FOLIAR UPTAKE OF ATMOSPHERIC REACTIVE NITROGEN POLLUTION
A Dissertation Presented to the Faculty of the Graduate School of Cornell University In Partial Fulfillment of the Requirements for the Degree of Doctor of Philosophy
by Dena M. Vallano August 2009
© 2009 Dena M. Vallano
FOLIAR UPTAKE OF ATMOSPHERIC REACTIVE NITROGEN POLLUTION
Dena M. Vallano, Ph. D. Cornell University 2009 Several studies have established that gaseous reactive N is capable of being incorporated directly into the leaves and assimilated into plant metabolism. However, no studies have directly quantified the magnitude of foliar N uptake at realistic atmospheric N concentrations or how this entry pathway may influence vegetative responses to increasing atmospheric N deposition. This body of work expands on previous research by explicitly quantifying the direct foliar uptake of atmospheric nitrogen dioxide (NO2) using foliar δ15N and the mechanisms driving the capacity for species to incorporate atmospheric reactive N directly through the leaves into biomass over multiple scales in laboratory, mesocosm, and natural systems.
In chapter one, I use foliar δ15N as a tool to quantify the magnitude of foliar NO2 uptake in tomato (Lycopersicon esculentum) and tobacco (Nicotiana tabacum). This chapter establishes that gaseous 15NO2-N may be directly incorporated into the leaves of vegetation and is assimilated as a nutritional source under N-limited conditions. In chapter two, I examine a suite of factors including morphological, stable isotope, and nitrate reductase activity in L. esculentum and N. tabacum to quantify the amount of N incorporated via foliar 15
NO2 uptake, the variation in foliar uptake capacity between species, and
determine whether realistic concentrations of 15NO2 influence plant N metabolism. This chapter provides evidence that the magnitude of foliar NO2
uptake has the potential to influence plant metabolism and that variation exists among species in the capacity to incorporate NO2 via the foliar uptake pathway. In chapter three, I investigate the roles of different root N sources (NO3- and NH4+) on enzyme expression and foliar uptake capacity of 15NO2 in N. tabacum. This chapter contributes to the mechanistic understanding of foliar uptake capacity by integrating relationships among several potential drivers of the foliar uptake pathway. Chapter four examines the variation in natural abundance foliar δ15N patterns and uses these measurements to estimate the importance of soil δ15N, mycorrhizal associations, and foliar N uptake on foliar δ15N patterns. This chapter supports the use of natural abundance foliar δ15N to detect the potential for foliar N uptake in several tree species in temperate forests.
BIOGRAPHICAL SKETCH Dena Vallano was born on 9 February 1981 in Greensburg, Pennsylvania. Dena grew up spending much of her childhood playing in the fields behind her house until they were plowed over to make room for a subdivision. During this time, she fell in love with nature and basketball, never daring to choose between the two.
In 1999, Dena traveled across the state to Philadelphia to pursue her BS in Honors Biology at Villanova University. She began her studies with aspirations of becoming a veterinarian, shifted her focus to bioethics, and eventually took a course in Introductory Ecology that reignited her childhood love and passion for learning about nature. During her sophomore year, she received a Howard Hughes Medical Institute Fellowship to work in the lab of Dr. Joseph Orkwiszewski. This experience completely changed her life and introduced her to the wonderful world of laboratory research and plant physiology. While at Villanova, she spent a semester abroad in Melbourne, Australia, an experience that cemented her passion for international travel and further reinforced her desire to study ecosystem ecology. Dena eventually completed her senior honors thesis research examining the effects of the plant hormones kinetin and gibberillic acid on shoot apical meristem development in corn. The mentorship and friendship Dena received while working for Dr. Orkwiszewski was the inspiration behind her decision to pursue graduate studies.
She decided to combine her love for plant physiology and ecology and pursue a graduate degree in Ecology and Evolutionary Biology at Cornell University. She joined the lab of Dr. Jed Sparks and began her journey into the iii
fascinating underworld of direct leaf incorporation of atmospheric reactive nitrogen, which became the topic of her doctoral dissertation. While at Cornell, she became immersed in the field of stable isotope ecology and pretty handy along the way. When not working on her project, Dena had opportunities to work in many amazing locations (Trinidad, Hawaii, etc.) in fields other than her own, which kept her sane and broadened her scientific horizons. She also participated in several outreach and educational opportunities which were incredibly rewarding and memorable experiences.
Since defending her dissertation, Dena has moved to Santa Cruz for a postdoctoral position with Dr. Erika Zavaleta at the University of California Santa Cruz.
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To my mother, Linda.
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ACKNOWLEDGMENTS
I cannot express how thankful I am to all of the people that have supported me professionally and personally throughout my experience at Cornell. It is with sincere appreciation and pleasure that I acknowledge them now.
First and foremost, I thank my family. They have given me so much unconditional love throughout my life and in all of my endeavors. My mother, Linda, has always encouraged me to pursue my dreams and believed in me no matter where life has taken me. She is an inspiration and a hero to me in so many ways. My sisters, Elissa and Talia, have always kept me grounded and been my biggest cheerleaders. Their support and occasional field assistance has been pivotal throughout this process. My Aunt Mary Lou has also been a tremendous source of comfort and support during my time at Cornell. I love you all.
Iʼd like to thank my graduate committee for their intellectual support and dedicated mentorship that has shaped me into the scientist that I am today. All three contributed to my experience at Cornell in unique ways and fundamentally refined the way I view science and greatly expanded my perspectives and ability to think critically and clearly about my research. Dr. Jed Sparks has been steadfastly supportive and integral to my development as a scientist over the years. I am grateful for all the impromptu meetings, both during the week and on the weekends, as well as the last minute feedback on countless grants, proposals, talks, and manuscripts. I am especially thankful for the laughter and many unforgettable experiences that Iʼve had with him
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over the years – many of which cannot be retold here. Dr. Thomas Brenna broadened my understanding of stable isotope techniques and taught me how important it is to not only understand the output of your data but also the process behind getting those numbers. Dr. Thomas Owens is one of the best critical thinkers and mentors that I have encountered during both my undergraduate and graduate careers. His advice and insight were invaluable to me throughout my dissertation and I am truly thankful for the time that he has spent teaching me to think critically about my research.
Without a doubt, I would like to thank my undergraduate thesis advisor at Villanova, Dr. Joseph Orkwiszewski. He has been my science “rock”, without which I would certainly be a different person and scientist than I am today. His unconditional faith in my abilities and character has encouraged me to always strive for the best and realize that I deserve it. He has been an advisor, a mentor, and a friend over the years. I am forever grateful for his role as my “Dutch Uncle”.
Many other people contributed to my research over the years: Kim Sparks for her organizational prowess and endless knowledge of all things plant ecophysiology, Art Kasson for running gazillions of isotope samples for me on very short notice, Gary Oltz for all his handiwork, and Paul Cooper for his greenhouse expertise. I thank my lab members for their guidance, support, and advice on my crazy presentation color choices: Allyson Eller, Carrie McCalley, Danica Lombardozzi, Dave Baker, Jocelyn Lavallee and Dr. Brian Strahm (post-doc extraordinaire). Gary Lovett, Steve Thomas, and Eric Hazelton have also provided advice and assistance on many aspects of my vii
dissertation work. I am also incredibly fortunate to have been a part of two amazing communities while at Cornell, the Department of Ecology and Evolutionary Biology and the Program in Biogeochemistry and Environmental Complexity. The faculty and graduate students in these groups have made my time here so socially rewarding and intellectually stimulating the entire time. I look forward to many more years of friendship with all of you.
I also would like to acknowledge the steadfast and amazingly efficient support of the administrative and research staff of the Department of Ecology and Evolutionary Biology. I am especially indebted to Alberta Jackson, LuAnne Kenjerska, Dee Dee Albertsman, Carol Damm, Janeen Orr, Brian Mlodzinski, and John Howell over the years.
It is with great pleasure that I thank the unseen family members that I have gained along this amazing journey. The girls – and now women – that became my family at Villanova are a blessing that I feel honored to have in my life. Finally, I am so grateful for the friends I have made at Cornell over the years. I am truly lucky to have encountered such a dynamic, vivacious, intelligent group of graduate students that have provided countless hours of laughter until I was crying and encouraged copious amounts of wine drinking. They made this experience unforgettable and have been such an enormous source of joy to me. Youʼve have made me a better person in so many, countless, ways. Thank you.
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TABLE OF CONTENTS
BIOGRAPHICAL SKETCH ……...…………….....………….......…………………iii DEDICATION …...................………....…………….…………………………….... v ACKNOWLEDGEMENTS…………....…………….…………………………….... vi LIST OF FIGURES …………………………...…………………………………… xi LIST OF TABLES ……………………………………………….........…………... xiii PREFACE ................................…………………….........……………………… xiv CHAPTER 1. FOLIAR δ15N VALUES AS INDICATORS OF FOLIAR UPTAKE OF ATMOSPHERIC NITROGEN POLLUTION ……..….......1 Abstract ......………………………………………………………………….………1 Introduction ..……………………………………………...…………….…...……...2 Nitrogen sources to terrestrial vascular plants ..………………..………...……..6 Foliar assimilation of atmospheric nitrogen pollutants ..……………...............12 Contribution of foliar uptake to total plant nitrogen assimilation ......…………14 Foliar δ15N as a tool to determine the magnitude of foliar nitrogen uptake ………..…………………………………………...….................16 Current challenges for using foliar δ15N as a tool to determine the magnitude of foliar nitrogen uptake ………………..………………....…….18 Promising methods using foliar δ15N as an indicator of direct leaf nitrogen assimilation ……..……………..………………...…………...…….19 Future research ……………..………………..…………………...………………26 Conclusions ……………..…………………………………………...………........29 References ....................................................................................................30 CHAPTER 2. QUANTIFYING FOLIAR UPTAKE OF GASEOUS NITROGEN DIOXIDE USING ENRICHED FOLIAR δ15N VALUES ....................…..….......39 Abstract ........…………………………………...…………………………….……39 Introduction..……………………………………………...……………......……...40 Materials and methods ....................................………………..………...……..44 Results ..........................................................................……………...............49 Foliar uptake and N source partitioning ........................................…………49 Morphology and biomass allocation ……………………...……...…..............52 Nitrate reductase activity ......................……………..…...…....….........…….56 Discussion .....................................................................................................58 References ....................................................................................................65 CHAPTER 3. INFLUENCE OF DIFFERENT ROOT NITROGEN SOURCES ON THE CAPACITY FOR DIRECT FOLIAR NO2 UPTAKE AND ENZYME EXPRESSION IN NICOTIANA TABACUM......................................…..….......70 Abstract ........…………………………………...…………………………….……70
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Introduction..……………………………………………...……………......……...71 Materials and methods ....................................………………..………...……..75 Results ..........................................................................……………...............80 Foliar uptake and N source partitioning ........................................…………80 Morphology and biomass allocation ……………………...……...…..............82 Enzyme activity.....................................……………..…...…....….........…….85 Discussion .....................................................................................................88 CHAPTER 4. PARTITIONING THE RELATIVE INFLUENCE OF SOIL N, MYCORRHIZAL ASSOCIATIONS, AND DIRECT FOLIAR NITROGEN UPTAKE ON FOLIAR δ15N PATTERNS: CAN WE DETECT FOLIAR UPTAKE OF REACTIVE NITROGEN? ...........................................................…..….....104 Abstract ........…………………………………...…………………………….…104 Introduction ..……………………………………………...……………......…..105 Materials and methods ....................................………………..………...…..109 Results ..........................................................................…………….............115 Potted plant mesocosm experiment ..............................................………115 Field sampling ..................................……………………...……...…............116 Discussion ...................................................................................................124
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LIST OF FIGURES Figure 1.1. Primary source pools and uptake pathways in the N cycle.............8 Figure 1.2. Primary metabolic processes involved in the foliar uptake and assimilation of atmospheric ammonia and nitrogen dioxide ..........................13 Figure 1.3. Design of an individual hydroponics fumigation system used for seedlings for long term experiments..............................................................21 Figure 1.4. Proportion (%) of total N content attributable to the direct foliar uptake of 15NO2-N in L. esculentum ..............................................................22 Figure 1.5. Proportion (%) of total N content attributable to the direct foliar uptake of 15NO2-N in N. tabacum and L. esculentum ....................................24 Figure 2.1. Proportion (%) (a), and amount (in mg) (b) of total N content attributable to the direct foliar uptake of 15NO2-N in N. tabacum and L. esculentum for each treatment ......................................................................50 Figure 2.2. Amount (in mg) of total N content attributable to the direct foliar uptake of 15NO2-N in N. tabacum (a) and L. esculentum (b) for root, shoot and leaf tissue for each treatment .................................................................52 Figure 2.3. Total plant biomass (a) and root to shoot ratio (b) of N. tabacum and L. esculentum over all treatments ...........................................................54 Figure 2.4. Carbon to nitrogen ratio for each treatment in N. tabacum and L. esculentum ................................................................................................55 Figure 2.5. Foliar δ13C across treatments in N. tabacum and L. esculentum .56 Figure 2.6. Leaf (a) and root (b) NR activity for each treatment in N. tabacum and L. esculentum .........................................................................................57 Figure 2.7. Leaf NR activity versus mg N incorporated from foliar 15NO2 uptake in L. esculentum and N. tabacum ......................................................58 Figure 3.1. Amount (in mg) of total N content attributable to the direct foliar uptake of 15NO2-N in N. tabacum and partitioning of N in root, shoot and leaf tissue for each treatment ...............................................................................81
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Figure 3.2. Total plant biomass (a) and root to shoot ratio (b) in N. tabacum for each treatment..........................................................................................83 Figure 3.3. Carbon to nitrogen ratio (a) and foliar δ13C (b) in N. tabacum......84 Figure 3.4. Root (a) and leaf (b) NR activity in N. tabacum ............................86 Figure 3.5. Root (a) and leaf (b) GS activity in N. tabacum ............................87 Figure 3.6. Relationship between leaf NR activity versus mg N incorporated via foliar 15NO2 uptake (a) and leaf GS activity and mg N incorporated from 15 NO2 uptake (b) in N. tabacum .....................................................................88 Figure 4.1. Site locations in New York State ................................................109 Figure 4.2. Relationship between ambient atmospheric NO2 concentration (a) and bulk soil δ15N (b) with foliar δ15N in red maple seedlings .....................115 Figure 4.3. Relationship between ambient atmospheric NO2 concentration and foliar δ13C (a) and foliar N content (b) in red maple seedlings with and without mycorrhizal associations .................................................................117 Figure 4.4. Relationship between Δ15N and ambient atmospheric NO2 concentration in adult trees .........................................................................118 Figure 4.5. Relationship between foliar δ15N and bulk soil δ15N (a, c, e, g) and ambient atmospheric NO2 concentration (b, d, f, h) in adult trees ...............120 Figure 4.6. Partial residual plots for the predictor variables included in the multiple regression model explaining variance in foliar δ15N patterns .........121 Figure 4.7. Relationship between foliar δ15N and ambient atmospheric NO2 concentration (a) and bulk soil δ15N (b) in red maple trees .........................122 Figure 4.8. Relationship between ambient atmospheric NO2 concentration and foliar δ13C (a) and foliar N content (b) in adult trees ..............................123
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LIST OF TABLES
Table 4.1. Site descriptions ..........................................................................110
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PREFACE
Nitrogen (N) is central to the molecular structures of proteins, nucleic acids, and enzymes that regulate metabolic processes in all living organisms. Traditionally, plants are thought to exclusively access reactive N via root uptake of ammonium (NH4+), nitrate (NO3-), and dissolved organic N from the soil pool. Most of the incorporated reactive N is assimilated in the roots and transported to the shoots in the form of amino acids. These compounds are used in a variety of metabolic processes within the plant that are necessary for nutrition, growth, and photosynthesis. And, in many terrestrial ecosystems, reactive N is often the critical element that limits plant growth. Therefore, the relevance of atmospheric reactive N deposition in ecosystems became increasingly pivotal during the latter half of the 20th century and continues to drive research in the fields of plant ecophysiology, biogeochemistry, and stable isotope ecology (Galloway et al. 2004, Hill 1971, Nussbaum et al. 1999).
The dramatic increase in atmospheric concentrations of reactive N oxides (NOy) from human activities (car exhausts, factory emissions, etc.) has rapidly increased the amount of reactive N deposition to vegetation (Vitousek 1994). Subsequently, the traditional view that plants incorporate reactive N exclusively through the root uptake pathway was further examined, as this additional N source of atmospheric reactive N became increasingly available to plants. It was discovered that these pollutant compounds were, in fact, being deposited and eventually assimilated into vegetation via two interrelated pathways: directly via foliar uptake and indirectly via the root uptake
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pathway (Gessler et al. 2002, Hanson & Lindberg 1991). In the foliar uptake pathway, gaseous reactive N compounds enter the leaf through the stomata, are rapidly dissolved in the apoplast, and are eventually metabolized. Thus, it is intriguing to ask how this uptake pathway impacts plant physiological processes through influences on growth, photosynthesis, and the plant N budget and further examine the subsequent potential implications on the structure, function and composition of terrestrial ecosystems.
Nitrogen isotope ratios (15N:14N) became a practical tool for scientists to investigate multiple N sources to plant metabolism, estimate the amount of atmospheric N compounds in an ecosystem, and identify the roles of the various uptake pathways into a plant (Robinson 2001, Robinson et al. 1998). Stable nitrogen isotopes are especially useful for these applications because the isotopic composition (δ15N) of pollutant compounds may considerably differ from the natural N δ15N in the soil. For instance, if the leaf incorporated N source has a significantly different δ15N compared to the root incorporated N source, then the foliar δ15N value will represent both sources and can be used to quantify the proportion of each N source assimilated into plant metabolism. Therefore, the importance of various N sources for plant metabolism could be determined at the leaf level (Ammann et al. 1999, Muller et al. 1996). Researchers began to use stable isotope techniques to investigate the linkage between gaseous N deposition and N cycling in plants and as a tool to trace the movement and pools of pollutant N compounds within plants (Dawson et al. 2002).
Several studies established that gaseous reactive N was capable of being
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incorporated directly into the leaves and assimilated into plant metabolism (Boyce et al. 1996, Okano et al. 1988, Siegwolf et al. 2001, Wellburn 1990). Additional work yielded limited insight to the variation in gaseous reactive N assimilation existing across and within species (Gessler et al. 1998, Morikawa et al. 1998, Sparks et al. 2003) and the processes that potentially govern the ability to incorporate reactive N directly through the leaves (Eller & Sparks 2006, Ramge et al. 1993). However, to date, few studies have successfully used this tool to examine the magnitude of leaf N uptake at realistic atmospheric N concentrations or how this entry pathway may influence vegetative responses to increasing human-caused N deposition. Further, the mechanisms influencing species variation in the capacity to directly assimilate reactive N via the foliar uptake pathway have not been rigorously examined.
This body of work expands on previous research by explicitly quantifying the direct foliar uptake of atmospheric nitrogen dioxide (NO2; a common gaseous pollutant) using foliar δ15N measurements and the mechanisms driving the capacity for various species to incorporate atmospheric reactive N pollutants directly through the leaves into biomass over multiple scales in laboratory, mesocosm, and natural systems.
Chapter one examines the use of foliar δ15N values as a tool to quantify the magnitude of foliar NO2 uptake and the potential role of the foliar N uptake pathway in N metabolic processes in two herbaceous species, tomato (Lycopersicon esculentum) and tobacco (Nicotiana tabacum). Using a customdesigned hydroponics-fumigation system, both species were exposed to various concentrations of enriched gaseous 15NO2 to the shoot systems and
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NO3- to the root systems. Exposure to and assimilation of 15NO2 into plant tissue stimulated whole-plant biomass production in both species. Both species were capable of incorporating up to 15% of the total assimilated N into biomass directly via the foliar uptake pathway. No significant variation in foliar uptake capacity was observed between species at the whole-plant level. This chapter establishes that gaseous 15NO2-N may be directly incorporated into the leaves of vegetation, assimilated as a nutritional source under N-limited conditions, and may be successfully quantified using foliar δ15N measurements in a laboratory setting.
Chapter two examines a suite of factors including morphological, stable isotope, and nitrate reductase activity in N. tabacum and L. esculentum to quantify the amount of N incorporated via foliar 15NO2 uptake, the variation in foliar uptake capacity between species, and determine whether realistic concentrations of 15NO2 influence plant N metabolism. N. tabacum and L. esculentum assimilated 15 and 11%, respectively, of 15NO2-N into total biomass via the foliar uptake pathway under low NO3- supply. Neither L. esculentum nor N. tabacum linearly increased total biomass accumulation (in g DW) in response to NO2 exposure. Both species exhibited differences in the ratio of root to shoot (stem + leaf) tissue in response to differences in root NO3supply. There were no strong effects on leaf δ13C values, or leaf and root nitrate reductase activity (NRA) in response to 15NO2 exposure in either species. Despite only small amounts of variation in NRA, a positive relationship existed between leaf NRA and the amount of NO2-N incorporated via the foliar uptake pathway in both L. esculentum and N. tabacum. This chapter provides further evidence that foliar uptake is a significant pathway of
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NO2 entry, the magnitude of foliar NO2 uptake has the potential to influence plant metabolism, and that some variation exists among species in the capacity to incorporate NO2 via the foliar uptake pathway.
Chapter three investigates the roles of different root N sources (NO3- and NH4+) on enzyme expression and the capacity for foliar uptake of gaseous enriched 15NO2 in N. tabacum. The proportion and amount of NO2-N incorporated into biomass via direct foliar uptake was not influenced by differences in the root N source, suggesting that the overall rate of foliar NO2 uptake is not strongly impacted by the form of N supplied to the root system in N. tabacum. When exposed to gaseous NO2, plants grown solely on NO3- or NH4+ incorporated 12.0 ± 0.6 and 12.4 ± 0.6% of total plant N through foliar uptake, respectively. N. tabacum did not increase total biomass accumulation (in g DW) in response to NO2 exposure, regardless of root N supply. Fumigated plants grown under NO3- supply showed a decrease in the ratio of root to shoot (stem + leaf) tissue compared to non-fumigated plants, but this result was not observed in fumigated plants grown under NH4+ supply. Interestingly, we found altered enzyme activities associated with availability of different root N sources and NO2 exposure. Overall, leaf NR activity decreased and leaf GS activity increased in plants exposed to NO2, regardless of the root N source. Results suggest that plant performance and foliar uptake capacity are independent of the root N form and that enzyme expression within the primary N reduction pathways may be correlated with foliar uptake capacity in N. tabacum. This chapter contributes to the mechanistic understanding of direct foliar uptake capacity by integrating relationships among several potential drivers of the foliar uptake pathway.
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Chapter four examines the variation in natural abundance foliar δ15N patterns among several dominant hardwood tree species and uses these measurements to estimate the relative importance of soil δ15N, mycorrhizal associations, and foliar N uptake on foliar δ15N patterns in response to different levels of atmospheric reactive N deposition in temperate forest ecosystems. Using a potted plant mesocosm system, the relationship between foliar δ15N, soil δ15N and ambient atmospheric N concentration was determined while controlling for mycorrhizal associations in red maple (Acer rubrum) seedlings. Along the same temporal and spatial scales, foliar and soil δ15N patterns were determined in adult stands of several dominant hardwood species. The potential influence of direct foliar N uptake, soil δ15N, and mycorrhizae on foliar δ15N patterns were examined in the following species: American beech (Fagus grandifolia), black birch (Betula lenta), red maple (A. rubrum), and red oak (Quercus rubra). Significant relationships existed between soil δ15N, foliar δ15N, and foliar N content versus NO2 concentration in red maple seedlings across sites. Soil δ15N was the dominant driver of foliar δ15N patterns in red maple seedlings, regardless of mycorrhizal treatment. In adult tree species, there were significant correlations between foliar δ15N and both soil δ15N and NO2 concentration across sites. Using a multiple regression model, NO2 concentration explained 0%, 69%, 23%, and 45% of the residual variation in foliar δ15N remaining in American beech, red maple, red oak, and black birch, respectively, after accounting for soil δ15N. This chapter supports the use of natural abundance foliar δ15N to characterize the influence of soil δ15N, mycorrhizal fractionation, and foliar N uptake on plant N status in temperate forests.
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Overall, these studies demonstrate the importance of the foliar N uptake pathway on plant metabolic processes and represent the first quantification of the magnitude of direct foliar uptake of gaseous NO2 pollution in vegetation. Furthermore, this work indicates that if plants are able to assimilate significant quantities of gaseous N pollutants and incorporate them into organic compounds, direct foliar uptake of atmospheric N pollution may play a crucial role in the global N and carbon cycles through the alteration of photosynthesis and the removal of N pollutants from the atmosphere.
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REFERENCES Ammann M., Siegwolf R., Pichlmayer F., Suter M., Saurer M. & Brunold C. (1999) Estimating the uptake of traffic-derived NO2 from 15N abundance in Norway spruce needles. Oecologia, 118, 124-131. Boyce R.L., Friedland A.J., Chamberlain C.P. & Poulson S.R. (1996) Direct canopy nitrogen uptake from N-15-labeled wet deposition by mature red spruce. Canadian Journal of Forest Research-Revue Canadienne De Recherche Forestiere, 26, 1539-1547. Dawson T.E., Mambelli S., Plamboeck A.H., Templer P.H. & Tu K.P. (2002) Stable isotopes in plant ecology. Annual Review of Ecology and Systematics, 33, 507-559. Eller A. & Sparks J.P. (2006) Predicting leaf-level fluxes of ozone and nitrogen dioxide: the relative roles of diffusion and biochemical processes. Accepted in Plant, Cell, and Environment. Galloway J.N., Dentener F.J., Capone D.G., Boyer E.W., Howarth R.W., Seitzinger S.P., Asner G.P., Cleveland C.C., Green P.A., Holland E.A., Karl D.M., Michaels A.F., Porter J.H., Townsend A.R. & Vorosmarty C.J. (2004) Nitrogen cycles: past, present, and future. Biogeochemistry, 70, 153-226. Gessler A., Rienks M. & Rennenberg H. (2002) Stomatal uptake and cuticular adsorption contribute to dry deposition of NH3 and NO2 to needles of adult spruce (Picea abies) trees. New Phytologist, 156, 179-194. Gessler A., Schneider S., Von Sengbusch D., Weber P., Hanemann U., Huber C., Rothe A., Kreutzer K. & Rennenberg H. (1998) Field and laboratory experiments on net uptake of nitrate and ammonium by the roots of spruce (Picea abies) and beech (Fagus sylvatica) trees. New Phytologist, 138, 275-285. Hanson P.J. & Lindberg S.E. (1991) Dry deposition of reactive nitrogencompounds - a review of leaf, canopy and non-foliar measurements. Atmospheric Environment Part A-General Topics, 25, 1615-1634. Hill A.C. (1971) Vegetation: a sink for atmospheric air pollutants. Journal of the Air Pollution Control Association, 21, 341.
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Morikawa H., Higaki A., Nohno M., Takahasi M., Kamada M., Nakata M., Toyohara G., Okamura Y., Matsui K., Kitani S., Fujita K., Irifune K. & Goshima N. (1998) More than a 600-fold variation in nitrogen dioxide assimilation among 217 plant taxa. Plant Cell and Environment, 21, 180-190. Muller B., Touraine B. & Rennenberg H. (1996) Interaction between atmospheric and pedospheric nitrogen nutrition in spruce (Picea abies L Karst) seedlings. Plant Cell and Environment, 19, 345-355. Nussbaum S., Ammann M. & Fuhrer J. (1999) Foliar absorption and use of airborne oxidized nitrogen by terrestrial plants. In: Nitrogen nutrition and plant growth (eds H. Srivastava & R. Singh), pp. 103-172. Science Publishers, Inc., Enfield, NH. Okano K., Machida T. & Totsuka T. (1988) Absorption of atmospheric NO2 by several herbaceous species - estimation by the N-15 dilution method. New Phytologist, 109, 203-210. Ramge P., Badeck F.W., Plochl M. & Kohlmaier G.H. (1993) Apoplastic antioxidants as decisive elimination factors within the uptake process of nitrogen-dioxide into leaf tissues. New Phytologist, 125, 771-785. Robinson D. (2001) δ15N as an integrator of the nitrogen cycle. Trends in Ecology & Evolution, 16, 153-162. Robinson D., Handley L.L. & Scrimgeour C.M. (1998) A theory for 15N/14N fractionation in nitrate-grown vascular plants. Planta, 205, 397-406. Siegwolf R.T.W., Matyssek R., Saurer M., Maurer S., Gunthardt-Goerg M.S., Schmutz P. & Bucher J.B. (2001) Stable isotope analysis reveals differential effects of soil nitrogen and nitrogen dioxide on the water use efficiency in hybrid poplar leaves. New Phytologist, 149, 233-246. Sparks J.P., Roberts J.M. & Monson R.K. (2003) The uptake of gaseous organic nitrogen by leaves: A significant global nitrogen transfer process. Geophysical Research Letters, 30, 2189-2193. Vitousek P.M. (1994) Beyond global warming - ecology and global change. Ecology, 75, 1861-1876.
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Wellburn A.R. (1990) Why are atmospheric oxides of nitrogen usually phytotoxic and not alternative fertilizers? New Phytologist, 115, 395429.
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CHAPTER 1: FOLIAR δ15N VALUES AS INDICATORS OF FOLIAR UPTAKE OF ATMOSPHERIC NITROGEN POLLUTION*
Abstract
Vegetation is a significant sink for atmospheric nitrogen (N) pollutants in terrestrial ecosystems and, in N-limited systems, the foliar uptake of reactive N may represent a considerable source of N to plants. However, little research has examined the variation in foliar N assimilation among species at realistic concentrations or how this assimilation may influence foliar δ15N values. An understanding of the magnitude of foliar N uptake and whether foliar δ15N values may be used to track this assimilation of anthropogenic N is imperative for understanding the future N cycle. In the following chapter, I discuss several of the more promising laboratory and field efforts using δ15N to quantify foliar N uptake. In particular, I have used combined hydroponics-fumigation systems to address several of the unresolved questions associated with the foliar uptake of reactive N. Using this system, I evaluated tomato (Lycopersicon esculentum) and tobacco (Nicotiana tabacum), which contrast in the reduction site for metabolism, under low or high (50μM and 500μM) root NO3- supply and exposed shoot systems to filtered or 15NO2-enriched air (0 ppb, 20 ppb,
*Vallano D.M. and J.P. Sparks. 2007. Foliar δ15N values as indicators of foliar uptake of atmospheric nitrogen pollution. In Dawson T. and Siegwolf R. (eds) Isotopes as Indicators of Ecological Change, Elsevier Academic Press. 93109.
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and 40 ppb) for four weeks. A key finding from this work is that both species are capable of incorporating up to 15% of their total assimilated N using the foliar uptake pathway, although no significant variation in foliar uptake capacity between species was observed at the whole-plant level. Results indicate that NO2 pollution may be used as a nutritional source under N-limiting conditions and may be traced through foliar δ15N values.
Introduction
Among the cycles of elements important to life, the nitrogen (N) cycle has been the most perturbed beyond its natural state by human activities (Hopkins et al. 1998, Vitousek 1994, Vitousek et al. 1997). One manifestation of this perturbation is a dramatic increase in the amount of reactive N in the atmosphere. Anthropogenic emissions of atmospheric reactive N now exceed natural emissions and are the predominant source of atmospheric reactive N deposition to terrestrial ecosystems. Asia, Europe, and North America account for the majority of the current anthropogenic emissions and the highest levels of deposition. However, substantial increases in atmospheric reactive N are projected over the next 50 years for several other regions of the world as industrialization increases (Galloway et al. 2004, Phoenix et al. 2006). It is crucial that we improve our understanding of the potential sinks for reactive N as global inputs continue to increase.
Vegetation appears to be a significant sink for atmospheric reactive N and the direct foliar uptake of atmospheric N pollution may represent a considerable
2
source of N to plants (Hanson & Lindberg 1991, Hill 1971, Rondon & Granat 1994, Rondon et al. 1993). Plants are capable of assimilating the N in gaseous N compounds, including the reactive N oxides (“NOy”, including NO, NO2, HNO3 and organic nitrates) and ammonia gases (“NHx”, including NH3 and particulate NH4+), and incorporating this N into organic compounds for growth (Gessler et al. 2002, Morikawa et al. 1998). The capacity for foliar uptake of reactive N is likely influenced by plant species characteristics and reactive N concentrations in the atmosphere (Morikawa et al. 1998). In this chapter, we review the potential of N stable isotopes, specifically foliar δ15N values, as an innovative and important tool for determining the magnitude and ultimate fate of atmospheric reactive N deposition to foliage globally.
Reactive N generation and redistribution globally is altering ecosystem function and the composition of natural plant assemblages both through growth-stimulating and phytotoxic effects (Bobbink & Roelofs 1995, Lea 1998, Pearson & Soares 1995, Vitousek et al. 1997, Wellburn 1990). Increasing the total reactive N input to an ecosystem can lead to several negative effects on ecosystem function such as soil and water acidification, increased loss of nutrients from the soil, changes to competitive interactions among plants, reductions in biodiversity, increased vulnerability of invasion, and nutrient imbalances in vegetation (Asner et al. 1997, Bobbink et al. 1998, Gilliam 2006). Moreover, plant performance can also be deleteriously affected through reductions in net photosynthesis, respiration, stomatal conductance, enzyme activities, and growth (Takahashi et al. 2005). Positive effects of increased reactive N deposition may include fertilization of N-limited ecosystems promoting increased carbon sequestration and plant growth (Beedlow et al.
3
2004). Although numerous studies have shown the impacts of total N deposition on ecosystem processes (Aber et al. 2003, Fahey et al. 1999, Galloway et al. 2003, Lovett 1994, Wesely & Hicks 2000), few have investigated the magnitude or influence of atmospheric N pollution assimilated directly by foliage (Sparks et al. 2001).
The ability of vegetation to incorporate N into plant metabolism from both leaves and roots suggests the fate of N deposited to an ecosystem may be fundamentally different depending upon the entry pathway. For example, the chemical products of foliar N assimilation (most commonly NO3-) are plant nutrients and are directly incorporated from the leaf apoplast and used for growth (Gessler et al. 2000, Morikawa et al. 1998). In contrast, N additions to the soil surface during deposition may be incorporated into microbial biomass, revolatilized, or leached. Furthermore, additional N to the soil surface may have other effects (e.g., changes in soil fertility, soil acidification, stripping of base cations, etc.). Thus, the direct entry of N through leaves is intriguing in that it could fundamentally change plant productivity through a direct influence on photosynthesis without the concomitant changes in soil chemistry often observed under high levels of soil N additions. However, it is still unclear if, and how, plant performance differs under N deposition to the soil (i.e., fundamentally altering the reactive N supply to root systems) versus changes in N additions from the atmosphere (i.e., changes in the magnitude of foliar uptake). Both forms of N entry into an ecosystem will likely increase together under most scenarios of intensified industrialization. However, various forms of gaseous reactive N have significantly different atmospheric lifetimes, suggesting plant communities arrayed across the landscape experience not
4
only unequal total N deposition, but different proportions of N through the root and foliar pathways.
There is some evidence to suggest the proportion of N incorporated through the foliar versus soil pathway may vary by environment. For instance, atmospheric reactive N is dominated by relatively reduced forms such as nitrogen dioxide (NO2) in industrialized areas. These compounds deposit more readily through the leaf stomata than to other surfaces in the environment (Sparks et al. 2001). At distances further from sources of N pollution, much of the reactive N is further oxidized to organic forms such as peroxyacetyl and isoprene nitrate. These compounds also readily enter plant stomata (Sparks et al. 2003) at a rate higher than deposition to other surfaces. In regions still further from sources of reactive N, the air mass is often dominated by nitric acid (HNO3-). Nitric acid is very soluble in water and tends to deposit readily to many surfaces in the environment at a rate much faster than entry through the stomata (Byterowicz et al. 1998). Since the composition of atmospheric reactive N gases can vary across the landscape and are more or less likely to enter plant stomata, terrestrial ecosystems may receive differential amounts of N through the foliar uptake pathway.
Nitrogen stable isotope ratios (15N/14N) are a useful tool for estimating the amount of anthropogenic N in the system and quantifying the uptake pathway into the plant [e.g., through soil NO3- or NH4+, dissolved organic N, foliar uptake via stomata, cuticular diffusion, etc. (Robinson et al. 1998, Robinson et al. 2000)] because forms of N available to plants often have varying isotopic compositions (Evans 2001, Kolb & Evans 2002). This may be especially
5
relevant to industrialized, urban regions where pollutant compounds contribute a significant amount of N input via dry and wet deposition to ecosystems. Consequently, the difference in source isotopic composition can provide a natural tracer for the assessment of foliar uptake and assimilation of reactive N pollution in terrestrial ecosystems (Handley & Raven 1992, Handley & Scrimgeour 1997). Specific to foliar uptake, if the foliar incorporated source is significantly different from the soil source, then the foliar δ15N value will be representative of both sources and can be used to quantify the proportion of each N source incorporated into plant biomass. If plants are able to assimilate significant quantities of gaseous N pollutants and incorporate them into organic compounds, foliar δ15N will be an important tool to examine the role foliar uptake of atmospheric N pollution plays in the global N and carbon cycles through the alteration of photosynthesis and the removal of N pollutants from the atmosphere.
In this chapter, I discuss the use of foliar δ15N values as a tool for determining the magnitude and impact of gaseous reactive N directly entering the foliage of vegetation. Although not an exhaustive treatment, I place this discussion within the context of the causative factors defining foliar δ15N values. I also describe the current understanding of foliar δ15N values to provide a background upon which foliar uptake of reactive N can be considered.
Nitrogen sources to terrestrial vascular plants
The N sources assimilated by plants often have different isotopic compositions and many studies have used foliar δ15N values to indicate the δ15N of the
6
dominant N source used by a plant and as a measure of N sources from both soil and atmospheric deposition (Evans 2001). However, we now know that foliar δ15N values represent the integration of a range of processes (Evans 2001, reviewed by Handley & Scrimgeour 1997, Robinson 2001, Stewart et al. 2002) and can vary by as much as ±10‰ in co-existing species (Handley & Scrimgeour 1997). The presence of multiple N sources with distinct isotopic values, differential processing of soil N pools (e.g. temporal and spatial patterns of net mineralization and nitrification), temporal and spatial variation in N availability, mycorrhizal interactions, and alterations in plant N demand can all influence foliar δ15N (Figure 1.1). Identifying and differentiating the sources of N available to plants is necessary for a variety of ecological questions including accurately interpreting foliar δ15N values. Further, additional understanding of how specific N species deposited to terrestrial ecosystems impact foliar δ15N values will provide us a tool to link plant physiological responses, terrestrial ecosystem function, and atmospheric chemistry. In this section, we briefly discuss various sources of plant-available N and their potential influences on foliar δ15N.
Soil nitrogen sources
Plants derive N primarily from the soil in the form of ammonium (NH4+), nitrate (NO3-), and, secondarily, from organic N. Plant available soil N originates from decomposition of organic matter, biological N fixation, additions of N in organic or inorganic fertilizers, and deposition originating from fossil fuel combustion and animal production (Hopkins et al. 1998). There is enormous variability in the isotopic composition of the soil N pool available to plants. For example,
7
Figure 1.1. Primary source pools and uptake pathways in the N cycle. Boxes represent pools and arrows represent fluxes. Various N transformations in the soil and atmosphere lead to varying amounts of 15N enrichment or depletion in natural ecosystems, which can vary greatly with geographical location and environmental conditions. SOM represents soil organic matter derived from plant, microbial, and faunal organic tissues (adapted from Dawson et al. 2002).
evidence suggests that bulk soil δ15N and the δ15N of soil NO3- and NH4+ diverge from each other at various timescales. This suggests that foliar δ15N may be interpreted as the integrated signal of δ15N of all plant-available N forms over the growth history of a particular leaf. However, the leaf signal may not represent the current status of soil δ15N at the time of sampling. Symbiotic interactions with fungi and N2-fixing bacteria may also obscure the interpretation of the foliar δ15N value (Evans 2001). Further, observations have suggested that much of the variation in foliar δ15N among co-existing species may be due to the internal metabolism of fungi participating in the mycorrhizal
8
association (Handley et al. 1998). For example, under N-limited conditions, plants interacting with mycorrhizae will have a lower δ15N compared to the fungus due to fractionation during N transfer from the symbiont to the host plant (Evans 2001).
Differential uptake of multiple N sources with different δ15N values, differences in fractionation during uptake and assimilation, and physiological differences between plants and mycorrhizal symbionts are significant obstacles to the use of foliar δ15N values as a tool to assess the foliar uptake of reactive N. Currently, there is no adequate method for isolating and analyzing δ15N in soil N pools that are available for plant uptake. Consequently, the influence of foliar uptake on foliar δ15N is confounded by variations in the soil system. Variability in substrate supply, abiotic conditions, organism assemblages and their demand for N influence foliar δ15N values to a degree beyond that of pollution additions to the soil surface or foliage, making most field based studies challenging. Ultimately, enhancing our understanding of the fractionations and transformations occurring in and between soil N pools and whether there are systematic correlations among pools will dramatically improve our ability to use foliar δ15N as an indicator and allow us to examine foliar N uptake in natural systems.
Atmospheric nitrogen sources
Atmospheric N deposition can affect plant functioning via both foliar and soil uptake, with different effects on plant growth depending on the mode of entry and the type of N compound and plant species (Stulen et al. 1998).
9
Atmospheric reactive N can enter terrestrial ecosystems via wet, dry, and occult deposition. Most forms of atmospheric reactive N are generated from the partial oxidation of N2 at high temperatures and via the volatilization of animal waste. Total oxidized reactive N (NOy) and ammonia gases (NHx) are the most prevalent atmospheric reactive N sources, can be transported long distances, and are eventually deposited on terrestrial ecosystems as NO3- and NH4+ in precipitation, as gases, or as particles. Major sources of NOy typically include power production, transportation sources, off-road mobile sources, and other industrial sources (EPA 1998). Sources of NHx include both natural plant and soil emissions and anthropogenic emissions from agricultural and animal waste products (Srivastava 1992). In addition to inorganic N sources, there are several sources of naturally-occurring organic N, including plant pollen and sea-spray droplets (Driscoll et al. 2003). Atmospheric organic N may also be derived from anthropogenic inorganic N compounds reacting with hydrocarbons or organic particles in the atmosphere (Prospero et al. 1996) and generally comprises 30% of total atmospheric reactive N deposition (Neff et al. 2002). All of these compounds have distinct deposition velocities, travel different distances from point sources of N pollution, and have different probabilities of being deposited to the soil or directly to vegetation. For example, recent work by Sparks et al. (2003) and Turnipseed et al. (2005) has shown that peroxyacetylnitrate (PAN), an organic N pollutant and the dominant form of reactive N transported long distances in the atmosphere, is readily taken up through plant foliage, primarily deposits directly to vegetation via foliar uptake and is less likely to be deposited to the soil N pool, making it a potential source of N to the plant that predominantly enters through the foliar uptake pathway.
10
Wet deposition
Most forms of reactive gas-phase N will eventually be oxidized to a form [most commonly nitric acid (HNO3)] that is highly soluble and easily scavenged by particles or water droplets. These compounds are deposited to ecosystems within precipitation, low clouds and fog. This process is collectively termed wet deposition and refers generally to the transfer of soluble compounds from the atmosphere to the earth's surface. Wet reactive N deposition contains a variety of N compounds, most of which are available for biological utilization, including inorganic (NO3–, NO2, NH4+) and organic (amino acids, PAN, urea) chemical species (Peierls & Paerl 1997). Cloud deposition, which occurs through impaction of fog droplets on exposed surfaces, can contribute between 25% and 50% of total N deposition in high-elevation areas of the Northeastern United States (Anderson et al. 1999). These compounds can be deposited directly onto leaf surfaces and later incorporated into the plant tissue or deposited to the soil surface and added to the soil pool.
Dry deposition
In the absence of an aqueous phase, gases and dry particles can be transported to and deposited into terrestrial ecosystems in the form of dry deposition. Dry deposition of gaseous N pollutants has been shown to be of a similar magnitude (or potentially greater) to wet deposition in many regions (Holland et al. 2004). Monitoring efforts in North America and Western Europe report that reactive N deposition flux to terrestrial ecosystems via dry deposition is dominated by NO2, NH3, HNO3 and a relatively small amount of
11
particle-nitrate. However, most monitoring efforts only assess inorganic forms of N and Day et al. (2003) suggest that the dry deposition of organic N could be comparable to that of HNO3. Dry deposition estimates are still highly uncertain in most areas, and methodology is still an active area of research. It is desirable for ecologists to not only evaluate the magnitude of gaseous N deposition on terrestrial ecosystems, but to also monitor the isotopic composition of the deposited N and the resulting impact on foliar δ15N values.
Foliar assimilation of atmospheric nitrogen pollutants
The impact of atmospheric reactive N deposition to leaves depends on the physical and chemical characteristics of the pollutant and the plant species, including adsorption of the compound to the leaf surface, compound solubility within the internal plant cell wall and its reactivity with cellular components, and physiological factors such as stomatal conductance and the rate of N metabolism. Foliar uptake occurs through two routes: stomatal diffusion and uptake via the cuticle, although the latter is negligible for most chemical species. Cuticular uptake of N compounds (most commonly HNO3) refers to the active or passive transport of a molecule through the lipid bilayer of the plasmalemma into the cytosol. Gaseous N pollutants enter the leaf through one of these two routes and are assimilated by different biochemical pathways (Figure 1.2). For example, ammonia gas diffuses through the stomata and is dissolved in the apoplastic space to form NH4+. Ammonium is transported into the cytoplasm where it is either partitioned into storage or assimilated by the glutamine synthetase/glutamate synthase cycle (Stulen et al. 1998). In contrast, after uptake via diffusion through the stomata, NO2 enters the
12
substomatal cavity of the leaf and disproportionates to form NO3- and nitrite (NO2-). Eventually, NO2- is reduced to NH2 via nitrate reductase in the cytosol and nitrite reductase in the chloroplast, used in amino acid synthesis, and incorporated into primary N metabolism (Weber et al. 1998, Wellburn 1994).
Figure 1.2. Primary metabolic processes involved in the foliar uptake and assimilation of atmospheric ammonia (NH3) and nitrogen dioxide (NO2). NH3 and NO2 enter the apoplast via stomatal or cuticular uptake, are assimilated into primary N metabolism by various biochemical pathways. Dashed lines indicate the possible role of cytosolic GS1 or chloroplastic GS2 in NH4+ assimilation derived from atmospheric NH3 (adapted from Stulen et al. 1998).
13
Contribution of foliar uptake to total plant nitrogen assimilation
Vegetation has the capacity to assimilate atmospheric sources of N both through root and foliar pathways. Through modeling, direct foliar uptake of reactive N has been estimated to contribute 3 - 16% of the plant N demand for new growth tissue (Holland et al. 2004). Although this is a small proportion of the total N required for growth, N directly assimilated via foliar uptake is a direct addition to plant metabolism, would not necessarily influence soil acidity, and would likely influence plant productivity and carbon assimilation in Nlimited terrestrial ecosystems. This additional source of N to plants could also increase plant growth under increased atmospheric carbon dioxide (CO2). Carbon dioxide fertilization effects, when they are observed, are often not sustained as the plant community becomes limited by some secondary limitation. This limitation is often hypothesized to be N availability in many ecosystems (Beedlow et al. 2004). Plants become N deficient because of the relatively higher availability of carbon substrate for photosynthesis. Under such a scenario, N available through foliar uptake would enhance or at least sustain more rapid growth under elevated CO2. Experiments directly quantifying the proportion of N assimilated through foliage or demonstrating a positive relationship between foliar uptake of N and elevated CO2 have not yet been forthcoming.
Variation in foliar nitrogen assimilation via the foliar uptake pathway
The ability of vegetation to incorporate atmospheric reactive N via foliar uptake has been demonstrated, but few studies have examined the variation in leaf N
14
assimilation among plant species. For example, Morikawa et al. (1998) documented a 600-fold variation in leaf N assimilation among 217 plant taxa. However, most studies, including this one, have been limited by the use of unnaturally high fumigation levels and limited mechanistic explanations as to why some species have a higher capacity for foliar uptake than others.
Reactive N uptake by leaves is strongly controlled by diffusion processes (Thoene et al. 1991, Weber & Rennenberg 1996) and variation in stomatal dynamics among species likely explains some of the observed variation in foliar uptake capacity. However, variation in foliar uptake of reactive N among species is likely controlled by factors in addition to diffusional resistance. For example, if stomatal conductance remains fairly constant, variations in the biochemical capacity for NO2 assimilation would alter the sub-stomatal air cavity NO2 concentration, and subsequently influence the NO2 concentration gradient into the leaf and the rate of foliar NO2 uptake (Hereid & Monson 2001). This ʻinternalʼ resistance to uptake is often termed the mesophyll resistance and has been suggested to be controlled by a number of factors including: (1) the permeability of NO3-/NO2- ions through cell membranes and walls (Ammann et al. 1995), (2) the site of the activity in the primary nitrate assimilation pathway through which NOy is metabolized (Wellburn 1990), and (3) the activity of ascorbic acid and other antioxidant compounds in the leaf apoplast (Teklemariam & Sparks 2004).
Recent work by Eller and Sparks (2006) has simultaneously investigated the hypothesized controlling factors over reactive N flux into leaves. In this study, the investigators measured stomatal conductance as an estimate of the
15
diffusional resistance to gases entering the leaf, the concentration of apoplastic ascorbate to examine the reaction rates between gases and chemicals within cell walls of leaf tissue, and the activity of the enzyme nitrate reductase (NR) as a proxy for the transport rate of NO3- out of the apoplast and then incorporated the strengths of each of these controlling factors into a multiple regression model describing leaf uptake of NO2. As in previous studies, stomatal conductance was found to be the primary control over the foliar uptake of NO2. However, NR activity and leaf ascorbate concentrations explained significant variance in NO2 leaf flux, suggesting that enzyme activities and antioxidant concentrations in the cell wall likely underlie the differences we observe across species in mesophyllic resistance to NO2 uptake. For example, NR activity is variable across species and such differences could account for the broad variation in rates of NO2 uptake (Morikawa et al. 1998) because plants expressing higher NR activities may have higher NO2 uptake capacity at elevated NO2 levels where the cell wall may be saturated with NO3-/NO2- (Eller & Sparks 2006). Furthermore, antioxidants such as ascorbate could also contribute to uptake capacity depending on their concentration and associated ability to react with oxidants such as NO2. Ultimately, the concentration of ascorbate may control the steady-state flux of NO2 into a leaf. Increasing our knowledge of these leaf biochemical processes will aid in our understanding of the existing variation in foliar uptake capacity among species.
Foliar δ15N as a tool to determine the magnitude of foliar nitrogen uptake
The differences in δ15N between atmospheric N, plant-available soil N, and N
16
derived from biological fixation may be used as a natural tracer in plant systems. Further, the δ15N of atmospheric N compounds produced from anthropogenic activities may be significantly different from the natural, background N sources in the soil (Freyer 1991). This anthropogenic signal in foliar δ15N has been reported in several studies (Ammann et al. 1999, Qiao & Murray 1997, Siegwolf et al. 2001) and may be a reliable indicator of N pollution addition. For example, the δ15N of atmospheric NOy pollution from traffic exhausts and industrial combustion processes is mostly positive (-1% to +5%) and atmospheric NH3 originating from the volatilization of animal waste is usually very depleted (Heaton 1986, Macko & Ostrom 1994) compared to natural sources of inorganic N (e.g., NO3- from mineralization) that are usually negative [between -5‰ and -2‰ (Nadelhoffer & Fry 1994)]. Because of these differences, the relative magnitude of different N sources can be observed at the leaf level (Ammann et al. 1999, Siegwolf et al. 2001). Only if there are differences in δ15N among potential N sources can δ15N be used as an indicator for any of these sources (Robinson 2001).
Assuming the δ15N of the various sources are known, the amount of N assimilated by the plant can be quantified. The final isotopic composition of the leaf tissue is a combination of the various source values and masses. Take, for example, a plant that receives N from two sources: root uptake of soil NO3and foliar uptake of atmospheric NO2. If the isotopic ratios of both sources are known, we can use a two ended mixing model and the proportion of N obtained from foliar uptake (Xfoliar) is:
δ15 Nplant −δ15 Nsoil Xfoliar = 15 δ Natmosphere −δ15 Nsoil
€
17
[1]
where δ15Nplant, δ15Nsoil, and δ15Natmosphere are the N isotope compositions of the leaf, soil, and atmospheric N sources, respectively.
Current challenges for using foliar δ15N as a tool to determine the magnitude of foliar nitrogen uptake
The obstacles for using mixing models like the one described above and interpreting the foliar δ15N value lie in the presence of simultaneously available multiple N sources and currently unknown fractionation events that occur during N transformation, uptake, and assimilation. As discussed in section 2.2, this is a significant issue in the soil system allowing the generation of various foliar δ15N values driven by differences in soil N cycling. Although the mechanisms are largely unknown, differences between the δ15N of plant material vs. soil δ15N suggest some within-plant fractionation of N (Comstock 2001, Robinson et al. 1998). In addition, foliar uptake of reactive N presumably exhibits an array of largely unknown fractionation events as well. For example, N gases assimilated through the stomata both have variable values in the atmosphere and are likely influenced by the same fractionation events caused by diffusion that influence the incorporation of CO2 during photosynthesis. Therefore, a large fractionation is expected to occur during the diffusion of NO2 into the leaf. Fractionations associated with transport into plant cells and N reduction enzymes, although not yet quantified, are also likely to occur. Further fractionation likely occurs during diffusion and biochemical assimilation pathways by a fine-scale separation of pools. As an example, in the substomatal cavity, NO2 rapidly disproportionates in water within the cell wall and is converted to NO3-. Accurately defining the difference in isotopic
18
composition between the gaseous NO2 in the air and NO3- in the cell wall would describe the fractionation due to stomatal diffusion and disproportionation of NO2. However, for foliar uptake of reactive N, none of these fractionation events have been adequately resolved. Developing a more complete understanding of the source values and fractionation events associated with foliar δ15N in the plant-soil system is necessary to predict the importance of various N sources on the plant N budget (Handley et al. 1998). Promising techniques are currently being developed and used to elucidate some soil processes, such as dual-isotope techniques [e.g., δ15N and δ18O in NO3- (Durka et al. 1994)]. Ultimately, a solid understanding of these processes will be necessary for the production of a mechanistic model describing how δ15N changes during plant N acquisition and assimilation from both foliar and soil uptake pathways (Dawson et al. 2002).
Promising methods using foliar δ15N as an indicator of direct leaf nitrogen assimilation
A possible strategy for quantifying the foliar uptake of atmospheric reactive N pollution using foliar δ15N values is to enhance the magnitude of differences in δ15N in the various N pools against a background level such that the difference in signal may provide a quantifiable tracer. In the laboratory, the external chemical and isotopic composition of the N source(s) can be identified and controlled. We describe this type of laboratory experiment in the following section. In the field, a significant 15N-enrichment of the gas-phase N source would be necessary to create a large enough signal separation. The naturally
19
occurring range of δ15N found in nature is generally no more than ±20‰ (Handley & Scrimgeour 1997). Therefore, the 15N-enrichment of the gas source would need to be large enough to overcome this natural variation. Systems enriched in 15N ameliorate many of the problems of interpretation brought about by fractionation among N pools and are useful for following the movement of N through a plant and for determining rates of N assimilation within the plant (Dawson et al. 2002, Nadelhoffer & Fry 1994). In the following sections, we discuss several of the more promising laboratory and field efforts using 15N to quantify foliar N uptake.
Hydroponics
Hydroponics systems allow for the control of the isotopic composition of the nutrient solution N and, if adequately replenished, remain constant through time. Traditional hydroponics systems have been used to quantify foliar δ15N values (Evans et al. 1996, Pritchard & Guy 2005, Qiao & Murray 1997, Qiao & Murray 1998, Yoneyama & Kaneko 1989) and to explore the impact of NO2 fumigation on root uptake of N (Muller et al. 1996). However, these studies did not directly address the magnitude nor the consequences of foliar uptake of reactive N.
We have used combined hydroponics-fumigation systems to address several of the unresolved questions associated with the foliar uptake of reactive N (Figure 1.3).
20
Figure 1.3. Design of an individual hydroponics fumigation system used with seedlings for long-term (weeks) experiments. The root system is suspended in the nutrient solution while the shoot system is contained within the fumigation chamber. The seedling is sealed at the root-shoot junction with modeling clay to isolate the N sources. The nutrient solution is refreshed on a weekly basis and aerated by an aquarium pump. The NO2 concentration of the incoming air to the fumigation chamber is held constant by mixing concentrated 15NO2 gas with clean air by using mass flow controllers. It is vigorously mixed in the fumigation chamber by a fan.
Plants are transplanted into the system with the root systems suspended in nutrient solution and the shoot systems enclosed in a fumigation chamber. Using this system, we can simultaneously and independently control the forms, external concentrations, and isotopic compositions of N sources available to plant root and shoot systems. After specific exposure protocols (i.e., different combinations of N gas concentrations to the leaves and N fertilizer to roots), plant biomass and isotopic composition measurements are used to quantify the proportion of each N source utilized and the overall influence of the exposure protocol on plant growth. Using this system, I have
21
determined the partitioning of plant N among sources and the potential magnitude of NO2 assimilation in tomato (Lycopersicon esculentum). Depending on the root availability of NO3-, NO2 derived N in leaf tissue varied
%N Incorporated from Foliar 15NO2 Uptake
Tomato
between 1 and 7% at the end of the fumigation (Figure 1.4). 10
b
8
6
c 4
2
a a
0
0 ppb NO
2
40 ppb NO
2
0 ppb NO
Low NO3-
2
High
40 ppb NO
2
NO3-
Treatment Figure 1.4. Proportion (%) of total N content attributable to the direct foliar uptake of 15NO2-N in L. esculentum. Data are means ± 1 SE. Data with distinct lowercase letters indicate significant differences. From Vallano and Sparks (unpublished).
We have also begun to address the variation in foliar N assimilation among plant species at realistic concentrations and how this assimilation may influence foliar δ 15N values. Using the same hydroponics-fumigation system
22
to quantify species differences in foliar NO2 assimilation, we have examined differences in growth and foliar δ15N values in two species that contrast in their N reduction strategies: tomato (Lycopersicon esculentum) and tobacco (Nicotiana tabacum). In natural and cultivated systems, L. esculentum primarily reduces N in the roots, while N. tabacum primarily reduces N in the leaves, suggesting that N. tabacum may have an increased capacity to incorporate and assimilate atmospheric N sources via the leaves. Seed-grown plants were transplanted into the system and exposed to low or high (50μM and 500μM) root NO3- supply and filtered or 15NO2-enriched air (0 ppb, 20 ppb, and 40 ppb) for four weeks. A key finding from this work is that both species are capable of incorporating up to 15% of their total assimilated N using the foliar uptake pathway, although no significant variation in foliar uptake capacity between species was observed at the whole-plant level (Figure 1.5).
This result suggests that the foliar uptake of NO2 contributes a significant proportion of N to plant metabolism and is consistent with modeled values of foliar uptake in the literature (Ammann et al. 1999, Muller et al. 1996). We also observed a NO2-induced stimulation in whole-plant biomass production in both species similar to those observed in some fumigation studies (Muller et al. 1996, Siegwolf et al. 2001). These results suggest that NO2 is potentially beneficial to biomass production and can change whole-plant allocation patterns under certain conditions. Further analysis of NR activity is necessary to highlight any species differences in foliar uptake capacity. Obviously, hydroponics-fumigation experiments like the ones we describe here are not an adequate simulation of natural conditions. However, they do lend significant insight into the transport of N pollutants into plants, and the physiological
23
%N Incorporated from Foliar 15NO2 Uptake
Tomato
18 Tomato Tobacco
16 14
b
12 10 8 6
c
4 2 0
a a 0 ppb NO
2
Low
40 ppb NO
2
0 ppb NO
NO3-
2
High
40 ppb NO
2
NO3-
Treatment Figure 1.5. Proportion (%) of total N content attributable to the direct foliar uptake of 15NO2-N in N. tabacum (●) and L. esculentum (○). Data are means ± 1 SE. Data with distinct lowercase letters indicate significant differences for both species. From Vallano and Sparks (unpublished).
mechanisms that influence the capacity of various plant species to incorporate atmospheric reactive N pollution via foliar uptake. Our future work using these systems will include multiple species comparisons of foliar uptake capacity, combined applications of multiple atmospheric N pollution sources, and quantification of the fractionation events associated with the foliar N uptake and how this influences foliar δ15N values.
24
Field studies
Controlled laboratory experiments have yielded promising results for using foliar δ15N measurements as indicators of reactive N uptake through leaves. However, can we use these, or similar, methods to estimate the proportion of foliar uptake of anthropogenic reactive N in natural systems? Multiple field studies have used foliar δ15N values to estimate the contribution of total atmospheric N pollution in terrestrial ecosystems (Ammann et al. 1999, Gessler et al. 2002, Siegwolf et al. 2001, Wania et al. 2002). However, this work did not attempt to separate uptake pathways and most were likely complicated by many of the soil factors discussed earlier (e.g., mycorrhizal effects, N2 fixation, heterogeneity of soil N sources and transformations). Currently, no consistent mechanistic relationship has been reported between atmospheric N pollution, foliar uptake of atmospheric N pollution, and foliar δ15N values.
In temperate forest ecosystems, high concentrations of total N pollution addition to ecosystems have resulted in the 15N enrichment of soil and plants (Stewart et al. 2002). Measurements of foliar δ15N in pine needles representing both the soil and foliar uptake of N deposition in heavily polluted areas of Germany varied by an order of magnitude depending on the plants proximity to various emission sources (Jung et al. 1997). It has been suggested these trends are the result of the loss of 15N depleted N from soil and plants due to volatilization causing enrichment of the residual N fraction (Hogberg et al. 1999). This is likely true, but the role and importance of foliar uptake in determining the foliar δ15N value and what this means in terms of
25
plant function is still unknown and likely an important factor in these measurements.
Ammann et al. (1999) attempted to quantify the influences of soil and foliar uptake pathways on pollutant N assimilation by sampling whole plant communities over a growing season and examining the resulting patterns of foliar δ15N. They estimated NO2 uptake in Picea abies by analyzing atmospheric NO2, foliar δ15N, and soil δ15N values along a highway in Switzerland. This study showed that leaves of plants growing in close proximity to an emission source reflected the δ 15N values of the N pollution (Ammann et al. 1999). Specifically, the foliar δ15N values increased from 3.7‰ to -1.5‰ within one growing season. Total anthropogenic inputs of both soil-derived N (-3.7‰) and gaseous NO2 (5.7‰) contributed about 25% to the total N budget of vegetation adjacent to the emissions source. However, the experimental methodology did not allow for the separation of soil vs. foliar uptake sources of anthropogenic N. Additionally, whether the differences in δ15N are due to differential soil cycling, fractionations within the plant, or foliar uptake is unknown. It is critical that future research address the importance and resulting impacts of these fundamentally different modes of anthropogenic N uptake to terrestrial ecosystems.
Future research
Foliar δ15N measurements have the potential to be powerful indicators of the relative contribution of soil vs. foliar uptake of atmospheric reactive N inputs to terrestrial ecosystems in both rural and urban environments. Natural
26
abundance foliar δ15N measurements provide us with an integration of the δ15N of the N sources and fractionation events that occur during foliar N uptake, allocation, and emissions of N compounds from vegetation. However, the current understanding of sources and fractionation events is incomplete and using foliar δ15N as a tool to quantify a single process, like foliar uptake, is currently challenging. Although recent modeling efforts have made significant headway in identifying processes that influence foliar δ15N (Comstock 2001), no work has examined the fractionation events explicitly associated with foliar N uptake. For foliar δ15N measurements to become an effective research tool in determining foliar uptake of N pollution, we must further develop models for 15
N uptake and discrimination by leaves to a point analogous to those
developed for plant carbon isotope composition where all sources and fractionation events are considered (Evans 2001, Farquhar et al. 1982).
Additionally, the use of dual isotope methods promises to be a useful tool to investigate the impact of reactive N exposure; not only foliar uptake of atmospheric N pollution, but also as a way to analyze changes in physiology, morphology and carbon partitioning in vegetation exposed to atmospheric reactive N sources (Durka et al. 1994, Siegwolf et al. 2001). Recent advances in methodology using 13C/15N and 18O/15N dual isotope approaches (Cernusak et al. 2002, Gaudinski et al. 2005, McIlvin & Casciotti 2006) provide great potential to expand our knowledge of N source partitioning and fractionation events along multiple spatial and temporal scales. In particular, natural abundance ratios of 18O/16O in atmospheric NO3- sources differ from those of nitrification derived NO3- and may be a powerful diagnostic tool. Together with 15
N/14N, stable oxygen isotope analysis of leaf water could be very useful in
27
determining the fate of anthropogenic N sources to vegetation. This method has successfully been applied to identifying anthropogenic N sources to stream water (Piatek et al. 2005), but has not yet been applied to plant N dynamics, an area ripe for further investigation.
Laboratory based hydroponics-fumigation systems provide a foundation to test and verify our current understanding of the foliar uptake of pollutant N and how this additional source of reactive N to plants will affect the plant N budget. It may be possible to use a parallel strategy to that applied to hydroponicsfumigation systems in the field. Examining the divergence between natural abundance foliar δ15N and the δ15N of extractable NO3-/NH4+ (plant-available soil N) along a gradient of anthropogenic N deposition will assess whether the difference in magnitude between these measurements reflects the uptake of atmospheric N pollution directly via the leaves. Any such effort will need to account for the influence of mycorrhizae and verify the methodology of accurately quantifying soil solution δ15N.
Currently, there are no published experiments that have directly quantified the proportion of N assimilated through foliage or quantitatively demonstrated a positive relationship between foliar N uptake and anthropogenic N pollution in natural systems. Clearly, as atmospheric reactive N deposition increases across the globe, we need to develop effective methods to identify and trace the resulting ecological change due to human-mediated additions of reactive N. Future efforts must include both estimates of total N deposition and archives of foliar δ15N values across a variety of ecosystems.
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Conclusions
This chapter highlights the importance and utility of the foliar δ15N value as a tool for determining the magnitude and impact of atmospheric N pollution directly entering the foliage of vegetation. Substantial progress has been made in furthering our understanding of the mechanisms and environmental factors that control foliar δ15N and the contribution of foliar uptake of anthropogenic N pollution to individual plant N budgets. Recent hydroponics-fumigation studies have successfully used foliar δ15N as a tool for investigating and quantifying the proportion of foliar uptake of atmospheric N pollution. In situ coupled measurements of natural abundance foliar and soil solution δ15N across a range of ecosystems and species will shed further light on the validity of foliar δ15N as an accurate indicator of ecological change in plant N dynamics. A firm understanding of both the magnitude of foliar N uptake and whether foliar 15N values may be used as a tracer to quantify pollutant N assimilation is imperative for understanding the future global N cycle.
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Qiao Z. & Murray F. (1997) The effects of root nitrogen supplies on the absorption of atmospheric NO2 by soybean leaves. New Phytologist, 136, 239-243. Qiao Z. & Murray F. (1998) The effects of NO2 on the uptake and assimilation of nitrate by soybean plants. Environmental and Experimental Botany, 39, 33-40. Robinson D. (2001) δ15N as an integrator of the nitrogen cycle. Trends in Ecology & Evolution, 16, 153-162. Robinson D., Handley L.L. & Scrimgeour C.M. (1998) A theory for 15N/14N fractionation in nitrate-grown vascular plants. Planta, 205, 397-406. Robinson D., Handley L.L., Scrimgeour C.M., Gordon D.C., Forster B.P. & Ellis R.P. (2000) Using stable isotope natural abundances (δ15N and δ13C) to integrate the stress responses of wild barley (Hordeum spontaneum C. Koch.) genotypes. Journal of Experimental Botany, 51, 41-50. Rondon A. & Granat L. (1994) Studies on the dry deposition of NO2 to coniferous species at low NO2 concentrations. Tellus Series BChemical and Physical Meteorology, 46, 339-352. Rondon A., Johansson C. & Granat L. (1993) Dry deposition of nitrogendioxide and ozone to coniferous forests. Journal of Geophysical Research-Atmospheres, 98, 5159-5172. Siegwolf R.T.W., Matyssek R., Saurer M., Maurer S., Gunthardt-Goerg M.S., Schmutz P. & Bucher J.B. (2001) Stable isotope analysis reveals differential effects of soil nitrogen and nitrogen dioxide on the water use efficiency in hybrid poplar leaves. New Phytologist, 149, 233-246. Sparks J.P., Monson R.K., Sparks K.L. & Lerdau M. (2001) Leaf uptake of nitrogen dioxide (NO2) in a tropical wet forest: implications for tropospheric chemistry. Oecologia, 127, 214-221. Sparks J.P., Roberts J.M. & Monson R.K. (2003) The uptake of gaseous
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CHAPTER 2: QUANTIFYING FOLIAR UPTAKE OF GASEOUS NITROGEN DIOXIDE USING ENRICHED FOLIAR δ15N VALUES*
Abstract
The magnitude and impact of gaseous nitrogen dioxide (NO2) directly entering the leaves were investigated using foliar δ15N values in tomato (Lycopersicon esculentum) and tobacco (Nicotiana tabacum). Using a hydroponicsfumigation system, 15NO2 (20 ppb and 40 ppb) was supplied to shoot systems and NO3- (50 μM and 500μM) was supplied to root systems. Morphological, stable isotope and nitrate reductase activity (NRA) analyses were used to quantify foliar NO2 uptake and examine whether realistic concentrations of NO2 influenced plant metabolism. Nicotiana tabacum and L. esculentum incorporated 15 and 11%, respectively, of 15NO2-N into total biomass via foliar uptake under low NO3- supply. On a mass basis, N. tabacum and L. esculentum incorporated 3.3 ± 0.9 and 3.1 ± 0.8 mg of 15NO2-N into biomass, respectively, regardless of NO3- availability. There were no strong effects on biomass accumulation or allocation, leaf δ13C values, or leaf and root NRA in response to NO2 exposure. Foliar NO2 uptake may contribute a significant amount of N to plant metabolism under N-limited conditions, does not strongly influence growth at 40 ppb, and may be traced using foliar δ15N values.
*Vallano, D.M. and J.P. Sparks. 2008. Quantifying foliar uptake of gaseous nitrogen dioxide using enriched foliar δ15N values. New Phytologist 177(4): 946-955.
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Introduction
Nitrogen (N) deposition to ecosystems is potentially both an important source of nutrients to vegetation and a detriment to ecosystem function (Vitousek et al. 1997). The dramatic increase in atmospheric concentrations of reactive N oxides (NOy) and ammonia gases (NHx) from human activities has increased the input of N deposition to plants and soil (Gessler et al. 2002, Morikawa et al. 1998). These pollutant compounds are deposited to and incorporated into vegetation via two interrelated pathways: directly through the leaves and indirectly through the soil. In N-limited ecosystems, atmospheric reactive N compounds may represent a considerable source of N to vegetation and may have a significant impact on plant metabolism (Hill 1971, Siegwolf et al. 2001, Sparks et al. 2001).
Although numerous studies have shown the significant impacts of total N deposition on ecosystem processes (e.g. Aber et al. 2003, Galloway et al. 2003, Lovett 1994), few have investigated the amount of atmospheric reactive N incorporated directly by leaves. If foliar N uptake has a differential influence on plant metabolism compared with increased soil N availability, then understanding the source N partitioning between these two pathways will be necessary for predicting future ecosystem function.
The traditional view is that plants meet their N demand exclusively by root uptake of ammonium (NH4+), nitrate (NO3-), and dissolved organic N from the soil. However, plants are capable of incorporating atmospheric reactive N compounds, including NOy and NHx, directly through the leaves and
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assimilating this N into plant metabolism (Hereid & Monson 2001, Sparks et al. 2001, Vallano & Sparks 2007). The influence of N deposition on the root uptake pathway in isolation and the simultaneous uptake through leaves and roots has been well documented (Collier et al. 2003, Gessler et al. 1998, Muller et al. 1996). However, independently quantifying foliar N uptake and determining if this N is assimilated into biomass have remained challenging. In this study, I used N stable isotopes to independently quantify uptake and assimilation through leaves at realistic atmospheric N concentrations.
The ability of vegetation to incorporate N into metabolism from both leaves and roots suggests that the fate of N deposited to an ecosystem may differ fundamentally depending upon the entry pathway. For example, the chemical products of foliar N assimilation are nutrients and are directly incorporated from the leaf apoplast and used for growth (Gessler et al. 2000, Morikawa et al. 1998). By contrast, N soil additions during deposition may be incorporated into microbial biomass, revolatilized, or leached. Therefore, the direct entry of N into metabolism through leaves has the potential to fundamentally change plant productivity through a direct influence on photosynthesis. However, it is unclear if, and how, plant performance is differentially affected by changes in N deposition to the soil versus changes in N additions directly to leaves. Both forms of N entry into an ecosystem will likely increase under most scenarios of intensified industrialization. However, various gaseous reactive N forms have significantly different atmospheric lifetimes, suggesting that plant communities arrayed across the landscape experience not only unequal N deposition, but different chemical species depending upon the distance from a reactive N source.
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Some evidence suggests that the proportion of N incorporated through the foliar versus soil pathway may vary across the environment. For example, atmospheric reactive N is dominated by relatively reduced forms (e.g. NO2) near industrial sources. These compounds deposit more readily through the stomata than to other surfaces in the environment (Sparks et al. 2001). At distances further from N pollution sources, much of the reactive N is oxidized to organic forms such as peroxyacetyl nitrate. These compounds also readily enter plant stomata (Sparks et al. 2003) at a rate higher than that of deposition to other surfaces. In regions still further from reactive N sources, the atmosphere is often dominated by nitric acid (HNO3). HNO3 is soluble in water and tends to deposit to many surfaces in the environment at a faster rate than entry through the stomata (Byterowicz et al. 1998). As the composition of atmospheric reactive N compounds can vary across the landscape, with the result that they will be more or less likely to enter stomata, ecosystems may receive differential amounts of N through the foliar uptake pathway.
Nitrogen stable isotope ratios (15N:14N) are a potential tool for investigating N sources to plant metabolism (Robinson et al. 1998, Robinson et al. 2000), estimating the amount of atmospheric reactive N in a system, and identifying the uptake pathway into the plant. For instance, N sources available to plants often have varying N isotopic composition (δ15N) values (Kolb & Evans 2002). Foliar δ15N values may reflect the isotopic value of the N sources (Stewart et al. 2002). Therefore, the relative importance of various N sources for the plant N budget can be determined at the leaf level using N stable isotopes (Ammann et al. 1999). However, the variation in source δ15N values and an incomplete understanding of fractionation events make it challenging to
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interpret foliar δ15N values in natural systems because it is difficult to distinguish the δ15N values of multiple N sources (Evans 2001). One potential strategy for accurately quantifying foliar N uptake is to use a single N source enriched in δ15N as a tracer by enhancing the magnitude of differences in δ15N in the various N pools available to the plant in a controlled laboratory system. This strategy ameliorates many of the interpretation problems caused by fractionation among N pools by creating a large signal separation and is useful for following the movement of N through a plant and for following the movement of N through a plant and for determining rates of N assimilation (Dawson et al. 2002).
In this study, a hydroponics-fumigation system was used with enriched 15NO2, a common gaseous reactive N pollutant, as a tracer to create a separation in δ15N between N sources and to track foliar uptake and assimilation of NO2-N through plants. This system allowed me to simultaneously and independently control the forms, concentrations, and δ15N of the N sources. Using this strategy, I focused on three questions:
1. What proportion and amount of N can be attributed to direct foliar uptake of NO2? 2. How is this response influenced by NO3- availability through the root system? 3. Is there variation in foliar N assimilation among species, as indicated by differences in NO3- reduction location and activity? −
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The goals of this study were to quantify the magnitude of plant tissue N originating from foliar uptake and to begin to explore the physiological mechanisms underlying species-level differences in foliar uptake capacity. I hypothesized that, at realistic levels of atmospheric NO2 exposure, a significant amount of N entering metabolism could come from direct foliar uptake. Further, I expected that increased NO3- availability to roots would −
diminish the magnitude of foliar NO2 uptake and the impact of this pathway on N metabolism. Finally, I predicted variation in foliar N assimilation as a result of differences in NO3- reduction strategies among species.
Materials and Methods
Plants and treatments
Tomato (Lycopersicon esculentum Mill.) and tobacco (Nicotiana tabacum L.) seeds were purchased from a plant producer (Paramount Seeds, Inc., Palm City, FL, USA). Tomato and tobacco were selected for this study because both are fast-growing, herbaceous species that tend to express nitrate reductase differentially in the plant body, and grow well in hydroponics systems. Seeds of both species (n =36 per species) were germinated and seedlings were grown in pots in perlite (Sun-Gro Horticulture, Bellevue, WA, USA) in a climatecontrolled growth chamber (EGC, Chagrin Falls, OH, USA) at day and night temperatures of 27 and 21°C, respectively, under moderate light (500 µmol m-2 s-1) and a 16-h photoperiod. Plants were watered daily to saturation for 2 weeks and fertilized with a Hoagland solution including NO3- (δ15N = -2.7‰). Twenty-four 2-week-old plants were transplanted into the hydroponics-
44
fumigation system and used for measurements. The remaining 12 plants per species were harvested and analyzed for biomass, morphology, and isotopic composition to provide a baseline for the experimental plants.
The hydroponics-fumigation system consisted of four 50-L opaque polyethylene nutrient tanks (120 ×58 ×16 cm) each fitted with three separate airtight Plexiglas fumigation chambers (36 ×25 ×43 cm) with an opaque base. The system was located in a glasshouse with day and night temperatures of 27 and 21°C, respectively, relative humidity of 60-70%, and moderate light conditions (800 ± 75 µmol m-2 s-1) using natural and supplemental metal halide lighting (400 MH Econo Cool Grow Light; Sunlight Supply, Vancouver, WA, USA) under a 16-h photoperiod. Plants were transplanted into the system with roots suspended in nutrient solution (20°C) in a hydroponics tank and the shoots enclosed in a fumigation chamber (ambient temperature). Each plant was stabilized and fitted with modeling clay (Loctite; Henkel Consumer Adhesives, Avon, OH, USA) at the root-shoot junction at the base of each chamber, to ensure an impermeable seal between the fumigation and nutrient solution system components.
Two N regimes (50 and 500 µM) were applied to the roots using nutrient solutions with fixed concentrations of NO3- as the sole root N source in a modified quarter-strength Hoaglandʼs solution (Smart & Bloom 1993). The difference in concentration between the low-N (LN; 50 µM) and high-N (HN; 500 µM) regimes simulated N-limited and N-surplus conditions, respectively. The mean concentrations for the LN and HN regimes were 46 ± 2 and 481 ± 28 µM, respectively. The δ15N of the nutrient solution was -2.7 ± 0.2‰. The
45
nutrient solution was vigorously aerated at all times to provide adequate oxygenation and to ensure complete mixing. Solution pH was maintained at 5.8-6.2 by the addition of either KOH or H2SO4. A 130-L reservoir of stock solution was used for each experimental NO3- treatment. Nutrient solutions were replaced weekly to minimize microbial activity and to prevent N depletion. The nitrate concentration was measured weekly using an automatic autoanalyzer (Astoria Pacific, Inc., Clackamas, OR, USA).
Fumigation chambers were supplied with filtered (using activated charcoal) ambient air using a reciprocating air compressor (Model C403L; Gardner Denver, Quincy, IL, USA) at a flow rate of 15 L min-1. Enriched 15NO2 from a compressed tank (1% NO2 : 99% N2; Scott Marrin, Inc., Riverside, CA, USA) was diluted into the filtered air of selected chambers using high-precision rotometers and mass flow controllers (Models 03216-34 and 32044-00; ColeParmer, Vernon Hills, IL, USA) at fixed partial pressures (20 and 40 ppb). The δ15N of the enriched 15NO2 was 1720 ± 17‰, providing a large signal separation from the NO3- source. NO2 was selected as the atmospheric N source because it is a common atmospheric reactive N compound. Treatments were selected to simulate suburban (20 ppb) and urban (40 ppb) atmospheric NO2 conditions. Four randomly selected chambers were selected as controls and were not exposed to NO2 fumigation (0 ppb). Therefore, there were six treatments: (1) 0 ppb NO2 and 50 µM NO3-, (2) 20 ppb NO2 and 50 µM NO3-, (3) 40 ppb NO2 and 50 µM NO3-, (4) 0ppb NO2 and 500 µM NO3-, (5) 20 ppb NO2 and 500 µM NO3-, and (6) 40 ppb NO2 and 500 µM NO3-. Within each treatment, there were two chambers with four plants per chamber (n =8). NO2 and NO concentrations were monitored using a chemiluminescence NO-
46
NO2-NO analyzer (TECO Model 42; Thermo Environmental Instruments, Inc., Franklin, MA, USA). Non-incorporated NO2 was filtered using activated charcoal and exited the system through an output line extending outside the glasshouse. Temperature and relative humidity within the chambers were monitored using humidity and temperature probes (Model HMP45A; Vaisala, Inc., Boulder, CO, USA) connected to a datalogger (Model CR10x; Campbell Scientific, Inc., Logan, UT, USA).
Morphological and stable isotope analysis
Plants were harvested 28 d after the start of fumigation (or at the beginning of the experiment for baseline plants) and fresh tissue was weighed using a balance (Model M-2200; Denver Instrument, Denver, CO, USA). Whole-plant samples were separated into leaf, shoot, and root tissue. Shoot length and leaf number were measured for each individual and leaf area was estimated using a leaf area meter (LI-3100 Area Meter; Li-Cor, Inc., Lincoln, NE, USA).
After fresh weight morphological analyses, plant samples were dried and weighed, and subsamples were analyzed for tissue N and C contents and δ15N. Previously separated tissues samples were rinsed with deionized water to remove any NO2 deposited to the leaf surface and dried for 72 h at 55°C in a drying oven. The dried tissue samples were weighed and ground to a fine powder with a mortar and pestle, and sub-samples of 2.55-3.15 mg were weighed using a microbalance (Model 4504MP8; Sartorius Corp., Edgewood, NY, USA). Tissue N and C contents were measured using a CHN elemental analyzer (Model Carlo Erba NC2500; Thermo Finnigan, San Jose, CA, USA).
47
Tissue δ15N and δ13C were measured using a continuous flow isotope ratio mass spectrometer (Model Delta Plus; Thermo Finnigan, San Jose, CA, USA). All analyses were conducted at the Stable Isotope Laboratory at Cornell University.
Nitrate reductase assay
Nitrate reductase activity (NRA) was assayed in vitro on frozen leaf and root tissues from eight seedlings per treatment according to Scheible et al. (1997). NRA was determined as the rate of NO2- accumulation and was measured at 540 nm using a spectrophotomer (DU 640; Beckman, Fullerton, CA, USA). The samples were compared with a standard curve generated using a KNO2 stock solution.
Calculation of N source partitioning
Partitioning of plant N among sources (gaseous NO2 and nutrient solution NO3- ) was calculated using a two-ended linear mixing model (Dawson et al. 2002) and published fractionation factors for root NO3- assimilation (Kolb & Evans 2003). Because we used an artificially high enrichment of 15N in the NO2, fractionation events associated with leaf uptake were not detectable (i.e. the signal separation generated by the tracer was larger than all natural fractionations). Using this model, we can define the total amount of NO2 incorporated via direct foliar uptake during the fumigation period.
48
Statistical analysis
Data represent three separate experiments conducted within a single year, each time using a new set of experimental plants (20 September to 19 October 2005, 25 January to 20 February 2006, and 18 May to 13 June 2006). No significant differences were detected among experiments, and, for simplicity, the stable isotope and morphological data were pooled across experimental runs. Data were statistically evaluated using a multifactorial ANOVA, with NO3- in the nutrient solution and NO2 fumigation treatments as the two factors. The experimental design included six total treatments and twelve fumigation chambers, resulting in two replicates per treatment per experiment. Statistical analysis was carried out using data from three experimental runs, resulting in means calculated from a total of six fumigation chambers. Significance levels for all tests were P ≤ 0.05. Statistical analyses were performed using SAS software (Version 9.0) and SAS JMP software (Version 3.2.5; SAS Institute, Cary, NC, USA).
Results
Foliar uptake and N source partitioning
Both species incorporated N via foliar uptake into total biomass under NO2 fumigation (Fig. 2.1a,b). For both L. esculentum and N. tabacum, there was a significant proportion of N incorporated via foliar uptake in individuals exposed to NO2 regardless of NO3- supply (P < 0.05; Fig. 2.1a). Regardless of species, the highest proportion of N incorporated via foliar uptake was observed under
49
low NO3- (LN) conditions and the highest NO2 fumigation treatment (40 ppb NO2). Under this treatment, L. esculentum and N. tabacum incorporated 11.0
%N Incorporated from Foliar 15NO2 Uptake
mg N Incorporated from Foliar
Tomato
15
NO2 Uptake
Tomato
± 0.9 and 14.9 ± 0.8% of total plant N through foliar uptake. The δ15N values 18 16
(a)
Tomato Tobacco
14 12 10 8 6 4 2 0 0 ppb NO2 20 ppb NO2 40 ppb NO2 0 ppb NO2 20 ppb NO2 40 ppb NO2
4
(b)
Low NO3-
High NO3-
Treatment
3.5 3 2.5 2 1.5 1 0.5 0
0 ppb NO 20 ppb NO 40 ppb NO 2
2
Low NO3
2
0 ppb NO 20 ppb NO2 40 ppb NO2
-
2
High NO3-
Treatment
Figure 2.1. Proportion (%) (a), and amount (in mg) (b) of total N content attributable to the direct foliar uptake of 15NO2-N in N. tabacum (●) and L. esculentum (○) for each treatment. Data are means ± 1 standard error (n = 16 for L. esculentum; n = 12 for N. tabacum). Significance level was set at P ≤ 0.05. An asterisk indicates significant differences between species.
50
for L. esculentum and N. tabacum under this treatment were 188.5 ± 15.5‰ and 257.0 ± 14.6‰, respectively. Individuals of both species fumigated with NO2 and grown under LN conditions incorporated a significantly higher proportion of N via foliar uptake compared with individuals grown under high NO3- (HN) (P < 0.05). We observed significant differences in foliar NO2-N uptake between the two different NO2 concentrations (20 and 40 ppb) under LN conditions in N. tabacum (P = 0.0064) but not under HN conditions. We did not observe any such differences for L. esculentum regardless of NO3availability. Species differences were only observed in individuals exposed to 40 ppb NO2 under LN conditions. Under this treatment, N. tabacum incorporated 4% more N from foliar uptake than L. esculentum (P = 0.003).
When examined on a mass rather than a proportion basis (Fig. 2.1b), L. esculentum and N. tabacum incorporated 3.1 ± 0.8 and 3.3 ± 0.9 mg of NO2-N, respectively. In contrast to the proportional measurements, the total mass of NO2-N incorporated into plant biomass via foliar uptake was similar regardless of root NO3- conditions. Furthermore, there were no differences in the amount of N incorporated via foliar uptake between NO2 fumigation levels (20 vs 40 ppb) for either species. There were also no significant species differences in the amount of N incorporated from foliar uptake.
The amount of N derived from foliar uptake of NO2 in both species varied among root, shoot, and leaf tissues (Fig. 2.2a,b). In both species, the amount of total N incorporated via foliar uptake was significantly higher in leaf tissue compared with root and shoot tissues. In fumigated individuals of N. tabacum, leaf tissue contained 78–82% more N (in mg) from foliar NO2 uptake than root
51
Figure 2.2. Amount (in mg) of total nitrogen attributable to the direct foliar uptake of 15NO2-N in N. tabacum (a) and L. esculentum (b) for root (), shoot (☐), and leaf () tissue for each treatment. Data are means ± 1 standard error (n = 16 for L. esculentum; n = 12 for N. tabacum).
and shoot tissues (Fig. 2.2a). In fumigated individuals of L. esculentum, leaf tissue contained 69-93% more N (in mg) from the foliar NO2 uptake than root and shoot tissues (Fig. 2.2b). Similar to the total plant measurements, the total mass of NO2-N incorporated into specific tissue types was not influenced by root NO3- conditions for either species.
Morphology and biomass allocation
Neither L. esculentum nor N. tabacum linearly increased total biomass accumulation (in g DW) in response to NO2 exposure (Fig. 2.3a). However, in N. tabacum, NO2 exposure increased biomass accumulation at 20 ppb NO2 under both LN and HN supply (by 55 and 92%, respectively), suggesting a fertilizer effect of NO2 exposure. Interestingly, individuals fumigated with 40
52
ppb NO2 showed similar biomass accumulation to non-fumigated individuals. By contrast, NO2 exposure decreased biomass accumulation at 20 ppb NO2 in L. esculentum under LN and HN supply (by 59 and 32%, respectively), suggesting a toxic effect of NO2 exposure. Similar to N. tabacum, individuals fumigated with 40 ppb NO2 exhibited similar biomass accumulation compared with non-fumigated individuals (Fig. 2.3a). Species differences in total biomass across all treatments were only observed between individuals exposed to 20 ppb NO2 under LN conditions. Under this treatment, N. tabacum incorporated 70% more total biomass than L. esculentum (P = 0.012). Overall, no significant differences in growth were observed between experimental runs.
Both L. esculentum and N. tabacum exhibited significant differences in the ratio of root to shoot (stem + leaf) tissue in response to differences in root N supply. Further, N. tabacum also showed changes in biomass allocation in response to differences in NO2 exposure under LN conditions (P < 0.05, Fig. 2.3b). In both species, higher NO3- availability was accompanied by a change in whole-plant C partitioning, favoring shoot growth relative to root growth regardless of NO2 fumigation concentration and resulting in a lower root:shoot ratio compared with individuals under LN availability (P < 0.05). NO2 exposure stimulated shoot relative to root growth in N. tabacum, causing a decrease (by 40%) in the root:shoot ratio in individuals exposed to 40 ppb NO2 and LN conditions. However, NO2 exposure did not affect root:shoot ratios in esculentum. A decrease in the root:shoot ratio in N. tabacum at 20 ppb NO2 L. (by 14%) was also observed, but this difference was not significant. There were no significant species differences in biomass allocation observed for any treatments. Leaf area did not differ under any treatment for either species.
53
4.0
Total Biomass (g DW)
3.5
(a)
Tomato Tobacco
3.0 2.5 2.0 1.5 1.0 0.5 0 0 ppb NO2 20 ppb NO2 40 ppb NO2 0 ppb NO2 20 ppb NO2 40 ppb NO2
0.35
(b)
Treatment
0.30 Root:Shoot (g g-1)
High NO3-
Low NO3-
0.25 0.20 0.15 0.10 0.05 0 0 ppb NO
2
20 ppb NO
40 ppb NO
2
Low NO3
2
0 ppb NO
-
2
20 ppb NO
40 ppb NO
2
High NO3
2
-
Treatment
Figure 2.3. Total plant biomass (a) and root/shoot ratio (b) of N. tabacum () and L. esculentum (☐) over all treatments. Data are means ± 1 SE (n = 16 for L. esculentum; n = 12 for N. tabacum). Significance level was set at P ≤ 0.05. An asterisk indicates significant differences between species.
54
Differences in C and N allocation patterns were observed in response to differences in NO3- availability in L. esculentum and N. tabacum, but no effects of NO2 exposure were observed for either species (Fig. 2.4). Both species expressed lower C:N ratios under HN availability compared with LN availability (P < 0.05). Overall, N. tabacum exhibited higher C:N ratios under LN supply
18 Tomato Tobacco
16
C:N of Leaf Material
14 12 10 8 6 4 2 0 0 ppb NO
2
20 ppb NO
2
Low
40 ppb NO
2
0 ppb NO
NO3-
2
20 ppb NO
2
High
40 ppb NO
2
NO3-
Treatment Figure 2.4. Carbon to nitrogen ratio for each treatment in N. tabacum () and L. esculentum (☐). Data are means ± 1 SE (n = 16 for L. esculentum; n = 12 for N. tabacum). Significance level was set at P ≤ 0.05. An asterisk indicates significant differences between species. compared with L. esculentum and lower C:N ratios under HN supply compared with L. esculentum across all NO2 fumigation treatments. There was no withinspecies variation in C or N concentrations between NO2 fumigation levels regardless of NO3- supply for either species.
55
L. esculentum exhibited differences in the pattern of leaf δ13C in response to differences in NO3- availability (P < 0.05), but N. tabacum did not exhibit any differences (Fig. 2.5). Neither species showed differences in leaf δ13C
-21 Tomato Tobacco
13
δ C of Leaf Material
-22 -23 -24 -25 -26 -27 -28 -29 -30 -31 0 ppb NO
2
20 ppb NO 40 ppb NO 2
2
0 ppb NO
Low NO3-
2
20 ppb NO 40 ppb NO 2
2
High NO3-
Treatment Figure 2.5. Foliar δ13C across treatments in N. tabacum () and L. esculentum (☐). Data are means ± 1 SE (n = 16 for L. esculentum; n = 12 for N. tabacum). in response to differences in NO2 exposure. Species differences in leaf δ13C between N. tabacum and L. esculentum were observed for all treatments, with N. tabacum exhibiting higher leaf δ13C values than L. esculentum (P < 0.05).
Nitrate reductase activity
There were no differences in leaf or root NRA in response to differences in
56
NO3- availability or NO2 exposure in either species (Fig. 2.6a,b). Species differences in NRA were observed only in roots and only between individuals
1.0
-1
2
Tomato Tobacco
(a)
-
Leaf NRA (µmol min g FW NO )
1.2
Tomato
-1
0.8 0.6 0.4 0.2 0 (b)
2
Low NO3-
High NO3-
Treatment
0.6
Tomato
-1
-1
-
Root NRA (µmol min g FW NO )
0 ppb NO2 20 ppb NO2 40 ppb NO2 0 ppb NO2 20 ppb NO2 40 ppb NO2
0.4
0.2
0.0
0 ppb NO
2
20 ppb NO 40 ppb NO 2
Low
2
0 ppb NO
NO3-
2
20 ppb NO 40 ppb NO 2
High NO3
2
-
Treatment Figure 2.6. Leaf NRA (a) and root NRA (b) for each treatment in N. tabacum ( ) and L. esculentum ( ). Data are means ± 1 SE (n = 12 for L. esculentum; n = 12 for N. tabacum).
57
exposed to 20 ppb NO2 and HN supply (P = 0.018). Under this treatment, N. tabacum showed higher root NRA than L. esculentum (Fig. 2.6b). Despite only small amounts of variation in NRA, we tested for a positive relationship between leaf NRA and the amount of NO2-N incorporated via the foliar uptake pathway in both L. esculentum and N. tabacum (Fig. 2.7a,b). We observed a positive relationship between these parameters in L. esculentum (P = 0.019), although the correlation was not statistically significant in N. tabacum (P = 0.312). Tomato
-
Tobacco
20-LN 40-LN 20-HN 40-HN
1.2
-1
1.0
-1
-1
(a)
2
Leaf NRA (µmol min g FW NO )
(a) 1.2
-1
-
1.4
2
Leaf NRA (µmol min g FW NO )
1.4
0.8
0.6
0.4
0.2
0.0
0
4
8
12
16
1.0
0.8
0.6
0.4
0.2
0.0
20
0
4
8
12
16
20
15
mg N Incorporated from Foliar NO Uptake 2
Figure 2.7. Leaf NRA vs. mg N incorporated from foliar 15NO2 uptake in L. esculentum (a) and N. tabacum (b) for each treatment. Data are means ± 1 SE (n = 12 for L. esculentum; n = 12 for N. tabacum).
Discussion
The potential for foliar N uptake is established in the literature (Gessler et al. 1998, Muller et al. 1996, Takahashi et al. 2005). However, it remains challenging to independently quantify foliar uptake, rather than examining a
58
tracer applied simultaneously to leaves and soil, and to definitively determine if the N is assimilated into biomass. Further, most studies have evaluated foliar uptake at unrealistically high concentrations. The NO2 fumigation concentrations used in this study (20 and 40 ppb) are relevant to ambient NO2 exposure and trends. Currently, the United States Environmental Protection Agency Office of Air Quality Planning and Standards (OAQPS) has established a National Ambient Air Quality Standard for NO2 pollution at 53 ppb. Additionally, the national average has steadily declined since 1980 to an average of ∼20 ppb (EPA 2007).
We show that foliar uptake is a significant pathway of NO2 entry and that the magnitude of foliar NO2 uptake has the potential to influence plant metabolism. Further, there is some variation among species in their capacity to incorporate NO2 via foliar uptake, an area of research that has yet to be thoroughly addressed (although see Morikawa et al. 1998). The average proportion of NO2 incorporated via foliar uptake quantified in this study is similar in magnitude to values modeled for other plant species (Ammann et al. 1999, Muller et al. 1996). Specifically, Muller et al. (1996) estimated proportions of 10–15% in Norway spruce (Picea abies), Ammann et al. (1999) reported similar proportions of 10– 25% in P. abies, and Siegwolf et al. (2001) estimated values in the range of 15–20% of total plant N in hybrid poplar (Populus euramericana) originating from NO2 applied to both leaves and soil. However, these studies did not attempt to separate uptake pathways (soil versus foliar) and were likely complicated by many of the soil factors discussed in the Introduction. This study extends previous work by clearly
59
isolating and quantifying the direct impact of the foliar uptake pathway on plant metabolism.
We observed that foliar NO2 incorporation in both species and the magnitude of this uptake appeared to be related to soil N availability (Fig. 2.1a), perhaps suggesting that plants regulate the capacity for foliar uptake in response to changes in overall N availability. We also observed significant differences in foliar NO2-N uptake between the NO2 concentrations (20 and 40 ppb) under LN conditions in N. tabacum (P = 0.0064), suggesting that foliar uptake capacity in N. tabacum may be more sensitive to changes in NO2 concentrations than L. esculentum. However, when the incorporated foliar NO2-N was examined on a mass basis (Fig. 2.1b), we observed similar total amounts of NO2-N under both NO3- and NO2 treatments. Further, we observed no differences in the amount of incorporated foliar NO2-N in individuals fumigated with 20 or 40 ppb NO2, which may suggest that plants have a maximum threshold for the amount of N assimilated via foliar uptake. These results also indicate that foliar uptake rates are similar and any differences seen in the proportions of foliar uptake across NO3- treatments are likely driven by dilution into a larger plant N pool rather than an increase in foliar uptake rate in response to a decrease in soil N availability. Nitrogen from foliar uptake was partitioned differentially across tissue components (i.e. leaf, shoot, and root) in both L. esculentum and N. tabacum. Not surprisingly, we observed a greater amount of NO2 incorporated into leaf tissue compared with roots and shoots under all treatments (Fig. 2.2a,b). This pattern of N assimilation suggests that most of the NO2-N assimilated by the plant is directly incorporated into metabolism in the leaves and that there is
60
only limited reallocation of N to other plant organs. This pattern of N assimilation makes intuitive sense given the large sink for N in leaves and suggests a potential impact on plant productivity through a direct influence on photosynthesis.
No toxic effects on growth were observed, suggesting that the NO2 concentrations applied were not detrimental to growth (Fig. 2.3). In fact, the biomass information suggests that NO2 fumigation enhanced growth under some circumstances. For example, biomass production in N. tabacum increased by ∼92% under the 20 ppb NO2 and LN availability treatment. However, both species, when exposed to 40 ppb NO2, exhibited growth similar to that of control plants. This suggests that any beneficial fertilization effect occurs only at relatively low concentrations. The 40 ppb NO2 fumigation did not appear to enhance or inhibit growth. This may mean that increased growth as a result of additional N availability is counteracted by increased toxicity, but this requires further evaluation across a wider range of fumigation levels. An NO2-driven influence on biomass allocation was observed in N. tabacum (Fig. 2.3b); a decrease in root:shoot ratio in response to an increase in NO2 exposure under LN availability. This result is consistent with those of other studies (Muller et al. 1996, Siegwolf et al. 2001) that reported an increased C investment into leaves at high atmospheric N availability. This observation, as noted by others (Nussbaum et al. 1999, Siegwolf et al. 2001), suggests that a change in root:shoot ratio induced by NO2 may have indirect consequences for growth and survival. For example, root:shoot ratio often varies across plants based on water and nutrient availability to balance water uptake and loss.
61
Therefore, changes in root:shoot ratio induced by NO2 may negatively impact plant tolerance to drought or low nutrient conditions.
We examined the effects of NO2 exposure on foliar δ13C values (Fig. 2.5) to explore the influence of exposure on the integrated ratio of external to internal CO2 concentration (Ci:C). Typically, a higher N supply can lead to increased stomatal conductance or higher leaf carboxylation capacity influencing the foliar δ13C value (Schulze et al. 1996). Across all treatments, we observed differences in foliar δ13C values in only L. esculentum in response to higher NO3- availability, but not NO2 exposure. Our results suggest that NO2 exposure at these concentrations did not impact stomatal conductance or photosynthetic capacity.
Although the ability of vegetation to incorporate atmospheric reactive N via foliar uptake has been demonstrated, only a few studies have started to examine the variation in foliar N uptake capacity among species (e.g. Morikawa et al. 1998). In this study, we observed differences in patterns of biomass accumulation and the proportion of NO2 incorporated through leaves between L. esculentum and N. tabacum with increasing NO2 fumigation (Figs. 2.1, 2.3), reinforcing the idea that variation in foliar N uptake exists and may be influenced by species-specific properties. Diffusion processes and variation in stomatal dynamics among species are the primary drivers of variation in foliar N incorporation (Sparks et al. 2001, Weber & Rennenberg 1996). However, variation in foliar N uptake among species is likely controlled by factors in addition to diffusional resistance. For example, if stomatal conductance remains fairly constant, variations in the biochemical capacity for
62
NO2 assimilation would alter the substomatal air cavity NO2 concentration, and subsequently influence the NO2 concentration gradient into the leaf and the rate of foliar NO2 uptake (Hereid & Monson 2001). This ʻinternalʼ resistance to uptake is often termed the mesophyll resistance and has been suggested to be controlled by various factors, including the permeability of NO3- and NO2ions through cell membranes and walls (Ammann et al. 1995), the activity of ascorbic acid and other antioxidant compounds in the leaf apoplast (Teklemariam & Sparks 2004), and the site of activity in the primary assimilation NO3- pathway through which NO2 is (Wellburn 1990). To further investigate the role of the location of N reduction within the plant as a possible factor controlling the variation in foliar uptake capacity in L. esculentum and N. tabacum, we examined the relationship between leaf NRA and foliar NO2 incorporation. The balance between root and shoot NO3reduction and the impacts of foliar NO2 uptake on that balance, if variable among species, may strongly influence species responses to NO2 exposure. It has been previously reported that L. esculentum and N. tabacum differ in the location of N reduction (Gojon et al. 1994). In N. tabacum, NO3- is reduced into an organic form primarily in the shoot, while L. esculentum reduces NO3mainly in the roots. Previous work (as cited in Tischner 2000) has suggested that NRA stimulation will follow NO2 exposure, although other studies have reported very little, if any, NRA stimulation in response to fumigation (Muller et al. 1996). We originally hypothesized that these species would differ significantly in leaf NRA activity and provided a system with which to examine differences in NO2 uptake driven by differential metabolism. However, we observed no differences in leaf or root NRA between these two species regardless of NO2 exposure, including controls (Fig. 2.6). Similarly, our results
63
did not show a correlation between leaf NRA and the NO2 fumigation level in either species, indicating that leaf NRA was not stimulated by NO2 exposure (Fig. 2.6). However, when the entire range of leaf NRA was considered (Fig. 2.7), we observed a positive relationship between leaf NRA and the amount of NO2 incorporated via foliar uptake in L. esculentum, indicating the potential of leaf NRA stimulation in response to NO2 exposure.
Foliar NO2 uptake is significant in magnitude and can potentially influence plant metabolism and the plant N budget in terrestrial ecosystems impacted by atmospheric reactive N pollution. The results of this study give a quantitative estimate of the mass of NO2 assimilated into biomass through the foliar uptake pathway. Obviously, hydroponics-fumigation experiments like the one we describe here are not an adequate simulation of natural conditions. However, they do provide significant insight into the transport of N pollutants into plants, and the physiological mechanisms that influence the capacity of various species to incorporate atmospheric reactive N via the foliar uptake pathway. Given the significant role that this pathway may play in the uptake and assimilation of NO2, it is necessary that future research include multiple species comparisons of foliar uptake capacity, combined applications of multiple atmospheric N sources, and quantification of the fractionation events associated with foliar N uptake and how this influences foliar δ15N values. Results from such work will be imperative for the successful prediction of vegetative responses to increasing anthropogenic atmospheric N deposition.
64
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Gessler A., Rienks M. & Rennenberg H. (2000) NH3 and NO2 fluxes between beech trees and the atmosphere - correlation with climatic and physiological parameters. New Phytologist, 147, 539-560. Gessler A., Rienks M. & Rennenberg H. (2002) Stomatal uptake and cuticular adsorption contribute to dry deposition of NH3 and NO2 to needles of adult spruce (Picea abies) trees. New Phytologist, 156, 179-194. Gessler A., Schneider S., Von Sengbusch D., Weber P., Hanemann U., Huber C., Rothe A., Kreutzer K. & Rennenberg H. (1998) Field and laboratory experiments on net uptake of nitrate and ammonium by the roots of spruce (Picea abies) and beech (Fagus sylvatica) trees. New Phytologist, 138, 275-285. Gojon A., Passard C. & Bussi C. (1994) Root/shoot distribution of NO3assimilation in herbaceous and woody species. . In: A whole plant perspective on carbon-nitrogen interactions (ed E. Garnier), pp. 131147. SPB Academic Publishing, The Hague. Hereid D.P. & Monson R.K. (2001) Nitrogen oxide fluxes between corn (Zea mays L.) leaves and the atmosphere. Atmospheric Environment, 35, 975-983. Hill A.C. (1971) Vegetation: a sink for atmospheric air pollutants. Journal of the Air Pollution Control Association, 21, 341. Kolb K.J. & Evans R.D. (2002) Implications of leaf nitrogen recycling on the nitrogen isotope composition of deciduous plant tissues. New Phytologist, 156, 57-64. Kolb K.J. & Evans R.D. (2003) Influence of nitrogen source and concentration on nitrogen isotopic discrimination in two barley genotypes (Hordeum vulgare L.). Plant Cell and Environment, 26, 1431-1440. Lovett G.M. (1994) Atmospheric deposition of nutrients and pollutants in North-America - an ecological perspective. Ecological Applications, 4, 629-650.
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Morikawa H., Higaki A., Nohno M., Takahasi M., Kamada M., Nakata M., Toyohara G., Okamura Y., Matsui K., Kitani S., Fujita K., Irifune K. & Goshima N. (1998) More than a 600-fold variation in nitrogen dioxide assimilation among 217 plant taxa. Plant Cell and Environment, 21, 180-190. Muller B., Touraine B. & Rennenberg H. (1996) Interaction between atmospheric and pedospheric nitrogen nutrition in spruce (Picea abies L Karst) seedlings. Plant Cell and Environment, 19, 345-355. Nussbaum S., Ammann M. & Fuhrer J. (1999) Foliar absorption and use of airborne oxidized nitrogen by terrestrial plants. In: Nitrogen nutrition and plant growth (eds H. Srivastava & R. Singh), pp. 103-172. Science Publishers, Inc., Enfield, NH. Robinson D., Handley L.L. & Scrimgeour C.M. (1998) A theory for 15N/14N fractionation in nitrate-grown vascular plants. Planta, 205, 397-406. Robinson D., Handley L.L., Scrimgeour C.M., Gordon D.C., Forster B.P. & Ellis R.P. (2000) Using stable isotope natural abundances (δ15N and δ13C) to integrate the stress responses of wild barley (Hordeum spontaneum C. Koch.) genotypes. Journal of Experimental Botany, 51, 41-50. Scheible W.R., Lauerer M., Schulze E.D., Caboche M. & Stitt M. (1997) Accumulation of nitrate in the shoot acts as a signal to regulate shootroot allocation in tobacco. Plant Journal, 11, 671-691. Schulze E.D., Kelliher F.M., Korner C., Lloyd J., Hollinger D.Y. & Vygodskaya N.N. (1996) The role of vegetation in controlling carbon dioxide and water exchange between land surface and the atmosphere. . In: Global change and terrestrial ecosystems. (eds B. Walker & W. Steffen), pp. 77-92. Cambridge University Press, New York. Siegwolf R.T.W., Matyssek R., Saurer M., Maurer S., Gunthardt-Goerg M.S., Schmutz P. & Bucher J.B. (2001) Stable isotope analysis reveals differential effects of soil nitrogen and nitrogen dioxide on the water use efficiency in hybrid poplar leaves. New Phytologist, 149, 233-246.
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Smart D.R. & Bloom A.J. (1993) The relationship between kinetics of NH4+ and NO3- absorption and growth in cultivated tomato (Lycopersicon esculentum Mill cv. T5). Plant Cell and Environment, 16, 259-267. Sparks J.P., Monson R.K., Sparks K.L. & Lerdau M. (2001) Leaf uptake of nitrogen dioxide (NO2) in a tropical wet forest: implications for tropospheric chemistry. Oecologia, 127, 214-221. Sparks J.P., Roberts J.M. & Monson R.K. (2003) The uptake of gaseous organic nitrogen by leaves: A significant global nitrogen transfer process. Geophysical Research Letters, 30, 2189-2193. Stewart G.R., Aidar M.P.M., Joly C.A. & Schmidt S. (2002) Impact of point source pollution on nitrogen isotope signatures (δ15N) of vegetation in SE Brazil. Oecologia, 131, 468-472. Takahashi M., Nakagawa M., Sakamoto A., Ohsumi C., Matsubara T. & Morikawa H. (2005) Atmospheric nitrogen dioxide gas is a plant vitalization signal to increase plant size and the contents of cell constituents. New Phytologist, 168, 149-153. Teklemariam T.A. & Sparks J.P. (2004) Gaseous fluxes of peroxyacetyl nitrate (PAN) into plant leaves. Plant Cell and Environment, 27, 1149-1158. Tischner R. (2000) Nitrate uptake and reduction in higher and lower plants. Plant Cell And Environment, 23, 1005-1024. Vallano D. & Sparks J.P. (2007) Foliar δ15N values as indicators of foliar uptake of atmospheric nitrogen pollution. In: Stable Isotopes as Indicators of Ecological Change (eds T. Dawson & R. Siegwolf), pp. 93109. Elsevier Academic Press, San Diego. Vitousek P.M., Aber J.D., Howarth R.W., Likens G.E., Matson P.A., Schindler D.W., Schlesinger W.H. & Tilman D.G. (1997) Human alteration of the global nitrogen cycle: sources and consequences. Ecological Applications, 7, 737-750.
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Weber P. & Rennenberg H. (1996) Exchange of NO and NO2 between wheat canopy monoliths and the atmosphere. Plant and Soil, 180, 197-208. Wellburn A.R. (1990) Why are atmospheric oxides of nitrogen usually phytotoxic and not alternative fertilizers? New Phytologist, 115, 395429.
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CHAPTER 3: INFLUENCE OF DIFFERENT ROOT NITROGEN SOURCES ON THE CAPACITY FOR DIRECT FOLIAR NO2 UPTAKE AND ENZYME EXPRESSION IN NICOTIANA TABACUM
Abstract
Direct foliar uptake of gaseous reactive N has been observed in several plant species, but the mechanistic factors driving variation in foliar uptake capacity across species have yet to be rigorously tested. In this study, we investigated the influence of different root N sources (NO3- vs. NH4+) on enzyme expression and the capacity for foliar uptake of gaseous enriched 15NO2 in tobacco (Nicotiana tabacum) in a hydroponics-fumigation system. We measured differences in plant morphology, foliar δ15N, nitrate reductase (NR) activity, and glutamate synthetase (GS) activity to explore the potential drivers of foliar NO2 uptake capacity. The amount of NO2-N incorporated into biomass via foliar uptake was not influenced by differences in the root N source. We observed altered enzyme activities and foliar C isotope ratios associated with NO2 exposure and the form of N supplied to the roots. Specifically, leaf δ13C decreased, NR activity decreased, and GS activity increased in plants exposed to NO2, regardless of root N source. These results suggest that foliar uptake capacity is independent of the N form supplied to the roots and that enzyme expression within the primary N reduction pathways may be correlated with foliar uptake capacity in N. tabacum.
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Introduction
Reactive nitrogen (N) originating from human activity has dramatically increased in the past 50 years (Galloway et al. 2003, Vitousek 1994). A large proportion of this anthropogenic reactive N is emitted to the atmosphere and eventually deposited to terrestrial and marine ecosystems (Galloway et al. 2004). In particular, atmospheric levels of reactive N oxides (NOy) from combustion processes in vehicles and industry and ammonia gasses (NHx) from fertilizer and animal production have greatly increased the input of N to terrestrial ecosystems (Gessler et al. 2002, Morikawa et al. 1998). Atmospheric reactive N addition to terrestrial ecosystems is potentially both an important source of nutrients to vegetation and a detriment to ecosystem function if N accumulates in excess of biological demand (Vitousek et al. 1997). In N-limited ecosystems, atmospheric reactive N deposition may represent a considerable source of N addition to vegetation and have a direct impact on plant metabolism (Hill 1971, Siegwolf et al. 2001, Sparks et al. 2001).
Although numerous studies have shown the ecological consequences of total N deposition on ecosystem processes (Aber et al. 2003, Galloway et al. 2003, e.g., Lovett 1994), few have partitioned and quantified the respective roles of different uptake pathways of N into vegetation. Two pathways of N addition that may play important roles in terrestrial ecosystems are root entry via bulk N deposition to the soil surface and the direct uptake of N into plant metabolism through the leaves. Although some plants meet their N demand exclusively by root uptake of ammonium (NH4+), nitrate (NO3-), and dissolved organic N from
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the soil, many species are capable of incorporating atmospheric reactive N compounds, including NOy and NHx, directly through the leaves and assimilate this N into plant metabolism (Hereid & Monson 2001, Morikawa et al. 1998, Sparks et al. 2001, Sparks et al. 2003). Phytoxicity resulting from direct foliar N uptake has been documented, although plants were exposed to unnaturally high levels of fumigation in these experiments (Zeevart 1976). In contrast, the chemical products of N assimilation via direct foliar uptake can be potential nutrients and are directly incorporated into metabolism to support growth at more realistic atmospheric reactive N concentrations (Gessler et al. 2000, Vallano & Sparks 2007). Since direct foliar N uptake may have a differential impact on plant metabolism compared to increased soil N availability and the subsequent root uptake of reactive N, then understanding the source N partitioning between these two pathways will be necessary for predicting future ecosystem function, structure and composition (Sparks 2009 and references therein).
Foliar uptake of nitrogen dioxide (NO2; a common atmospheric N pollutant) has been described in several studies, although few have explicitly tested the potential mechanistic factors driving the documented variation in foliar NO2 uptake capacity across species (Vallano & Sparks 2008 and references therein). Diffusion into the leaf, controlled by stomatal dynamics, appears to exert the strongest control on the rate of foliar N uptake and is likely the primary driver of variation in foliar N uptake capacity observed among plant taxa (Hanson & Lindberg 1991, Sparks et al. 2001, Weber & Rennenberg 1996). However, a wide variation in foliar uptake capacity still exists when stomatal conductance is constant across taxa (Morikawa et al. 1998). Thus, it
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is likely that foliar NO2 uptake capacity is also influenced by a mesophyllic resistance (Gut et al. 2002, Teklemariam & Sparks 2006, Thoene et al. 1996), apoplastic antioxidant reactions by compounds such as ascorbate, and the subsequent removal of the primary end products of NO2 incorporation [nitrate (NO3-) and nitrite (NO2-)] from the apoplast (Eller & Sparks 2006).
In this study, we specifically examined the influence of the rate of elimination of NO3- and NO2- from the apoplast in Nicotiana tabacum, a species that tends to express N reduction differentially in the plant body, with more than 80% of N reduction occurring in the leaves (Wakhloo & Staudt 1988). We investigated this potential control on foliar uptake capacity by providing the root system with different N sources (NO3- and NH4+) and exposing the shoot systems to gaseous NO2. Previous studies have reported that bean plants supplied with NO3- versus NH4+ at the same concentration have higher rates of NO2 assimilation (Qiao & Murray 1997) and lower NO2 injury (Srivastava et al. 1992) in leaves. However, to better understand the complete range of mechanisms affecting the variation in foliar NO2 uptake capacity across plant species, it is necessary to examine the linkages between root N supply, associated assimilatory enzyme activity, and foliar NO2 uptake capacity.
Different root N sources may result in varying capacities for foliar NO2 uptake due to the nature of the associated assimilatory enzyme activities. Ammonium incorporated though the root uptake pathway is assimilated via glutamine synthetase (GS) primarily in the roots, although a small proportion of NH4+ may also be transported to and assimilated in the leaves (Bloom 1988). In contrast, NO3- incorporated via root uptake can be reduced in either the root or
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shoot tissue via nitrate reductase (NR) and eventually incorporated into amino acids and plant proteins. Gaseous reactive N compounds such as NO2 can be incorporated directly via foliar uptake and incorporated into N metabolism as either NO3- or NO2- ions, reduced to NO2- via NR activity if in the form of NO3-, converted from NO2- to NH4+, and eventually assimilated into organic compounds via the GS assimilatory enzyme pathway. Thus, the site and magnitude of activities of the NR and GS assimilatory enzyme pathways in response to both root and foliar uptake of various reactive N sources may also contribute to the large variation in foliar uptake capacity that has been documented across plant taxa (Eller & Sparks 2006, Gut et al. 2002, Hereid & Monson 2001, Morikawa et al. 1998, Teklemariam & Sparks 2004, Teklemariam & Sparks 2006, Thoene et al. 1996, Wellburn 1990).
The overarching goal of this study was to determine whether variation in root N sources and N reduction activity associated with different N assimilation pathways was directly correlated with foliar uptake capacity in Nicotiana tabacum. We measured the importance of different root N sources and their assimilation pathways on plant N metabolism using foliar N stable isotope ratios (15N/14N) (Robinson 2001, Robinson et al. 1998), that often reflect the isotopic value of the N sources (Stewart et al. 2002). We used an established hydroponics-fumigation system in which plants were exposed to enriched gaseous 15NO2 as a tracer to create a separation in δ15N between N sources and allow for simultaneous and independent control of the concentrations and δ15N of multiple N sources (Vallano & Sparks 2007, Vallano & Sparks 2008). Using this system, we had the following explicit objectives: (1) to determine the influence of two different root N sources (e.g. NH4+ versus NO3-) on the
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magnitude of foliar NO2 uptake, the expression of enzyme activity, and plant performance, and (2) to examine the potential relationships among root N source, enzyme expression and the magnitude of foliar NO2 uptake. We predicted that differential processing and assimilation of available root N sources would significantly impact the capacity for foliar NO2 uptake in N. tabacum.
Materials and Methods
Plants and treatments
Tobacco (N. tabacum) seeds were purchased from a plant producer (Paramount Seeds, Inc., Palm City, FL, USA). Tobacco was selected for this study because it is a fast-growing herbaceous species that grows well in hydroponic systems. Seeds (n = 60) were germinated and seedlings were grown in pots in perlite (Sun-Gro Horticulture, Bellevue, WA, USA) in a climate-controlled growth chamber (EGC, Chagrin Falls, OH, USA) at day and night temperatures of 27 and 21 °C, respectively, under moderate light (500 μmol m-2 s-1) and a 16-h photoperiod. Plants were watered daily to saturation for two weeks and fertilized with a Hoagland solution including NO3- or NH4+ of known δ15N. The δ15N values of the NO3- and NH4+ nutrient solutions were 0.56 ± 0.1‰ and 0.89 ± 0.1‰, respectively. Forty-eight two-week-old plants were transplanted into the hydroponics-fumigation system and used for measurements. The remaining 12 plants were harvested and analyzed for biomass, morphology, and isotopic composition to provide a baseline for the experimental plants.
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The hydroponics-fumigation system consisted of four 50 L opaque polyethylene nutrient tanks (120 × 58 × 16 cm) each fitted with airtight Plexiglas fumigation chambers (36 × 25 × 43 cm) with an opaque base. The system was located in a greenhouse with day and night temperatures of 27 and 21 °C, respectively, relative humidity of 60-70%, and moderate light conditions (800 ± 75 μmol m-2 s-1) using natural and supplemental metal halide lighting (400 MH Econo Cool Grow Light, Sunlight Supply, Vancouver, WA, USA) under a 16 h photoperiod. Plants were transplanted into the system with roots suspended in nutrient solution (20 ºC) in a hydroponics tank and the shoots enclosed in a fumigation chamber maintained at ambient temperature. Each plant was stabilized and fitted with modeling clay (Loctite, Henkel Consumer Adhesives, Avon, OH, USA) at the root-shoot junction at the base of each chamber to ensure an impermeable seal between the fumigation and nutrient solution system components.
Two N regimes simulating low N supply (50μM) were applied to the roots using nutrient solutions with fixed concentrations of either NO3- or NH4+ as the sole root N source in a modified quarter-strength Hoagland solution (Smart & Bloom 1993). The NO3- and NH4+ in the hydroponics nutrient solutions had the same δ15N values as the nutrient solution provided to plants prior to being placed in the system. The nutrient solutions were vigorously aerated at all times to provide adequate oxygenation and ensure complete mixing. Solution pH was maintained at 5.8-6.2 by the addition of either KOH or H2SO4. A 130 L reservoir of stock solution was used for each experimental N treatment. Nutrient solutions for each treatment were replaced weekly to minimize
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microbial activity and prevent N depletion. NO3- and NH4+ concentrations were measured weekly using an auto-analyzer (Astoria Pacific, Inc., Clackamas, OR, USA).
Fumigation chambers were supplied with filtered (activated charcoal), ambient air using a reciprocating air compressor (Model C403L, Gardner Denver, Quincy, IL, USA) at a flow rate of 15 L min-1. Enriched 15NO2 from a compressed tank (1% NO2 : 99% N2, Scott Marrin, Inc., Riverside, CA, USA) was diluted into the filtered air of selected chambers using high-precision rotometers and mass flow controllers (Models 03216-34 and 32044-00, ColeParmer, Vernon Hills, IL, USA) at a fixed partial pressure of 40 parts per billion (ppb). The δ15N of the enriched 15NO2 was 1720‰ ± 17‰, providing a large signal separation from the NO3- and NH4+ sources. NO2 was selected as the atmospheric N source because it is a common atmospheric reactive N compound. The fumigation treatment was selected to simulate urban (40 ppb) atmospheric NO2 concentrations. Six randomly selected chambers were used as controls and were not exposed to NO2 fumigation (0 ppb). Therefore, there were four treatments: (1) 0 ppb NO2 and 50 μM NO3-, (2) 40 ppb NO2 and 50 μM NO3-, (3) 0 ppb NO2 and 50 μM NH4+, and (4) 40 ppb NO2 and 50 μM NH4+. Within each treatment, there were three chambers with four plants/chamber (n = 12). NO2 and nitric oxide (NO) concentrations were monitored using a chemiluminescence NO-NO2-NOx analyzer (TECO Model 42, Thermo Environmental Instruments, Inc., Franklin, MA, USA). Nonincorporated NO2 was filtered using activated charcoal and exited the system through an output line extending outside the greenhouse. Temperature and relative humidity within the chambers were monitored using humidity and
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temperature probes (Model HMP45A, Vaisala, Inc., Boulder, Colorado, USA) connected to a datalogger (Model CR10x, Campbell Scientific, Inc., Logan, Utah, USA).
Morphological and Stable Isotope Analysis
Plants were harvested 28 days after the start of fumigation (or at the beginning of the experiment for baseline plants). Whole-plant samples were separated into leaf, shoot, and root tissue. Shoot length and leaf number were measured for each individual and leaf area was estimated using a leaf area meter (LI3100 Area Meter, LI-COR, Inc., Lincoln, NE, USA). Plant tissue samples were dried, weighed, and sub-samples analyzed for tissue N and C content and δ15N and δ13C. Previously separated tissues samples were rinsed with deionized water to remove any NO2 deposited to the leaf surface and dried for three days at 55 °C in a drying oven. The dried tissue samples were weighed using a balance (Model M-2200, Denver Instrument, Denver, CO, USA), ground to a fine powder with a mortar and pestle, and sub-samples of 2.553.15 mg were weighed for isotopic analysis using a microbalance (Model 4504MP8; Sartorius Corp. Edgewood, NY, USA). Tissue δ15N and δ13C were measured using a continuous flow isotope ratio mass spectrometer (Model Delta Plus; Thermo Finnigan, San Jose, CA, USA). Tissue N and C contents were measured using a CHN elemental analyzer (Model Carlo Erba NC2500; Thermo Finnigan, San Jose, CA, USA). All analyses were conducted at the Stable Isotope Laboratory at Cornell University.
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Enzyme Assays
Samples were weighed and immediately stored in liquid N until the preparation of the extracts. Root and leaf NR activity was assayed in vitro from 12 plants per treatment according to Scheible et al. (1997). NR activity was determined as the rate of NO2- accumulation and was measured at 540 nm using a spectrophotomer (Beckman DU 640). The samples were compared with a standard curve generated using a KNO2 stock solution. Root and leaf GS activity was assayed in vitro from 12 plants per treatment according to Hazelton (2007). GS activity was determined as the rate of gammaglutamylhydroxamic acid (GHA) accumulation and was measured at 540 nm using a spectrophotomer (Beckman DU 640). The samples were compared with a standard curve generated using a GHA stock solution.
Calculation of N Source Partitioning
Partitioning of plant N among different uptake pathways (foliar uptake of gaseous NO2 and root uptake of nutrient solution NO3- or NH4+) was calculated using a two ended linear mixing model (Dawson et al. 2002) and published fractionation factors for root NH4+/NO3- assimilation (Kolb & Evans 2003). Because we used an artificially high enrichment of 15N in the NO2, fractionation events associated with foliar uptake were likely not detectable (i.e., the signal separation generated by the tracer was much larger than all natural fractionations). Using this model, we defined the total amount of NO2 incorporated via direct foliar uptake during the fumigation period.
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Statistical Analysis
Data were statistically evaluated using a multifactorial analysis of variance (ANOVA), with N in the nutrient solution (NO3- or NH4+) and NO2 fumigation treatments as the two factors. We used the Tukeyʼs HSD procedure to make pair wise comparisons. Significant relationships between two variables were determined by simple linear regressions. Statistical analyses were performed with SAS JMP software (Version 7.0, 2007) and using a significance level of α = 0.05.
Results
Foliar Uptake and Nitrogen Source Partitioning
We found no significant differences in the total proportion and amount of N incorporated via foliar NO2 uptake into total biomass between plants grown under NO3- versus NH4+ root supply (Fig. 3.1). When exposed to gaseous NO2, plants grown on NO3- or NH4+ incorporated 11.6 ± 1.4 and 11.5 ± 1.4 mg NO2-N, respectively, into total biomass via foliar NO2 uptake (P < 0.0001). This amount represents 12.0 ± 0.6 and 12.4 ± 0.6% of total plant N in fumigated plants grown on NO3- or NH4+, respectively. Regardless of root N supply, the amount of total NO2-N incorporated via foliar NO2 uptake was significantly higher in leaf tissue compared to root and shoot tissues. In fumigated individuals, 88% and 87% of NO2-N from foliar NO2 uptake was assimilated into leaf tissue compared to root and shoot tissues in plants grown under root NO3- and NH4+, respectively. Similar to the total plant measurements, the total
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amount of NO2-N incorporated into specific tissue types was not influenced by root N supply. Foliar δ15N values for fumigated plants grown under the NO3and NH4+ treatments were 206.1 ± 10.2‰ and 215.9 ± 10.6‰, respectively.
Figure 3.1. Amount (in mg) of total nitrogen content attributable to the direct foliar uptake of 15NO2-N in N. tabacum and partitioning of N by tissue type for roots (white), shoots (black), and leaves (grey). Proportion (%) of total nitrogen content attributable to the direct foliar uptake of 15NO2-N is highlighted above bars. Data are means ± 1 standard error (n = 12). Different lower-case letters indicate statistically significant differences (P ≤ 0.05) across treatments.
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Morphology and biomass allocation
N. tabacum did not show changes in total biomass accumulation in response to different root N sources (Fig. 3.2a). Further, plants fumigated with gaseous NO2 showed similar total biomass accumulation compared to non-fumigated individuals, regardless of the root N source. We observed a non-significant trend for greater total biomass accumulation in plants exposed to NO2 and grown under NO3- supply (by 29%, P = 0.16), but this weak pattern was not observed in plants grown on NH4+ supply. We did not find any differences in the ratio of root to shoot (stem + leaf) tissue in non-fumigated plants grown on NO3- versus NH4+ (Fig. 3.2b). However, NO2 exposure did stimulate shoot growth relative to root growth in plants grown under NO3-, causing a decrease (by 22%) in the root to shoot ratio (P = 0.003). We did not observe this pattern for fumigated plants grown under NH4+.
Differences in C and N allocation patterns were observed in response to both root N source and NO2 exposure (Fig. 3.3a). The C:N ratio was lower in nonfumigated plants grown under NH4+ versus NO3- (P = 0.047). However, this trend reversed under NO2 fumigation with the C:N ratio being higher in fumigated plants grown under NH4+ versus NO3- (P = 0.011). Fumigated plants exhibited higher a C:N ratio than non-fumigated plants grown under root NH4+ (P < 0.0001). However, there were no differences in the C:N ratio between fumigated and non-fumigated plants grown under NO3-. Overall, leaf δ13C values were higher in plants grown under NH4+ versus NO3-, regardless of NO2 exposure (P < 0.01, Fig. 3.3b). Further, leaf δ13C values were more negative in
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Figure 3.2. Total plant biomass (a) and root to shoot ratio (b) in N. tabacum. Open bars represent plants exposed to 0 ppb gaseous 15NO2 and closed bars represent plants exposed to 40 ppb gaseous 15NO2. Data are means ± 1 SE (n = 12). Different lower-case letters indicate statistically significant differences (P ≤ 0.05) across treatments.
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Figure 3.3. Carbon to nitrogen ratio (a) and foliar δ13C (b) in N. tabacum. Open bars and circles represent plants exposed to 0 ppb gaseous 15NO2 and closed bars and circles represent plants exposed to 40 ppb gaseous 15NO2. Data are means ± 1 SE (n = 12). Different lower-case letters indicate statistically significant differences (P ≤ 0.05) across treatments.
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fumigated plants grown under NO3- (P < 0.0001) and NH4+ (P = 0.027) when compared to non-fumigated plants.
Enzyme Activity
We found no differences in root NR activity between non-fumigated plants grown on NH4+ versus NO3- (Fig. 3.4a). However, root NR activity was higher in fumigated plants grown on NO3- compared to NH4+ (P = 0.002). Within either root N source treatment, there were no differences in root NR activity between fumigated and non-fumigated plants. Leaf NR activity was higher in plants grown on NO3- compared to plants grown on NH4+, regardless of NO2 exposure (P < 0.05, Fig. 3.4b). However, leaf NR activity decreased in fumigated plants grown under NO3- (P = 0.024) or NH4+ (P = 0.0002).
Root GS activity was higher in plants grown on NH4+ versus NO3-, regardless of NO2 exposure (P < 0.0001, Fig 3.5a). We found no changes in root GS activity in fumigated plants grown on NO3-, while root GS activity increased in fumigated plants grown on NH4+ supply (P = 0.0264). Overall, leaf GS activity was higher in plants grown on NH4+ compared to plants grown on NO3-, regardless of NO2 exposure (P < 0.05, Fig. 3.5b). Leaf GS activity increased in fumigated plants grown on NO3- (P = 0.0078) and NH4+ (P < 0.0001).
When we examined the entire range of leaf NR activity expressed under NO2 fumigation, we found a significant correlation between leaf NR activity and the amount of NO2-N incorporated via foliar uptake in plants grown on NH4+ (P = 0.04, r2 = 0.52), but no significant relationship was observed between these
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Figure 3.4. Root (a) and leaf nitrate reductase (NR) activity (b) in N. tabacum. Open bars represent plants exposed to 0 ppb gaseous 15NO2 and closed bars represent plants exposed to 40 ppb gaseous 15NO2. Data are means ± 1 SE (n = 12). Different lower-case letters indicate statistically significant differences (P ≤ 0.05) across treatments.
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Figure 3.5. Root (a) and leaf glutamine synthetase (GS) activity (b) in N. tabacum. Open bars represent plants exposed to 0 ppb gaseous 15NO2 and closed bars represent plants exposed to 40 ppb gaseous 15NO2. Data are means ± 1 SE (n = 12). Different lower-case letters indicate statistically significant differences (P ≤ 0.05) across treatments.
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factors in plants grown under NO3- (Fig. 3.6a). Overall, leaf GS activity was not correlated with the amount of NO2-N incorporated via foliar uptake, regardless of the root N source (Fig. 3.6b).
Figure 3.6. Relationship between leaf NR activity vs. mg N incorporated via foliar 15NO2 uptake (a) and leaf GS activity and mg N incorporated via foliar 15 NO2 uptake (b) in N. tabacum (n = 12). Plants were supplied with either NO3(closed circles and dashed lines) or NH4+ (open circles and solid lines) as their sole root N source.
Discussion
The existence of the foliar N uptake pathway and some potential factors influencing foliar uptake capacity across taxa has been documented in the literature (Gessler et al. 1998, Hereid & Monson 2001, Hill 1971, Muller et al. 1996, Sparks et al. 2001, Takahashi et al. 2005). This study extends previous work by differentiating between deposited N among separate uptake pathways (soil vs. foliar) and investigating the influence of different root N sources and changes in downstream enzyme activity on foliar uptake capacity. We show
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that the proportion and amount of NO2-N incorporated into biomass via direct foliar uptake was not influenced by differences in the root N source (Fig. 3.1), suggesting that the overall rate of foliar NO2 uptake is not strongly impacted by the form of N supplied to the root system in N. tabacum. This was the case in spite of the observation of altered enzyme activities associated with availability of different root N sources and NO2 exposure (Figs. 3.4, 3.5, 3.6).
Our results suggest that the rate of foliar NO2 uptake is limited by resistances other than the activity of downstream enzymes in the N reduction pathway. Several studies have reported that the rate of foliar NO2 uptake is highly dependent on stomatal conductance (Eller & Sparks 2006, Gessler et al. 2000, Hanson & Lindberg 1991, Thoene et al. 1991, Weber & Rennenberg 1996). Further, N availability may influence stomatal conductance, photosynthetic rate, or both simultaneously, subsequently influencing foliar δ13C (Farquhar et al. 1982, Schulze et al. 1996).We observed lower foliar δ13C in response to NO2 exposure regardless of the root N source (Fig 3.3b), which is indicative of an increase in the intercellular CO2 concentration (ci) compared to the ambient CO2 concentration (ca). This pattern may be due to an increase in photosynthetic capacity and stomatal conductance in response to increased N supply via the foliar NO2 uptake pathway. However, we observed no differences in leaf N content in response to NO2 exposure regardless of root N source. Therefore, it is likely that the decrease in foliar δ13C primarily reflects changes in stomatal conductance due to direct foliar NO2 uptake in fumigated plants.
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Additionally, mesophyllic resistances likely play an important role in regulating rates of foliar NO2 uptake (Rondon et al. 1993, Sparks et al. 2001, Thoene et al. 1991, Weber & Rennenberg 1996), although these resistances have not been conclusively identified. Internal resistance to foliar NO2 uptake has been documented in some studies (Thoene et al. 1991), but not in others (Neubert et al. 1993). For instance, internal diffusion of gaseous NO2 into the leaf during the liquid phase versus the gaseous phase may be limited by the internal surface area of the chloroplasts exposed to intercellular air spaces per unit leaf area (Evans 1999, Evans et al. 1994). Other internal resistances include the disproportionation reaction of NO2 to NO2- and NO3- ions in the leaf apoplast by free-radical scavenging compounds such as ascorbate (Sparks et al. 2001) and differential permeability of NO3-/NO2- ions through cell membranes and walls (Ammann et al. 1995). Any one of these factors or, more likely, a combination of them likely drives much of the species-level variation observed in plant foliar uptake of NO2 at constant leaf conductance (e.g. Morikawa et al. 1998).
Root N supply and NO2 exposure did not appear to have a strong influence on plant growth, although we observed some changes nutrient allocation (Fig. 3.2). For instance, fumigated plants grown on NO3- supply had decreased ratios of belowground to aboveground tissue. This observation, as noted by others (Nussbaum et al. 1999, Siegwolf et al. 2001), suggests that NO2 fumigation directly or indirectly induces changes in basic plant allocation patterns. Such a change could have indirect consequences for growth over the long term such as plant tolerance to drought, low nutrient conditions, or other forms of environmental stress.
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We observed changes in both root and leaf enzyme expression in response to different root N source treatments (Figs. 3.4, 3.5). It is well known that NO3reduction requires the activity of NR, which is a substrate-inducible enzyme (Matt et al. 2001). Higher root and leaf NR activities in plants grown on NO3versus NH4+ root supply were likely a consequence of a greater NO3- substrate pool in each plant organ due to NO3- being the primary root N source. In contrast, it has been documented that root N form has little or no effect on GS activity (Claussen & Lenz 1999) or may show an increase in GS activity in the presences of NH4+ (Cruz et al. 2006, Lasa et al. 2002). We found higher root GS activity in plants grown on NH4+ versus NO3-. One possible mechanism driving this observation would be supplying roots with only NH4+ leads to a large NH4+ substrate pool in the roots and necessitates high GS activity to avoid toxic accumulation of NH4+. However, we also observed higher leaf GS activity in plants grown on NH4+ versus NO3-, a result reported by other studies on orchids (Majerowicz et al. 2000) and soybean and pea (Forde & Clarkson 1999).
Leaf NR and GS enzyme activities differed in their responses to NO2 exposure, regardless of the root N source. Previous work has reported that NO2 exposure stimulates NR activity (Eller & Sparks 2006, Murray & Wellburn 1985, Rowland et al. 1989), although inhibition of foliar N uptake has also been documented (Qiao & Murray 1998 and references therein). We observed a decrease in leaf NR activity in response to NO2 exposure, indicating that NO2 exposure may be limiting NR activity and hence, NO3- assimilation in N. tabacum (Fig. 3.4b). In contrast, we observed a positive relationship between leaf GS activity and the amount of NO2 incorporated via foliar uptake
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regardless of root N supply (Fig. 3.5b). These results suggest the potential of leaf GS activity stimulation in response to an increase in the overall flux of N at the leaf level from NO2 exposure.
When the entire range of foliar NO2 uptake was considered, we found a positive correlation between leaf NR activity and the amount of NO2 incorporated via foliar uptake in plants grown under NH4+, suggesting the potential for leaf NR activity to stimulate foliar NO2 uptake under these conditions. However, there was no significant correlation between these factors in plants grown under NO3- (Fig. 3.6a). We did not find a significant relationship between leaf GS activity and the amount of NO2 incorporated via foliar uptake regardless of the root N source (Fig. 3.6b).
The decrease in leaf NR activity and increase in GS activity under NO2 exposure may be due to several factors. Preferential disproportionation of NO2 into NO2- rather than NO3- in the leaf apoplast based on the activity of ascorbate and other antioxidant compounds is one potential scenario that may be driving the decrease in NR activity under NO2 exposure (Eller & Sparks 2006, Ramge et al. 1993). Under these circumstances, NO2 entering the leaf apoplast via foliar uptake would react preferentially with ascorbate rather than water to yield more NO2- versus NO3-, respectively (Gessler et al. 1998, Sparks 2009). This proportionation reaction is irreversible and dependent on the apoplastic concentration of NO2- and NO3- ions and the presence of apoplastic antioxidants such as ascorbate that may scavenge these ions (Stulen et al. 1998). We suggest that if NO2- were the primary chemical product of gaseous NO2 uptake and dissolution in the apoplast, then this
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compound would enter the NO3- → NO2- → NH4+ pathway further downstream than NO3-, resulting in decreased NR activity and increased GS activity compared to non-fumigated plants.
Although we did not observe any detrimental effects of NO2 exposure on plant growth, another plausible mechanism for the observed patterns in enzyme activity is toxicity from NO2 exposure. Previous work has acknowledged the potential phytotoxic effects of NO2 exposure on plant metabolism at unrealistically high ambient concentrations of 4-11 parts per million (ppm), although these effects have yet to be documented at realistic concentrations (Wellburn 1990, Zeevart 1976). Specifically, the dissolution of NO2-N in the apoplast originating from foliar uptake may yield a net H+ production and increases in NO3- concentration, that can then exceed the proton buffering capacity in the apoplast and result in toxicity (Rennenberg & Gessler 1999, Stulen et al. 1998), ultimately leading to the inhibition of NR activity observed in our study. This scenario is consistent with Qiao and others (1997, 1998), who have also noted declines in the rate of foliar NO2 uptake likely due to accumulation of H+ in the leaf apoplast from NO2 exposure in plants grown on either root NO3- or NH4+ supply.
Feedback inhibition between the NR and the GS reduction pathways may also contribute to the observed patterns in enzyme activity. Most likely, NO3- uptake and the NO3- and NH4+ assimilatory pathways are closely interlinked processes (Matt et al. 2001). For instance, NR activity may have been influenced not only by NO2 exposure, but also by the presence of NH4+ incorporated via the roots and translocated to the leaves. The increase in leaf
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GS activity in response to NO2 exposure observed in this study (Fig. 3.4b) suggests that foliar NO2 uptake may increase GS activity either by inducing its synthesis or activity or by preventing degradation or inactivation of the enzyme. Enhancement of this step of the N assimilatory pathway may be directly linked to the decrease in leaf NR activity under NO2 exposure. NR activity is often suppressed in the presence of NH4+ (Srivastava & Ormrod 1984). Specifically, NH4+ can inhibit NO3- uptake, inactivate the NR system or stop or slow the synthesis of active NR enzyme (Gessler et al. 1998).
The results of this study suggest that plant performance and foliar uptake capacity are independent of the root N form and that enzyme expression within the primary N assimilatory pathways may be correlated with foliar uptake capacity in N. tabacum. However, the observed changes in enzyme activity associated with N reduction indicate that the effects of foliar NO2 uptake at the physiological level may have important long-term impacts on plant metabolism. It is clear that foliar NO2 uptake contributes a significant amount of N to plants at the whole-plant level, although the effects of this uptake pathway on enzymatic processes have yet to be conclusively determined. Further work focused on the comparisons of foliar uptake capacity across multiple species, potential mesophyllic resistances to foliar uptake, and quantification of the within-plant N fractionation events associated with foliar N uptake will be crucial for predicting how vegetation will respond to increasing atmospheric N deposition.
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Acknowledgements
This work was supported by the National Science Foundation (NSF), Award # DEB- 0237674, to J.P. Sparks, the Cornell NSF Interdisciplinary Graduate Education, Research, and Training (IGERT) in Biogeochemistry and Environmental Biocomplexity, Award # DGE 0221658 to D. Vallano and an Andrew. W. Mellon Student Research Grant. The authors would also like to acknowledge E. Hazelton and T. Knight at the University of Southern Maine, who provided the protocol and invaluable assistance to conducting the GS assays.
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CHAPTER 4: PARTITIONING THE RELATIVE INFLUENCE OF SOIL N, MYCORRHIZAE, AND DIRECT FOLIAR N UPTAKE ON FOLIAR δ15N PATTERNS: CAN WE DETECT FOLIAR UPTAKE OF REACTIVE N?
Abstract
Vegetation is an important sink for atmospheric reactive N in N-limited systems and may be capable of incorporating reactive N compounds directly into leaves through the foliar uptake pathway. A proxy for atmospheric reactive N entering vegetation would be useful to estimate the impact of direct foliar N uptake on plant metabolism. Natural abundance foliar N isotopic composition (δ15N) is a practical tool for this purpose because plant-available N sources often have different isotopic compositions. Current understanding of foliar δ15N suggests these values primarily represent the integration of soil δ15N, direct foliar N uptake, mycorrhizal fractionation, and within-plant fractionations. Using a potted plant mesocosm system, we estimated the influence of mycorrhizae on foliar δ15N patterns in red maple (Acer rubrum) seedlings along an N deposition gradient in New York State. We found that mycorrhizal associations altered foliar δ15N in red maple seedlings from 0.03 – 1.01‰ across sites. Along the same temporal and spatial scales, we examined the influence of soil δ15N, foliar N uptake, and mycorrhizae on foliar δ15N in adult stands of American beech (Fagus grandifolia), black birch (Betula lenta), red maple (A. rubrum), and red oak (Quercus rubra). Using multiple regression models, atmospheric NO2 concentration explained 0%, 69%, 23%, and 45% of the residual variation in foliar δ15N remaining in American beech, red maple, red
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oak, and black birch, respectively, after accounting for soil δ15N. Our results suggest that foliar δ15N may be used to estimate pollution-derived atmospheric reactive N entering vegetation via the foliar N uptake pathway.
Introduction
Over the last century, human activity has greatly increased the amount of biologically reactive nitrogen (N) entering the natural environment and this trend is predicted to continue over the coming decades. A large proportion of reactive N is transported to the atmosphere and eventually deposited to terrestrial and marine ecosystems. On a global scale, atmospheric reactive N deposition has increased from 31.6 to 103 Tg N yr-1 from 1860 to the mid1990s and is projected to increase to 195 Tg N yr-1 by 2050 (Galloway et al. 2003, Galloway et al. 2004). In particular, the rise in atmospheric levels of N oxides (NOy) and ammonia gases (NHx) originating from fertilizer use and fossil fuel combustion has significantly increased the input of wet and dry N deposition to terrestrial ecosystems. This drastic increase in atmospheric reactive N levels and the resulting N that is subsequently deposited to plants and soil is often associated with dramatic changes in the structure, function and composition of ecosystems (Vitousek et al. 1997).
The short and long-term ecological consequences of increased atmospheric reactive N deposition can be potentially beneficial to vegetation by increasing the availability of nutrient sources and detrimental to ecosystem function if N accumulates in excess of biological demand (Driscoll et al. 2003, Vitousek et al. 1997). For instance, increased atmospheric reactive N deposition has been
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associated with detrimental ecological effects, such as soil acidification, plant nutrient imbalances leading to forest decline, and changes in species composition (Aber 1992, Schaberg et al. 2002). In N-limited ecosystems, such as the temperate forests in the northeastern US, atmospheric reactive N deposition may represent a considerable source of N addition to plants and may have beneficial implications for plant metabolism (Hill 1971, Siegwolf et al. 2001, Sparks et al. 2001). Although numerous studies have shown the ecological consequences of total N deposition on ecosystem processes (Aber et al. 2003, Galloway et al. 2003, e.g., Lovett 1994), few have partitioned and quantified the respective roles of the primary uptake pathways of atmospheric reactive N into vegetation.
The pathways of N addition that play important roles in temperate forest ecosystems are root entry via N deposition to the soil surface and direct uptake through the stomata in leaves. Although plants may meet their N demand exclusively by root uptake of nitrate (NO3-), ammonium (NH4+), and dissolved organic N from the soil, many species are capable of incorporating atmospheric reactive N compounds, including NOy and NHx, directly through the leaves and assimilating this N into plant metabolism (Hereid & Monson 2001, Sparks et al. 2001, Vallano & Sparks 2007, Vallano & Sparks 2008). If the foliar N uptake pathway has a differential influence on plant metabolism compared to increased soil N availability, then understanding the source N partitioning between these two pathways will be necessary for predicting how forests will respond to the increasing atmospheric N deposition (Sparks 2009 and references therein).
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Stable N isotope techniques are useful for estimating the amount of atmospheric reactive N in a system and identifying the uptake pathway of N sources into plant metabolism (Evans 2001, Handley & Raven 1992, Robinson 2001). Several studies have examined the link between N isotopic composition (δ15N) in plant and soil material to identify the correlations between foliar δ15N patterns and anthropogenic N inputs to temperate forest ecosystems (Amundson et al. 2003, Emmett et al. 1998, Garten & Vanmiegroet 1994, Hogberg 1997, Kahmen et al. 2008, Templer et al. 2007). Such measurements serve as practical tool for investigating stable isotope patterns across regional scales and providing an integrated measure of N cycling within a system (Pardo et al. 2007, Pardo et al. 2006). Further, foliar δ15N values that reflect the δ15N of distinct N sources have the potential to provide important information regarding the causal relationships between plant uptake, assimilation, and allocation of N from natural and human-derived N sources. However, few studies have successfully used this tool to examine the partitioning of N between the soil and foliar uptake pathways, and in particular, the magnitude of foliar N uptake of atmospheric reactive N deposition.
Despite the potential to use foliar δ15N as an indicator of plant N uptake patterns and N cycling, variation in the δ15N of N sources, fractionation due to mycorrhizal associations, and an incomplete understanding of mass dependent fractionation within plants has made it extremely challenging to accurately interpret foliar δ15N (Dawson et al. 2002, Evans 2001, Handley et al. 1998). Additionally, differences in the capacity for direct foliar uptake among plant species may also significantly impact the observed foliar δ15N patterns. To successfully detect and estimate the relative role of the foliar
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uptake pathway, greater understanding of the integrated effects of these processes on foliar δ15N patterns is necessary. Assuming that foliar δ15N values represent the integration of factors such as soil δ15N, direct foliar uptake of atmospheric reactive N deposition, and fractionation due to mycorrhizal associations during plant N assimilation, we sought to determine whether natural abundance soil and foliar δ15N measurments can serve as reliable indicators of atmospheric reactive N inputs, specifically via direct foliar N uptake, in temperate forest ecosystems.
In this study, we attempted to partition and estimate the relative importance of soil δ15N, mycorrhizal associations, and foliar N uptake on foliar δ15N patterns on several hardwood tree species along a gradient of increasing atmospheric NO2 concentration in New York State. Assuming that within-plant processes (e.g. translocation and assimilation) were similar within a species, we addressed these relationships by using a linked approach: (1) we estimated the influence of mycorrhizal fractionation on foliar δ15N patterns in seedlings using a potted plant mesocosm experiment where the influence of mycorrhizal fungi was eliminated; (2) we conducted intensive field measurements of foliar and soil δ15N along the same timescale in adult stands of trees and assessed the predictive ability of soil δ15N versus atmospheric NO2 concentration on foliar δ15N patterns using multiple regression models. Additionally, we estimated the influence of stomatal conductance and photosynthetic capacity by measuring foliar δ13C patterns among species. Using this dual approach strategy, we sought to determine whether these measurements could be used to estimate the predictive role of these factors on variation in foliar δ15N patterns and assess the potential impact of foliar uptake of reactive N
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deposition on temperate forest ecosystems. Alternatively, differences in soil N processing and within-plant N fractionations among species could preclude the interpretation of foliar δ15N patterns and the detection of foliar N uptake in these systems.
Materials and Methods
Site descriptions
The study was conducted at seven sites in eastern New York State across a north-south gradient of increasing N deposition (Fig. 4.1). Each site contained a mixed-hardwood forest comprised of clusters of adult stands of target tree species (Table 4.1). The sites included stands dominated by one of the
Figure 4.1. Site locations in New York State.
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Table 4.1. Site descriptions.
following tree species: American beech (Fagus grandifolia), red oak (Quercus rubra), red maple (Acer rubrum), and black birch (Betula lenta). A similar transect was established to measure the effects of urbanization on forest ecosystems (Pouyat et al. 1995) and to determine patterns of bulk N deposition, oak forest throughfall, and particulate dust deposition (Lovett et al. 2000). All sites are in close proximity to regional pollutant monitoring networks (e.g. the National Atmospheric Deposition Program/National Trends Network and the Clean Air Status and Trends Network) so that total N deposition (wet + dry inorganic N) was monitored and estimated at each site. Passive air samplers (Ogawa and Co. USA Inc., Pompano Beach, FL, USA) were installed and measured ambient atmospheric nitrogen dioxide (NO2) concentrations at each site from June 3, to September 17, 2007 (Table 4.1). Samplers were placed approximately 1 m above ground level, and when possible, installed adjacent to a potted plant mesocosm system. Samplers were housed in a shelter to minimize effects from precipitation and duplicate
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samples were collected at each site (n = 2). After exposure, samplers were immediately stored at 4°C and analyzed for ambient NO2 concentrations using an Astoria Pacific series 300 autoanalyzer (Astoria-Pacific Inc., Clackamas, OR, USA).
Potted Plant Mesocosm Study
Red maple (A. rubrum) seedlings, approximately 0.3-0.4 m tall were obtained bare root from Pinelands Nursery and Supply (Columbus, NJ, USA). Seedlings were selected for uniformity of height, transplanted into 12-L pots containing a mixture of perlite and vermiculite (Sun-Gro Horticulture, Bellevue, WA, USA), and grown in a climate-controlled growth chamber (EGC, Chagrin Falls, OH, USA) at day and night temperatures of 27 and 21 °C, respectively, under moderate light (600 μmol m-2 s-1) and a 16-h photoperiod. Seedlings were watered daily to saturation for 12 weeks and fertilized regularly with a modified Hoaglandʼs solution (Smart & Bloom 1993) containing NO3- with a known δ15N. Measurements of seedling size (height and stem diameter at 2.5 cm above the root collar), δ15N, δ13C and N and C content were made on a subset of seedlings prior to the start of the experiment to provide a baseline for experimental plants (n = 20).
Potted plant mesocosm systems were established at five of the sites (The New York Botanical Garden, Louis Calder Center, Mianus River Gorge, Cary Institute of Ecosystem Studies, and Huntington Wildlife Forest) in June 2007 and maintained through September 2007. Mesocosm systems were established in open fields exposed to full sun. Each system was approximately
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2.5 x 4.0 m and enclosed by fencing to eliminate browsing by deer and other small mammals. At each site, seedlings (n = 10) were randomly placed in holes to the depth of the pot and filled in with natural soil. All holes were dug 50% beyond the depth of the pot and filled with gravel and sand to allow for ample drainage. A 15 cm diameter hole was cut from the bottom of each pot to further allow for maximum drainage. Seedlings were drip irrigated to match natural precipitation patterns throughout the experiment using a 95-L barrel as a reservoir and an automatic drip irrigation system (Raindrip Inc., Fresno, CA, USA). To minimize any effects of atmospheric reactive N deposited to the soil surface, pots were covered with an opaque plastic shield (Ithaca Plastics, Ithaca, NY, USA) suspended around the base of each seedling and open on all sides to allow for adequate airflow and soil evaporation. This planting strategy matched the rooting zone temperature to the site during the growing season, allowed for irrigation, and minimized soil N from the environment entering each pot by lateral flow.
The experimental system was composed of two treatments: ʻcontrolsʼ with natural levels of root infection by mycorrhizae and ʻfungicide-treatedʼ plants where the fungicide Rovral (10% w/w iprodione, Bayer CropScience, Research Triangle Park, NC, USA) was applied to eliminate the influence of mycorrhizae on foliar δ15N. Rovral was applied every 3 weeks at rates of 2.0 g/m2 of soil surface in 500 ml of deionized water to half of the seedlings (n = 5) to reduce mycorrhizal colonization of roots (Gange et al. 1990, St Clair & Lynch 2005). The fungicide treatments were applied according to the methods of Ganade and Brown (1997). Rovral has been shown to be effective in reducing infection by mycorrhizal fungi in plant roots for a wide variety of plant species (Gange et
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al. 1990, Gange et al. 1993, St Clair & Lynch 2005). For many of the previously tested species, the fungicide did not appear to directly affect the plant or seed performance (Gange et al. 1992). Further, no negative side effects were detected on a range of soil invertebrates (Ganade & Brown 1997, Gange et al. 1993). To assess the bulk soil δ15N, soil cores were taken every 3 weeks between June 6 and September 12 2007 from each pot at a depth of 010 cm using a soil corer.
Seedlings were harvested in September 2007 (or prior to the experiment for baseline seedlings) and separated into leaf, shoot, and root tissues. Shoot length and leaf number were measured for each individual and leaf area was estimated for a subset of sample (n = 3) using a leaf area meter (LI-3100 Area Meter, LI-COR, Inc., Lincoln, NE, USA).
Field Sampling
Leaf and soil samples were collected every 3 weeks between June 6 and September 12 2007 from all seven sites (Fig. 4.1, Table 4.1) from June 2007 through September 2007. Leaf samples from 3-4 dominant species were collected using a wrist rocket (Falcon 2, Stunning Sales, Garland, TX, USA) from 5 randomly selected trees with a diameter breast height ≥15 cm at each site (Table 4.1). To assess the bulk soil δ15N, soil cores were taken at every 3 weeks from the base of each sampled tree at a depth of 0-10 cm using a soil corer.
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Stable Isotopic Analysis
Plant tissue samples from both the potted plant mesocosm study and field sampling were rinsed with deionized water to remove any reactive N deposited to the leaf surface and dried at 55 °C for 72-h. The dried plant tissue samples were weighed, ground to a fine powder with a mortar and pestle, and subsamples of 2.55-3.15 mg were weighed for isotopic analysis using a microbalance (Model 4504MP8; Sartorius Corp. Edgewood, NY, USA). Bulk soil samples were dried at 65°C for 48-h, sieved using a 2-mm mesh screen, ground to a fine powder using a mortar and pestle, and stored at 4°C until the time of analysis. Bulk soil and leaf N and C concentration were measured using a CHN elemental analyzer (Model Carlo Erba NC2500; Thermo Finnigan, San Jose, CA, USA). δ15N and δ13C were measured using a continuous flow isotope ratio mass spectrometer (Model Delta V; Thermo Finnigan, San Jose, CA, USA). All analyses were conducted at the Stable Isotope Laboratory at Cornell University, Ithaca, NY.
Statistical Analysis
We examined differences in Δ15N (the difference between foliar and soil δ15N), δ15N, δ13C and %N values among samples using two-way analyses of variance (ANOVA), with mycorrhizal treatment and NO2 concentration as the main effects. Tukeyʼs HSD procedure was used to make pairwise comparisons (α = 0.05) within and between field sites. We used single linear regression models to compare potential relationships among Δ15N, δ15N, δ13C, %N, and NO2 concentration for the potted plant and field data. Multiple linear regression
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models were constructed using soil δ15N and NO2 concentration to explain variation in foliar δ15N. The r2 selection method was used to determine the combination of predictors that resulted in the best multiple regression models explaining foliar δ15N. Significance levels for all tests were P ≤ 0.05. Statistical analyses were performed using SAS JMP software (Version 7.0, 2007).
Results
Potted plant mesocosm experiment
We observed a positive relationship between foliar δ15N and increasing NO2 concentration in red maple seedlings with mycorrhizal associations (r2 = 0.28 P = 0.01), although there was no trend in plants without mycorrhizal associations (Fig. 4.2a). There were significant differences in foliar δ15N between
Figure 4.2. Relationships between ambient atmospheric nitrogen dioxide concentration (a) and bulk soil δ15N (b) with foliar δ15N in red maple seedlings without mycorrhizal associations (open circles) and with mycorrhizal associations (closed circles). Data are means ± 1 standard error (n = 5).
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mycorrhizal and non-mycorrhizal plants (P < 0.01) when considered across sites along the NO2 concentration gradient. When the influence of mycorrhizae was removed, foliar δ15N values were altered 0.03 – 1.01‰ depending on the site location. We found no significant correlation between foliar and soil δ15N in red maple, regardless of mycorrhizal associations (Fig. 4.2b). We estimated the influence of mycorrhizal fractionation on foliar δ15N values by calculating the difference in foliar δ15N between the mycorrhizal and non-mycorrhizal individuals for each site along the atmospheric NO2 concentration gradient. Defined as the mycorrhizal fractionation, the correction factor had a range of –0.055 to 0.741 ‰ across sites (data not shown).
There was no trend between foliar δ13C and increasing NO2 concentration in red maple, regardless of the mycorrhizal treatment (Fig. 4.3). Foliar N content was negatively correlated with increasing NO2 concentration in red maple with mycorrhizal associations (r2 = 0.30, P < 0.01) and without mycorrhizal associations (r2 = 0.45, P = 0.0004). However, this trend was driven completely by a single site location and is likely not strongly correlated with increasing NO2 concentration along the gradient. There were no differences in foliar N content between mycorrhizal and non-mycorrhizal treatments when considered across sites along the atmospheric NO2 concentration gradient.
Field Sampling
There were significant trends between Δ15N (the absolute difference between foliar and soil δ15N) and both soil δ15N and increasing NO2 concentration on foliar δ15N patterns in tree species. The Δ15N value for all species showed a
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Figure 4.3. Relationship between ambient atmospheric nitrogen dioxide concentration and (a) foliar δ13C in red maple seedlings without mycorrhizal associations (open circles) and with mycorrhizal associations (closed circles) and (b) foliar nitrogen content in seedlings without mycorrhizal associations (open squares) and with mycorrhizal associations (closed squares). Data are means ± 1 standard error (n = 5).
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negative relationship with increasing NO2 concentration (Fig. 4.4). Black birch and American beech showed a negative correlation between Δ15N and increasing NO2 concentration (P = 0.03 and P = 0.02, respectively). Although not significant at α = 0.05, we observed a negative trend between Δ15N and increasing NO2 concentration in red oak (P = 0.07). There was not a significant correlation between Δ15N and increasing NO2 concentration in red maple.
Figure 4.4. Relationship between Δδ15N (the absolute difference between soil δ15N and foliar δ15N) and ambient atmospheric nitrogen dioxide concentration in adult trees. Species include American beech (open squares), red maple (closed squares), red oak (open circles) and black birch (closed circles). Data are means ± 1 standard error (n = 20).
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Single linear regressions were used to determine the influence of soil δ15N and increasing NO2 concentration on foliar δ15N patterns in tree species. In general, we found significant correlations between foliar δ15N and both soil δ15N and increasing NO2 concentration when tested singly (Fig. 4.5). There were significant and positive relationships between foliar and soil δ15N values in the following species: American beech (Fig. 4.5a), red maple (Fig. 4.5c), red oak (Fig. 4.5e), and black birch (Fig. 4.5g). We also found that foliar δ15N was negatively correlated with increasing NO2 concentration in the following species: American beech (Fig. 4.5b), red maple (Fig. 4.5d), and red oak (Fig. 4.5f). However, there was not a significant relationship between foliar δ15N and increasing NO2 concentration in black birch.
Patterns in foliar δ15N were best explained by a multiple regression model containing both soil δ15N and the average NO2 concentration (in ppb) at each site as predictors (Fig. 4.6). Multiple linear regression models containing both factors explained 40%, 56%, 47%, and 43% in American beech, red maple, red oak, and black birch, respectively. Partial residual plots for each predictor were generated to determine how much of the variation in foliar δ15N was explained by increasing NO2 concentration after accounting for the effects of soil δ15N, the factor with the strongest relationship to foliar δ15N. After accounting for soil δ15N, increasing NO2 concentration explained 0%, 69%, 23%, and 45% of the residual variation in foliar δ15N remaining in American beech, red maple, red oak, and black birch, respectively.
Applying the mycorrhizal correction factor calculated using the potted plant mesocosm foliar δ15N data did not alter the field relationship between foliar
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Figure 4.5. Relationship between foliar δ15N and bulk soil δ15N (a, c, e, g) and ambient atmospheric nitrogen dioxide concentration (b, d, f, h) in adult trees. Species include American beech (open squares), red maple (closed squares), red oak (open circles) and black birch (closed circles). Data are means ± 1 standard error (n = 20).
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Figure 4.6. Partial residual plots for the predictor variables included in the multiple regression model explaining foliar δ15N. The correlation of bulk soil δ15N and the residuals of the model describing foliar δ15N with NO2 concentration (a, c, e, g) in tree species. The correlation of NO2 concentration and the residuals of the model describing foliar δ15N with bulk soil δ15N (b, d, f, h) in tree species. Species include American beech (open squares), red maple (closed squares), red oak (open circles) and black birch (closed circles). Data are means ± 1 standard error (n = 20).
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δ15N, soil δ15N and NO2 concentration in red maple (Fig. 4.7). When this simplistic mycorrhizal correction factor was applied, there was no significant difference in the relationship between foliar δ15N and NO2 concentration in corrected (r2 = 0.54, P < 0.0001) and uncorrected data (r2 = 0.52, P < 0.0001, Fig. 4.7a). Similarly, there was not a difference in the relationship between foliar and soil δ15N between corrected (r2 = 0.97, P < 0.0001) and un-corrected (r2 = 0.87, P < 0.0001) data (Fig. 4.7b).
Figure 4.7. Relationship between foliar δ15N and (a) ambient atmospheric nitrogen dioxide concentration and (b) bulk soil δ15N in red maple trees. Data represent original foliar δ15N (open squares) and foliar δ15N corrected for mycorrhizal fractionation (closed squares). Data are means ± 1 standard error (n = 20).
In general, we observed no relationship between foliar δ13C and increasing NO2 concentration in most species, with the exception of a negative trend between foliar δ13C and increasing NO2 concentration in red maple (r2 = 0.19, P < 0.01, Fig. 4.8a). Likewise, we found no relationship between foliar N content and increasing NO2 concentration in most species (Fig. 4.8b).
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However, there was a negative correlation between foliar N content and increasing NO2 concentration in red maple (r2 = 0.21, P = 0.006).
Figure 4.8. Relationship between ambient atmospheric nitrogen dioxide concentration and (a) foliar δ13C and (b) foliar nitrogen content in adult trees. Species include American beech (open squares), red maple (closed squares), red oak (open circles) and black birch (closed circles). Data are means ± 1 standard error (n = 20).
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Discussion
We found significant trends in natural abundance foliar δ15N values across sites along a transect of increasing atmospheric NO2 concentration in both red maple seedlings and in adult trees species of American beech, red maple, red oak, and black birch (Figs. 4.2, 4.5). This range of −4-2‰ was similar to previous values recorded in other temperate forest studies (Nadelhoffer & Fry 1994, Pardo et al. 2007, Pardo et al. 2006, Templer et al. 2007). Overall, we found strong patterns of decreasing foliar δ15N values across the following species: Red maple > red oak > American beech > black birch. These patterns in foliar δ15N observed in adult tree species may be potentially linked to lower nitrification rates in the soil N pool (Pardo et al. 2007). Additionally, it is likely these patterns are driven by systematic variation among species, possibly due to a suite of differences in rooting depth, availability of NH4+ versus NO3- in the soil N pool, phenology, local nitrification rates, and mycorrhizal associations (Nadelhoffer et al. 1999).
Variation in mycorrhizal fractionation among species is thought to be a controlling driver of foliar δ15N patterns, although our results did not show support for such a relationship in red maple seedlings. For instance, it has been shown that ecto- or ericoid mycrohizal species such as red oak typically show a greater degree of isotopic fractionation associated with root N uptake from the soil N pool (Hobbie & Hobbie 2008). However, red maple is a vesicular arbuscular mycorrhizal species and such species are assumed to discriminate only marginally against 15N during uptake (Handley & Raven 1992), which could explain the observed patterns found in this study. Further,
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applying the mycorrhizal correction factor calculated using the potted plant mesocosm foliar δ15N data did not alter the field relationship between foliar δ15N, soil δ15N and NO2 concentration in red maple, suggesting that mycorrhizal fractionation did not strongly influence foliar δ15N patterns in adult stands of red maple (Fig. 4.7).
In general, we did not find significant correlations between either foliar δ13C or foliar N content and increasing NO2 concentration across sites along the transect (Figs. 4.3, 4.8), suggesting that variation in integrated seasonal stomatal conductance was similar across sites along the transect. Additionally, these results indicate that it is unlikely differential rates of atmospheric NO2 flux into leaves were likely playing a role in driving the observed foliar δ15N patterns.
We used patterns in Δδ15N, or the absolute deviation of foliar δ15N from soil δ15N, to infer the sensitivity of plant species at each of the sites to increasing rates of human-derived reactive N deposition (Fig. 4.4). Consistent with previous studies, foliar δ15N values were depleted relative to surface soil δ15N, usually resulting from mineral soil N uptake which is depleted in 15N compared to sampled soil surface N or soil organic N (Kahmen et al. 2008, Robinson 2001). Regardless of species, Δδ15N decreased as the NO2 concentration increased across the transect of increasing NO2 concentration. These patterns may be interpreted as a shift in the dominant N source of species from mineral soil N sources to atmospheric reactive N sources from human-derived pollution. Variation in the capacity for foliar uptake of atmospheric reactive N among species may be driving the observed species differences in Δδ15N.
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Additionally, species differences in N uptake preferences may also contribute to the observed variation in Δ15N values across species (Kahmen et al. 2008).
We observed strong relationships between foliar δ15N, soil δ15N, and NO2 concentration in all species (Fig. 4.5). Overall, we found strong relationships between soil and foliar δ15N values across the following species: Red maple > red oak > American beech > black birch. The general trends among these species are consistent with previous studies that have found a strong relationship between foliar δ15N and relative rates of soil N cycling in temperate forest ecosystems (Emmett et al. 1998, Gebauer & Schulze 1991, Hogberg 1990, Pardo et al. 2002). We also observed similar relationships between increasing NO2 concentration and foliar δ15N values across the following species: Red maple > red oak > American beech > black birch. These results are consistent with previous work that has documented comparable foliar δ15N patterns in red maple and red spruce stands (Pardo et al. 2007).
Although results suggest that soil δ15N predominantly drives foliar δ15N patterns in several species, foliar uptake of atmospheric reactive N (as indexed by ambient atmospheric NO2 concentration) may also be a strong predictor of variation in foliar δ15N in several species.
The regression model including both soil δ15N and NO2 concentration as predictors explained up to 56% of variation in foliar δ15N patterns (Fig. 4.6). When we accounted for the influence of soil δ15N on patterns of foliar δ15N using multiple regression models, increased NO2 concentration explained a
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large proportion of the remaining variation in foliar δ15N across species. For instance, the strong correlation and explained variance for red maple foliar δ15N with increasing NO2 concentration suggests that this species is likely responding directly to foliar uptake of atmospheric NO2 or indirectly to physiological and biochemical consequences of increasing atmospheric NO2 concentration. Further, the predictive capability of NO2 concentration may highlight species-specific differences in the capacity for foliar uptake of atmospheric reactive N deposition.
Despite the strong relationships documented between foliar δ15N, soil δ15N, and NO2 concentration (Figs. 4.5, 4.6), the remaining variance not accounted for in our statistical analyses suggests that a possible suite of other factors is influencing foliar δ15N patterns. Soil characteristics, available sources, and N cycling processes likely play a strong role in the overall relationship between soil δ15N and foliar δ15N. Species differences in within-plant fractionation and differences in foliar N uptake rates may also have played a role in the variation in foliar δ15N patterns observed in this study. Within-plant fractionations of N can cause variation of 7-12‰ within individual plants (Evans 2001). Species differences in foliar N uptake of atmospheric reactive N could also influence within-plant variation in foliar δ15N (Morikawa et al. 1998). Climate factors such as precipitation, elevation, and land use change have also been shown to exert control on foliar δ15N values (Amundson et al. 2003). Further, the capability of plants to bypass mycorrhizal associations when N is more abundant or a shift in mycorrhizal species as N availability increases across sites cannot be definitely ruled out as a driver of the observed foliar δ15N patterns (Hobbie & Hobbie 2008, Hobbie et al. 2000).
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It is clear that human derived reactive N inputs have impacted rates N isotope cycling in many ecosystems, likely leading to significant changes in the way N is processed and incorporated into ecosystems (Vitousek et al. 1997). However, it is still not well understood how increasing atmospheric reactive N deposition has impacted the foliar δ15N value in ecosystems. In this study, we show that foliar δ15N may be used as an indicator of systematic changes in ecosystem N cycling to characterize the relative influences of various N inputs and transformations available to vegetation, although the complexity of sources, inputs and fractionation still lends itself to great uncertainty in these measurements. Very few, if any studies, have attempted to direct estimate the influence of foliar N uptake on foliar δ15N patterns in trees in temperate forests. Although not conclusive, our results indicate that total canopy uptake, and most likely direct foliar N uptake, of atmospheric reactive N deposition does appear to have an influence on foliar δ15N in several species and can be detected using natural abundance δ15N measurements of plant and soil material. Further work is necessary to clearly identify and quantify the role and δ15N of various plant-available soil N sources such as NO3-, NH4+, and organic N in the soil solution. Additional insight to the contribution of mycorrhizal associations to foliar δ15N patterns is also necessary to adequately assess ecosystem and plant N dynamics on local, regional, and global scales of varying N deposition.
Acknowledgements
This research was funded by a Kieckhefer Adirondack Fellowship and by the Cornell NSF-IGERT in Biogeochemistry and Environmental Biocomplexity,
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Award #DGE 0221658. We thank J. Lavallae, R. Doyle-Morin, D. Lombardozzi and K. Wagner for field support. G. Lovett provided advice throughout the project, A. Eller assisted in statistical analyses, and J. Arcate helped initiate the NYBG sampling. Special thanks to those who have assisted us with this project at the following sites: Adirondack Ecological Center, Fordham Universityʼs Louis Calder Center, the Cary Institute for Ecosystem Studies, Mianus River Gorge Preserve, Clarence Fahnestock State Park, New York Botanical Garden, and Inwood Hill Park.
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