70 34.1 0.7
372 111.96 - 58.1 1.8
60-100 107 80.23 1.08 9.5 0.3
372 94.84 - 24.6 0.9
W
0-25 60 326.68 2.24 58.1 1.8
366 321.02 0.80 68.2 2. 1
25-60 60 90.47 1.07 88.1 2.2
366 84.10 - 83.2 1.9
60-100 60 65.92 1.69 76.2 1.9
366 148.97 - 25.4 1.4

 

213

Table 6.14. Parameters developed from the modiﬁed exponential equation
applied to the CO2 mineralization data from long-term incubations of
soil from the sampling of leaf litter exchange plots. MRT is mean
residence time. C3011 is total soil C.

 

—
8011 Days C1 ()2
depth in K1 K2 C1 02 pg C g’l 11g C g-l
(cm) field d-l d-l MRT d-l MRT y-l soil soil
Sentemhenlitteunhanae
0-10 168 0.02 0.0005 46.6 5.7 308.0 5192.0
328 0.03 0.0005 33.2 5.7 185.7 5314.3
1025 168 0.02 0.0003 64.4 8.3 288.4 4761.6
328 0.02 0.0003 49.1 10.1 186.0 4864.0
JubhlitteLenhanse
0-10 168 0.01 0.0003 68.4 8.3 421.6 5078.4
328 0.02 0.0005 41.1 6.0 213.3 5286.7
1025 168 0.02 0.0004 42.9 6.4 123.0 4927.0

328 0.02 0.0003 59.3 8.8 177.2 4872.8

 

214

Table 6.14a Represents the amount of C found in the soil microbial biomass
of the leaf litter exchange experiment and C mineralized at 150 (C 150)
days. Csoil is total soil C. Biomass C is C in the soil biomass.

 

Soil C150 Biomass C/ Biomass C/
depth pg C g‘1 Cl/Csoil C150 Csoil
(cm) Day soil % % %

Waugh
0-10 1% 656.36 2.80 52.8 3. 1
3% 552.88 1.69 39.7 2.0
10- 1% 490.32 2.86 50.8 2.5
2510/25/ 1992
3% 370.29 1.84 44.2 1.6
lubdittauxchange
0-10 1% 619.74 3.83 57.4 3.2
328 560.25 1.94 48.4 2.5
10-25 1% 427.01 1.22 50.7 2. 1

3% 384.45 1.75 38.1 1.4

 

215

The amount of C mineralized at 150 days (C150) of incubation
decreases 3 fold with soil depth. Microbial biomass C decreases 6 fold with
depth (Tables 6.6 and 6.7). The microbial biomass C to C150 ranges from
0.1-0.88 from subsurface to surface soil, suggesting that the
microorganisms in subsurface soil appear to account for proportionately
more of the C mineralized during the long-term incubation. Both equations
1 and 2 did not describe C mineralization data from subsurface soil from
the third sampling, one year latter, for the labeled tree plots. Cumulative C
mineralization curves for these subsurface soils never approach asymptotic
values (Figure 6.2a). Soil handling procedures and storage may inﬂuence
subsurface soil to a greater extent because of their small potentially
mineralizable C pools. The potentially mineralizable C is generally less
than 3% of total soil C. This labile C pool is equivalent to the amount of C
contained in the microbial biomass (Table 6.13-6.14b). Microbial biomass C
accounts for approximately 50% of the C mineralized during 150 days of
incubation, especially in surface soil, showing the potential cycling of C
through the microbial biomass.

The ﬁrst order rate constants for labeled C are similar to the rate
constants for native C (Table 6.15 and 6.17a). However, the pool size of the
recently incorporated C dwarfs that of the native C. As a result, a greater
proportion of the small pool of recently incorporated C is mineralized
during the long-term soil incubations, 1400/14Csoi1. where 1400 is the
mineralizable 14C and 1403011 is total 14C in soil (Tables 6.17 and 6.17a).
The potentially mineralizable 14C and mineralization rate of more stable
soil 140 (02-140) was higher in leaf labeled soil. The labile 140 pool (C1-
14C) accounted for 1.8-12.6% of total soil 140 in the labeled tree plots and 5.4-
36% in leaf litter plots. The MRT of 01-140 is between 14-64 days for all soil

216

Table 6.15. Parameters developed from the modiﬁed exponential equation
applied to the 14CO2 mineralization data from long-term incubations of
soil from the sampling of labeled tree plots. MRT is mean residence

 

time.
Soil Days C1-14C (32,140:
(cm) field d-l d-l MRT d-l MRT 1'1 soil soil
«lubLlabeledJmes
0-25 107 0.04 0.0006 26.8 4.3 0. 14 1.3
372 0.03 0.0007 31.0 3.8 0.10 1.1
25-60 107 0.07 0.0002 13.9 12.5 0.03 0.6
372 0.07 0.0004 14.6 6.8 0.03 0.7
60-100 107 0.05 0.0002 20.6 14.4 0.02 0.5
372 0.07 0.0003 14.3 8.3 0.01 0.4
W
0-25 60 0.04 0.0006 22.7 4.4 0.31 2.2
366 0.02 0.0003 54.8 9.4 0.21 3.3
25-60 60 0.05 0.0011 21.7 2.6 0.10 0.9
366 0.05 0.0002 18.3 16.1 0.02 1.2
60-100 60 0.05 0.0010 19.6 2.8 0. 11 0.9
366 0.03 0.0005 29.9 5.7 0.03 0.5

 

* C2~14C calculated assuming no recalcitrant 1“IC has formed during the duration of the
experiment.

217

Table 6.16. Represents the amount of radiolabeled C found in the soil
microbial biomass and 14C mineralized at 150 days (14C150). 14Csoil is
total soil 14C. Biomass 14C is 14C in the soil biomass.

 

SO“ i“C150 C1-uC/ Biomass i4C/ Biomass uC/
depth Bq 14C 8'1 14(3soil 14”(31.50 14Csoil
(cm) Day soil % % %
MW
0-25 107 0.26 9.45 82.2 14.5
372 0.21 8.36 51.7 9.2
25-60 107 0.06 5.08 89.8 7.3
372 0.07 4.01 42.7 4.1
60-100 107 0.03 3.56 57.7 3.6
372 0.03 2.58 71.6 5.2
Septembenlabslsitnees
0-25 (1) 0.51 12.64 90.9 18.5
366 0.34 6.13 55.9 5.4
25-60 (1) 0.23 9.88 75.4 17.5
366 0.05 1.84 75.9 3.3
60-100 60 0.24 10.71 46.3 10.5
366 0.07 5.64 44.2 5.4

 

218

Table 6.17. Parameters developed from the modiﬁed exponential equation
applied to the 14C02 mineralization data from long-term incubations of

soil from the sampling of the litter exchange plots. MRT is mean
residence time.

 

Soil Days (31,140 02-140“
depth in K1 K2 (31.140 (32.140 Bq g-l Bq g-l
(cm) field d-l d-l MRT d-l MRT y-l soil soil
Wage
0-10 168 0.02 0.0008 44.7 3.3 1.77 4.7
3% 0.02 0.0005 63.8 5.4 ‘ 0.85 6.0
10-25 168 0.02 0.0006 44.2 4.4 0.12 0.4
3% 0.03 0.0001 34.5 22.8 0.03 0.5
Wm
0-10 1% 0.02 0.0011 46.1 2.4 0.83 1.8
328 0.02 0.0009 53.4 2.9 0.44 3.3
10-25 1% 0.03 0.0017 31.8 1.6 0.08 0.1
3% 0.02 0.0013 44.0 2.2 0.04 0.4

 

* C2-14C calculated assuming no recalcitrant 14C has formed during the duration of the

experiment.

 

 

6'91 99:. Z96 019 969
999 9'ZL 9699 116 991 9391
V91 62.9 1.911 1790 969
0'19 991. 6602 60'I 991 011)
MW
916 1196 999 1790 96s:
6661/96/0196
7‘88 1‘86 79178 910 991 -01
1'91 991. W61 131 999
6'96 6'11. 392% QZ'Z 991 011)
WWW
% % % llos 48a (m0)
“”011 0910171 “”0171 143 0171 ”8 tilde?
lam 889mm long 888111019 Ion-10 ”it 1103

“99131110191109 91121 111 0171 31071 993111018 '071 HOS 1910’) 9113090171
“(091 Q“) 091 is pazyfexaunn Qﬂ pus quampadxa eﬁueqoxe 161111 am JO

889111019 18190109111108 919 u! puma 0 991998101981 .10 mourv 9119 9199.1

613

2%

depths and plots. The C2-14C pool accounted for 37.4-48.2% of total soil 140
for both tree and leaf litter plots. The MRT for the C2-14C fraction was
between 2- 16 years.

The microbial biomass 14C content was 44-91% in tree plots and 68-
93% in leaf litter plots of the 14C mineralized during 150 days of incubation.
Again this shows the important aspect of cycling C through the microbial
biomass. The litter labeled soil contains more labeled C in both microbial
biomass and potentially mineralizable fractions than does the root labeled
soil. This suggests that leaf litter incorporates more C into both labile and
more stable SOM pools during the ﬁrst year following the incorporation of
labeled substrates to soil. However, as mentioned earlier, the incorporation
of C into soil from leaf litter is spatially limited to the surface soil.
Movement of labeled C through the soil proﬁle is becoming evident in leaf
labeled soil and can be seen in both biomass and fractions of SOM (Table
6.17b). The duration of the current experiment was not long enough to
predict the magnitude of the contribution of leaf litter C to subsurface soils
as compared to root derived C.

Table 6.18, shows the consistency of N mineralization over time and
soil depth. The microbial biomass N and N mineralized at 150 days (N150)
accounted for 4-8% of total soil N in the surface soil of all plots. The
microbial biomass N to N mineralized during 150 days of incubation was
from 0.5-1.6, indicating the importance of microbial N as a component of
labile N fraction and contribution to the potentially mineralizable N pool.
The ratio of C150 to N150 generally declines with soil depth indicating that
preferentially more C than N was mineralized deeper in the soil proﬁle.

The differences in C and N mineralization occurred at a relatively constant

221

Table 6.18. N mineralized was calculated by linear regression. N150 is the
N mineralized 150 days. N301] is total soil N. Biomass N/N15o is a ratio.

 

 

 

 

 

 

w
Soil Days N mineralized N150/ Biomass N/ Biomass N:
depth in at 150 days N soil- N soil N150
(C m) field ”£5180“ % %
Wm
0-25 107 35.85 4.48 4.01 0.90
372 41.35 5.17 4.36 0.84
25-60 107 3.28 0.99 1.33 1.34
372 6.17 1.87 2.39 1.28
60100 107 2.23 0.89 0.56 0.63
372 4.13 1.65 1.44 0.87
Sentemhenlabeledjmes
0-25 60 37.81 4.73 4.13 0.87
366 40.80 5.10 4.70 0.92
25-60 60 7.70 2.33 3.36 1.44
366 6.24 1.89 2.91 1.54
60-100 60 5.01 2.00 2.76 1.38
366 3.61 1.44 2.32 1.61
I 'I | l . |
Soil Days N mineralized N150] Biomass N/ Biomass N:
depth in at 150 days N soil- N soil N150
(cm) field “3'1 soil % %
WW9
0-10 1% %.87 6.82 5.97 0.88
3% 76.99 7.62 4.03 0.53
10-25 168 49.78 5.41 4.66 0.86
3% 51.61 5.61 3.26 0.58
luhLlimLenhange
0-10 1% 67.18 6.65 6.20 0.93
328 79.66 7.89 4.91 0.62
10-25 168 48.52 5.27 4.09 0.7 7

3% 52.67 5.73 2.96 0.52

 

222

C:N in the microbial biomass. This indicates that N may have been

limiting in subsurface soil.

DISCUSSION

This research was established to measure the contribution of tree
root and leaf litter to microbial biomass and SOM dynamics. The pools of C
and N were examined using long-term laboratory incubations of a ﬁeld
labeled tree-soil system. Additionally, I were interested in comparing and
estimating the amount of C and N in microbial biomass. There are a large
number of CFI and CFE studies, but these are rarely standardized against
direct microscopy, against each other or conducted on subsurface soil. The
intent of these comparisons was to select and verify a consistent assay for
both the determination of microbial biomass size and content of tracer 14C

and 15N.

Microbial Biomass Determinations

Fungal biomass values of 153-215 11g C g'1 soil and bacterial biomass
of 30-57.8 11g C g'1 soil in the temperate grasslands of Canada (data
recalculated from Paul et al., 1979) are similar to our results. Babiuk and
Paul (1970) found bacterial numbers of 2.2-4.6 109 g‘1 in 0-30 cm grassland
surface soil and 1.2 x 109 g“1 soil at 75-90 cm. My bacterial numbers are
similar for surface soils , but much lower for subsurface soils. Variations
in total hyphal lengths, bacterial numbers and diameters will be inﬂuenced
by soil physiochemical characteristics and nature of the plant community.
Hyphal diameters have been shown to decrease with decreasing amounts of

organic matter present in the soil (Schniirer et al., 1985). These

223

researchers found that hyphal diameters ranged from 1.6 to 2.7 pm in
arable soils including a fallow treatment. The hyphal diameters averaged
2.3 um for surface soils (0-25 cm) across a range of cultural treatments on
the same soil type (Harris, private communication). No hyphal diameter
estimates were done on subsurface soils in this study. Hyphal diameters
may have been inﬂuenced by the decreasing amount of organic matter in
the subsurface soil.

The degree of hyphal vacuolization can also inﬂuence fungal C
estimates. I did not separate active from inactive hyphae. The viability of
fungi can be determined with ﬂourecien diacetate which is cleaved to
ﬂourecien by cytoplasmic esterases in living hyphae. This technique has
shown that between 2-10 % of hyphae contain cytoplasm (Schniirer et al.,
1985; Paul and Clark, 1989). If the hyphal cytoplasmic content changed
with soil depth and cytoplasm represents such a low percentage of the fungi
we must ask why all of our microbial biomass measurements give such

well correlated results.

Long-Term Soil Incubations

The mineralization rate of C and N will be affected by the quality and
amount of organic C and N. The labeling of trees in the ﬁeld made it
possible to examine the steady state ﬂux of C and N from tree litters into two
of the active fractions (C1 and C2) of the SOM. The size of the microbial
biomass declines in all of the long-term soil incubations. A constant C/N
indicates that the representatives of the microbial population are
diminishing at a constant rate. The decline in biomass C and N is typical of
long-term soil incubations (Carter and Rennie, 1982; Bonds et al., 1988;
Robertson et al., 1988) and is thought to contribute to the C and N

224

mineralization (Paul 1984; Juma and Paul, 1984; Bonde and Rosswall,
1987).

In this study, a substantial amount of the N mineralized (3569 pg N
g'1 soil) could be accounted for by the decrease in microbial biomass N over
the long-term incubation in surface soil of all plots. In subsurface soil, the
amount of N mineralized during 150 days of incubation was equal to
biomass N. The decline in biomass C accounted for approximately half the
C mineralized during the incubations. Under ﬁeld conditions the biomass
is regenerated by the addition of new substrate. However, the size of the
microbial biomass is sufﬁciently large to act as a temporary pool of C and
N. This reservoir of C and N plays an important role in supplying
nutrients for plant uptake.

The rate of N mineralization remained unchanged during the long-
term soil incubations. As a consequence, pools of active and stable N were
mathematically undeﬁnable. The mineralization of recently incorporated
15N from leaf litter decomposition behaved similarly. However, the
calculated slopes from the mineralization of labeled and unlabeled N
indicated that the rates of mineralization differed. The difference in
mineralization rates of labeled and unlabeled N suggest that the recently
incorporated 15N resides in a separate pool of organic N such as microbial
biomass (J uma and Paul, 1984; He at al., 1988). My data indicated that a
decline in microbial biomass 15N corresponded to an increase in
mineralized 15N. The exchange of 15N between these pools exempliﬁes the
importance of the microbial biomass as an important labile reservoir of N.

The trees exhibited no change in total N from the second to third
growing season (Chapter 4). The loss of N through leaf fall amounted to
approximately 60% or 12.8 g N of total tree N. The microbial biomass

225

contained 19.4 g N m'2 or one and one half times one years litter fall. The
amount of N mineralized during 150 days of laboratory incubation was
equivalent to the amount of N found in the microbial biomass in the surface
soil. The comparison of 150 days of laboratory incubation to one year of ﬁeld
conditions indicates that sufficient N mineralization would be available to

replace the loss of N through leaf fall.

Potentially Mineralizable Pools of C

First order rate kinetics have been used in simulation models to
describe the decomposition of organic matter (Jenkinson and Rayner, 197 7;
Paul and van Veen, 1978). The use of ﬁrst order models implies that the
ability of the microbial biomass to decompose plant residues is not rate
limiting (Paul and Clark, 1989). These attributes of the microbial biomass
include large numbers and rapid growth rate (V oroney et al. , 1989). Models
used to describe potential C mineralization included both mixed (ﬁrst and
zero order) and modiﬁed double exponential components. The mixed order
and double exponential equations both described C and 14C mineralization
well. The unlabeled and labeled C mineralization curves were ﬁt well
using both models. Bonde et al. (1988) comparing both mixed and double
exponential models found that the mixed order model gave the best ﬁt to N
mineralization data except in straw amended soil. The mixed model
described the linear characteristics of the N mineralization in their soil.
The constant rate of N mineralization was attributed to a constant rate of
fungal hyphae growth without an equivalent increase in cytoplasm (Bonde
and Rosswall, 1987).

N mineralization failed to approach ﬁrst order kinetics during our

long-term soil incubations indicating that C and N mineralization kinetics

226

are different and that the N pool was very large. Nitrogen and C
mineralization also proceeded at different rates within the soil proﬁle.
Proportionately less N was mineralized in subsurface soil. Most studies on
N availability have concentrated on surface soils. This study indicates that
subsurface soil N mineralization behaves differently to surface soil. A
reﬁned approach to potential N mineralization potential should include N
mineralization in subsurface soil and C and N relationship of the microbial
biomass to accurately deﬁne potential N mineralization.

The labeled and unlabeled C had similar ﬁrst order decay rates
describing C1 fraction in the surface soil. In contrast, decay rates were
higher for recently incorporated C into the more stable C2 SOM fraction.
The higher decay rate for the recently added C in the C2 SOM pool indicates
that this C does not have the same characteristics as the entire resistant
fraction. This observation was seen for the incubation of labeled residues
(J enkinson 1965) and microbial products in soil (Shields et al., 1974; McGill
et al., 1975; van Veen et al. , 1987). These results can be explained by the fact
that the equations describe the C mineralization curves well, but may
oversimplify the processes occurring (V oroney et al., 1989). The equations
do not take into account the formation and decay of the microbial biomass,
production of stable organic matter and or protection and priming of
substances in the presence of others. Additionally, the individual reactions
inﬂuencing the rate of decomposition and stabilization of C into SOM
fractions may not follow ﬁrst order kinetics, rather sum into a ﬁrst order
description of the entire process.

It is difficult to exclude microbial biomass C and N from the measure
of the potentially mineralizable pools. Indeed, the microbial biomass maybe

an excellent indicator of the potentially mineralizable pools (Paul, 1984;

227

Bonde et al., 1988). The reproducibility of the biomass and mineralization
rates make their combination an excellent parameter in determining SOM
dynamics. These attributes were very useful in detecting the difference
between root and leaf litter labeled soil.

The root labeled soil from the September labeled trees contained more
potentially mineralizable labeled C than the July root labeled soils and is
similar to leaf labeled soil. The increased 14C in this soil was evident
shortly after the September labeled trees had assimilated 14C (Table 6.4).
The increased enrichment of microbial biomass with 14C is probably a
result of the increased export of C to the root system during autumn. The
rapid incorporation of 14C by the microbial biomass may have been a result
of the assimilated C leaking into the rhizosphere. An increase in soluble
sugars occurs in the root system at this time of year and may explain the
above scenario (Chapter 5). The mineralization rate of 14C and microbial

14C subside during the following season and are similar to the July root

labeled soil.

CONCLUSIONS

The comparison of microbial biomass determination methods
showed good agreement. No signiﬁcant differences and high correlation
values indicated that the different constants and variables used were
applicable across the soil depths analyzed. The use of a partial control for
the CFI method produced uncannily similar values to direct microscopy
results. The comparison of CFI and CFE methods were also very similar

despite the subtraction of a control in the CFE method.

2%

Long-term soil incubations of the root and leaf litter labeled soil was a
useful method to characterize the pools of SOM. In combination with
labeled C and N, this method is able to distinguish the fate of recently
incorporated soil C and N. The comparison of root versus leaf litter labeled
soil indicates that leaf derived C and N has enriched SOM pools to a greater
extent than root labeled soil. Both the kinetic analysis of pool size and
mineralization of recently incorporated C and N conﬁrm that in the year
following the dual labeling of trees with 15N and 140, leaf litter has
conserved more organic matter and contributed a greater quantity of C and
N to potential nutrient cycling processes.

The amount of N mineralized during the long-term incubations was
sufﬁcient to supply the N uptake of trees during the next season of growth.
The substrate to drive potential N mineralization in the initial years of
plantation establishment was determined to be dominantly leaf litter. This
conclusion was reached from results on C and N allocation patterns
determined for these trees. However, the root labeled soil has enriched the
SOM to greater soil depths. The spatial difference in the quantity of C and
N recycled by root and leaf litter suggests these litters may have different
roles or strategies that conserve SOM in the soil solum. Movement of
recently incorporated C from leaf litter past 25 cm soil will probably occur
as time progresses.

It is evident from the duration of this experiment that the ﬁnal
contribution of C and N from root and leaf litter to SOM conservation was
not determined. Future research in this area will require extended periods
of study beyond one year to fully assess the contribution of root and leaf litter

to the conservation of SOM and nutrient cycling processes.

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Fry (eds.) Academic Press, Inc., New York.

Voroney, RP. and E. A. Paul. 1984. Determination of kc and kn in situ for
calibration of the chloroform fumigation-incubation method. Soil
Biol. Biochem. 16:9-14.

Zak, D. R. and K. S. Pregitzer. 1990. Spatial and temporal variability of
nitrogen cycling in northern lower Michigan. Forest Sci. 36:367-380.

Chapter 7

CONCLUSION

The interpretation of C, N and other associated nutrient cycling
processes has become a necessity in systems managed for increased
productivity. The managed ecosystem often contains limited plant species
and is subject to an increase in cultural inputs. The practical management of
ecosystems designed to increase production ideally predicates embodying the
concept of sustainable nutrient cycles. Intensively managed ecosystems may
require a compromise between sustainability and reliance on cultural inputs
which may have environmental pollution control potential.

The study of litter turnover, nutrient cycling and SOM dynamics has
been a theme of countless studies endeavoring to elucidate the functions and
processes underlying the nutrient cycles of ecosystems. Unfortunately, the
lack of suitable techniques and limited examination of long-term ﬁeld
processes has led to an incomplete understanding of sustainable nutrient
cycles. Nutrient cycles are strongly inﬂuenced by the production and
turnover of necromass from primary producers. The simultaneous growth
and decomposition of nutrient-rich plant tissue is a difﬁcult process to resolve
using periodic sampling methods to examine the change or loss of standing
biomass. These problems are exempliﬁed in the study of below—ground
systems characterized by spatially complex root systems undergoing
continual production and turnover.

Field dual labeling with 14C and 15N was used to examine below-
ground C ﬂow and the allocation and turnover of N in the tree. The

advantage of isotope methods over samplings methods that estimate growth

234

235

and loss was that production and turnover processes could be simultaneously
isolated by following long-term tracer ﬂux and dilution. The use of isotope
methods provided a direct estimate of production and turnover of the root
system by following tracer dilution in standing biomass and ﬂux to soil
microbial biomass and organic matter. Though tracer techniques have been
used extensively, their application under ﬁeld conditions, such as in the
current study, to probe long-term nutrient cycling processes has been done
infrequently.

The application of tracers (14C and 15N) to ﬁeld grown two-year-old
trees with heights of greater than three meters to study nutrient cycles of the
above- and below-ground system was unique. The complexity of ﬁeld
engineering used was necessary to ensure that the ﬂux of photoassimilates
was characteristic of the ﬁeld environment. This complex system was chosen
over other 14C labeling techniques, such as short-duration pulse-chase and
open-top chambers, because of the degree of environmental control afforded
by closed systems.

The 15N was injected into the stem to preferentially label tree
components. This unusual approach was done to follow both internal N
retranslocation and ﬂux of N through plant biomass turnover. The dual
labeling approach provided the opportunity to determine the ﬂux of both
tracers in the plant biomass, especially the below-ground system. The
application of 1‘er and 15N and traditional methods of plant-soil sampling
under ﬁeld conditions represented a infrequently used quantitative approach
to studying C and N dynamics in tree-soil systems.

The standing biomass of leaf litter contained signiﬁcantly more C and
N than ﬁne root biomass. The standing biomass values in themselves

indicated than the ﬁne roots would need to turnover frequently to attain the

236

nutrient cycling potential of leaf litter. The standing biomass values did not
correlate to the amount of 140 found in soil from leaf and root derived C
sources. The discrepancy in the amount of 14C found in soil was further
complicated by the slow rate of 14C dilution in the cell wall fraction and
stable mass of ﬁne roots during the following year after labeling. However, a
dilution of 14C in nonstructural root components occurred between the
periods of labeling and the beginning of dormancy for the July labeling and
into the next growing season for the September labeling. During these
periods, the highest level of 140 was found in the soil containing labeled
roots.

The slow dilution of the root cell wall fraction could have resulted from
the synthesis of new roots from labeled reserves. Reserve material in the root
system was sufﬁcient to reconstruct the ﬁne root system 1.3 times. Above-
ground reserves may have contributed additional substrate for the growth of
roots. It is plausible to maintain the 14C content of the ﬁne root cell wall
fraction with no signiﬁcant changes in mass during the year following
labeling. In combination with the synthesis of new roots, additional loss of
14C may have occurred from sloughing, exudation and ﬂux to mycorrhizal
symbionts.

The 15N from leaf litter was detected in soil and microbial biomass
within 168 days in plots containing labeled leaf litter. In root labeled plots,
no 15N was detected in the soil. This result was apparent from the small
amount of N contained within the standing biomass of roots as compared to
leaves. The detection limit of 15N may have become compromised due to the
small levels of N associated with the growth and turnover of roots. Though

similar amounts of C are involved in both leaf and root turnover, leaf litter

237

turnover appears to dominant N cycling processes in the early stages of the
rotation.

The analysis of microbial biomass and associated soil organic fractions
was important in determining the contribution of C and N from leaf and root
turnover. The examination of different microbial biomass techniques
compared biomass estimates and tracer contents of soil microorganisms. In
general, direct microscopic counts and chloroform fumigation methods
(chloroform fumigation incubation, CFI and chloroform fumigation extraction,
CFE) were well correlated across time and soil depth. The subtraction of a
partial control from the CFI estimates resulted in remarkably similar
biomass values to that obtained by microscopic counting. The CFI and CFE
methods gave similar biomass C values, but differed in 14C content,
indicating that the techniques were sampling different components of the soil
microbial biomass.

The stabilization of 1‘er in soil is a measure of the amount of C
entering organic matter from litter turnover. Similar amounts of C entered
the soil from both leaf and root turnover. The difference between leaf and
root turnover lies in the spatial deposition of the stabilized C. The C entering
the soil from leaf litter decomposition was localized in the surface 0-10 cm
layer. An increase in the amount of 14C below 10 cm indicated that
eventually more C from leaf litter turnover would leach or move by faunal
action to deeper soil depths, especially during successive rotations.

Kinetic analysis of C mineralization during long-term soil incubations
showed that the labile C pools of both leaf and root labeled soil had similar
turnover rates. The intermediate resistant fraction also was not different for
the two treatments. The turnover of labile soil 1‘er occurred in 14-64 days.
The 14C stabilized into pools of intermediate resistance had a turnover time

238

of 2-16 years. In contrast, the unlabeled C fractions had turnover times of 11-
74 days for labile C and 6-23 years for fractions of intermediate resistance.
The intermediate resistant C pool turnover time increased with soil depth for
both litters. This showed that C stabilization increased with soil depth and
was probably associated with the change in amount, activity and makeup of
subsurface microbial populations. The discrepancy between the turnover
times of intermediate resistant C of recently incorporated 14C and native C
indicated that this pool should be further divided. Separating the stabilized
C into meaning fractions will enhance the understanding of the processes
involved in stabilization and mineralization of soil organic matter under ﬁeld
conditions.

The characterization of the tree-soil system with 14C and 15N
represented a complete tracer and total C and N budget. The combination of
traditional component analysis combined with tracers proved to be a powerful
approach in determining the role of the different litters in nutrient cycling
processes. This approach should be broaden to include a diversity of species,
cultural practices and natural systems to fully comprehend the processes

associated with nutrient cycling in ecosystems.

APPENDICES

APPENDIX A
Chapter 4

239

240

Table A.1. The carbon concentration of tree components sampled one year
after the dual labeling procedure on July 19, 1990. Standard error of the
mean shown in parentheses.

 

 

Samplinulate
2-Aug-90 2—Nov-90 25-Jul-91
stem 43.0 (0.6) 38.8 (1.2) 39.3 (0.1)
branches 42.5 (0.5) 39.4 (2.1) 38.9 (0.4)
leaves 40.4 (0.3) 38.0 (0.9) 38.6 (0.5)
root 0-25 cm
< 0.5 mm 27.1 (2.6) 25.8 (2.0) 20.9 (6.6)
.5-1 mm 36.9 (2.0) 29.8 (1.9) 34.2 (2.2)
1-3 mm 38.2 (1.7) 34.4 (1.5) 32.8 (3.8)
3-10 mm 40.3 (0.7) 36.2 (0.0) 34.4 (2.2)
> 10 mm 40.8 (0.1) 35.9 (2.0) 37.4 (0.2)
cutting 41.2 (1.2) 41.1(0.9) 39.2 NA
root 25-60 cm
< 0.5 mm 26.6 (3.6) 20.4 (1.1) 28.2 (4.4)
.5-1 mm 35.3 (2.1) 28.4 (0.7) 27.8 (2.9)
1-3 mm 37.7 (1.8) 31.5 (0.4) 33.4 (2.6)
3-10 mm 37.5 (1.1) 34.7 (0.2) 34.7 (0.1)
> 10 mm 40.4 (0.3) 35.4 (0.8) 34.0 NA
root 60-100 cm
< 0.5 mm 31.8 (1.2) 21.2 (2.0) 30.7 (0.7)
.5-1 mm 36.4 (2.1) 28.5 (1.9) 34.7 (0.4)
1-3 mm 38.8 (1.6) 33.8 (0.4) 32.3 (4.2)
3-10 mm 39.5 (1.1) 36.0 (0.8) 36.6 (0.3)
> 10 mm 43.4 NA 36.8 (1.1) 39.0 NA

 

241

Table A2. Carbon concentration of tree components sampled one year after
the dual labeling procedure on September 5, 1990. Standard error of the
mean shown in parentheses.

Sampling date

 

20-Sep-90 4-Nov-90 5-Sep-91
stem 40.0 (0.8) 39.7 (0.6) 40.2 (0.1)
branches 42.0 (2.0) 40.1 (0.1) 39.0 (0.4)
litter 39.5 (2.2) 38.9 (0.7) 40.3 (0.1)
root 0-25 cm
< 0.5 mm 30.2 (3.0) 27.2 (1.4) 28.4 (3.1)
.5-1 mm 32.1 (3.2) 31.5 (0.5) 33.6 (2.7)
1-3 mm 33.0 (0.4) 33.2 (1.0) 35.7 (0.1)
3-10 mm 36.5 (1.3) 34.0 (1.3) 34.5 (1.4)
> 10 mm 38.2 (1.6) 36.2 (1.3) 35.1 NA
cutting 43.0 (0.6) 40.3 (1.4) 39.4 (0.1)
root 25-60 cm
< 0.5 mm 26.7 (2.9) 24.4 (3.8) 33.7 (0.4)
.5-1 mm 31.9 (1.4) 31.5 (0.2) 35.6 (1.2)
1-3 mm 34.7 (1.4) 34.1(1.1) 36.8 (0.4)
3-10 mm 35.6 (0.7) 37.1 (0.9) 38.0 (1.1)
> 10 mm 36.0 (0.5) 37 .3 (0.3) 38.1(0.5)
root 60-100 cm
< 0.5 mm 25.8 (0.1) 25.6 (4.9) 34.8 (0.6)
.5-1 mm 32.3 (0.8) 30.2 (1.1) 37.1 (0.4)
1-3 mm 37.4 (0.7) 33.4 (1.5) 38.0 (0.0)
3-10 mm 37.6 (0.9) 35.3 (0.7) 37.1 (1.2)
> 10 mm 35.2 NA 35.2 (3.3) 38.6 NA

 

242

Table A.3. Nitrogen concentration of tree components sampled one year after
the dual labeling procedure on July 19, 1990. Standard error of the mean
shown in parentheses.

Sampling gate

 

2-Aug-90 2-Nov-9O 25-Jul-
91
stem 0.43 (0.03) 0.56 (0.02) 0.25 (0.01)
branches 0.59 (0.04) 0.86 (0.05) 0.41 (0.01)
leaves 2.41 (0.09) 1.58 (0.06) 1.74 (0.04)
roots 0-25 cm
< 0.5 mm 0.75 (0.07) 0.61(0.06) 0.40 (0.07)
.5-1 mm 0.75 (0.06) 0.69 (0.05) 0.48 (0.00)
1-3 mm 0.74 (0.06) 0.88 (0.04) 0.46 (0.04)
3-10 mm 0.78 (0.03) 1.13 (0.11) 0.57 (0.02)
> 10 mm 0.58 (0.07) 0.99 (0.02) 0.40 (0.13)
cutting 0.47 (0.02) 0.62 (0.03) 0.28 NA
roots 25-60 cm
< 0.5 mm 0.64 (0.07) 0.42 (0.01) 0.57 (0.11)
.5-1 mm 0.72 (0.07) 0.55 (0.01) 0.50 (0.08)
1-3 mm 0.73 (0.11) 0.68 (0.02) 0.47 (0.03)
3-10 mm 0.77 (0.09) 0.82 (0.01) 0.51(0.04)
> 10 mm 0.68 (0.12) 0.82 (0.02) 0.55 NA
roots 60-100 cm
< 0.5 mm 0.77 (0.06) 0.47 (0.06) 0.57 (0.03)
.5-1 mm 0.83 (0.09) 0.54 (0.03) 0.53 (0.06)
1-3 mm 0.74 (0.06) 0.66 (0.04) 0.53 (0.08)
3-10 mm 0.78 (0.03) 0.90 (0.13) 0.71(0.03)
> 10 mm 1.07 NA 1.04 (0.07) 0.58 NA

 

243

Table A.4. Nitrogen concentration of tree components sampled one year after
the dual labeling procedure on September 5, 1990. Standard error of the
mean shown in parentheses.

—W—

 

20-Sep- 4-Nov-90 5-Sep-91
90
stem 0.48 (0.01) 0.59 (0.02) 0.42 (0.06)
branches 0.87 (0.04) 0.96 (0.05) 0.72 (0.15)
leaves 2.44 (0.03) 2.05 (0.03) 1.97 (0.39)
roots 0-25 cm
< 0.5 mm 0.74 (0.02) 0.75 (0.03) 0.64 (0.17)
.5-1 mm 0.81 (0.08) 0.92 (0.03) 0.82 (0.12)
1-3 mm 0.96 (0.03) 1.16 (0.03) 0.88 (0.22)
3-10 mm 1.07 (0.02) 1.42 (0.06) 0.97 (0.25)
> 10 mm 0.88 (0.01) 1.29 (0.09) 1.25 NA
cutting 0.63 (0.05) 0.77 (0.02) 0.49 (0.15)
roots 25-60 cm
< 0.5 mm 0.71(0.07) 0.57 (0.06) 0.77 (0.15)
.5-1 mm 0.84 (0.05) 0.80 (0.04) 0.86 (0.21)
1-3 mm 0.95 (0.03) 1.03 (0.07) 0.91(0.20)
3-10 mm 1.09 (0.06) 1.22 (0.07) 1.18 (0.32)
> 10 mm 0.95 (0.03) 1.17 (0.11) 1.01(0.29)
roots 60-100 cm
< 0.5 mm 0.64 (0.02) 0.52 (0.11) 1.02 (0.12)
.5-1 mm 0.86 (0.03) 0.73 (0.07) 0.86 (0.13)
1-3 mm 0.90 (0.05) 0.90 (0.07) 1.01(0.28)
3-10 mm 1.06 (0.11) 1.20 (0.06) 1.34 (0.18)
> 10 mm 0.84 (0.03) 1.15 (0.17) 0.87 NA

 

lm.‘ “Anal-r '
E ..— ._ J

APPENDD( A

Chapter 6

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255

Table 3.11. N mineralization results are shown for the last two samplings of
tree labeled July 19, 1990. All soil depths are indicated. Standard error
of the mean shown in parentheses.

 

 

 

Soil depth
0-25 cm 25-60 cm 60-100 cm.
Cumulative Cumulative Cumulative
Days ug N [lg N pg N
Sampled incubated 5’1 soil 1 soil 1 soil
107 days 0 2.3 (0.1) 1.7 (0.2) 0.7 (0.1)
10 4.0 (0.4) 1.4 (0.2) 0.6 (0.3)
20 8.5 (0.5) 1.6 (0.6) 0.9 (0.2)
30 10.9 (2.3) 3.0 (0.9) 1.1 (0.4)
51 16.9 (4.0) 3.7 (0.3) 2.2 (0.4)
61 17.7 (2.9) 2.4 (0.2) 1.3 (0.4)
71 19.8 (2.8) 2.3 (0.1) 1.6 (0.4)
80 26.7 (1.8) 7.3 (0.9) 4.5 (0.7)
122 30.1 (0.8) 3.1 (0.1) 2.2 (0.5)
132 31.1 (1.3) 3.7 (0.2) 2.4 (0.3)
142 31.0 (0.6) 3.6 (0.1) 2.4 (0.5)
372 days 0 3.5 (0.0) 2.4 (0.2) 1.2 (0.3)
10 5.8 (0.2) 1.8 (0.4) 1.6 (0.5)
20 8.3 (0.5) 1.8 (0.4) 1.3 (0.4)
42 14.1(2.1) 2.3 (0.3) 1.6 (0.4)
55 16.2 (0.1) 2.7 (0.2) 1.9 (0.3)
65 22.8 (0.0) 3.8 (0.1) 2.4 (0.5)
75 25.6 (1.0) 4.2 (0.0) 3.0 (0.3)
90 26.3 (1.6) 4.1 (1.4) 2.5 (0.5)
149 43.5 (1.6) 5.1 (0.5) 3.3 (0.9)
159 40.8 (0.4) 6.8 (0.6) 4.8 (0.4)
169 47.0 (1.3) 6.0 (0.9) 4.1 (0.9)

 

256

Table B.12. N mineralization results are shown for the last two samplings of
tree labeled September 5, 1990. All soil depths are indicated. Standard
error of the mean shown in parentheses.

 

 

 

Soil depth
0-25 cm 25-60 cm 60-100 cm.
Cumulative Cumulative Cumulative
Days pg N pg N pg N
sampled incubated g'l soil 1 soil E1 soil
60 days 0 3.8 (0.7) 3.1 (0.5) 1.4 (0.4)
10 6.0 (0.9) 2.9 (0.5) 2.5 (1.2)
20 9.5 (0.4) 3.6 (0.3) 2.2 (0.2)
30 12.5 (1.0) 3.9 (0.3) 2.4 (0.4)
51 19.2 (1.9) 5.6 (0.8) 3.7 (0.5)
61 22.9 (2.6) 5.7 (0.8) 3.5 (0.4)
71 23.7 (2.5) 5.1 (0.7) 3.3 (0.4)
80 25.1 (1.6) 6.0 (0.4) 3.6 (0.3)
122 30.0 (2.1) 6.6 (1.2) 4.3 (0.5)
142 32.3 (2.7) 7.1 (1.7) 4.6 (0.8)
366 days 0 3.1 (0.3) 2.3 (0.5) 0.8 (0.0)
42 17.4 (0.1) 5.5 (0.1) 3.7 (0.0)
61 20.4 (0.4) 3.3 (0.2) 1.4 (0.0)
71 20.2 (0.6) 5.2 (0.2) 2.8 (0.0)
81 28.4 (0.1) 5.0 (0.5) 2.3 (0.1)
96 27.5 (2.2) 4.9 (0.9) 2.5 (0.6)
151 42.1 (0.0) 7.0 (1.2) 3.8 (0.3)

 

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259

Table B.15. N mineralization results are shown from both samplings of
September and July litter exchange experiment. Soil was collected from
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September labeled litter plot July labeled litter
plot
Day 168 Day 168
Sumulative émnulative
Days ug N Days ug N
incubated g1 soil incubated 5-1 soil
0 2.7 (0.2) 0 2.7 (0.0)
10 6.0 (0.1) 10 5.7 (0.1)
20 9.1 (1.0) 20 8.3 (0.3)
55 21.4 (0.8) 55 18.8 (0.8)
70 23.2 (0.4) 70 19.7 (1.0)
80 27.8 (1.2) 80 23.0 (1.6)
90 35.4 (0.8) 90 32.2 (1.2)
101 35.1 (1.0) 101 32.4 (1.2)
143 45.6 (1.7) 143 43.0 (1.3)
153 55.1 (1.7) 153 52.5 (1.3)
163 50.4 (1.9) 163 48.8 (1.5)
Day 328 Day 328

0 4.2 (0.2) 0 4.3 (0.1)
10 8.1 (0.9) 10 8.4 (0.4)
20 12.8 (0.3) 20 15.2 (1.1)
55 20.5 (0.5) 55 21.1 (0.9)
65 26.5 (0.5) 65 27.0 (1.4)
75 31.0 (2.8) 75 29.5 (1.1)
92 34.2 (0.9) 92 34.0 (2.1)
124 36.1 (0.9) 124 37.6 (1.8)

134 53.3 (1.5) 134 50.1 (1.5)

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