Root-Derived Bicarbonate Assimilation in Plants 9819941245, 9789819941247

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Yanyou Wu Sen Rao

Root-Derived Bicarbonate Assimilation in Plants

Root-Derived Bicarbonate Assimilation in Plants

Yanyou Wu · Sen Rao

Root-Derived Bicarbonate Assimilation in Plants

Yanyou Wu Institute of Geochemistry Chinese Academy of Sciences Guiyang, Guizhou, China

Sen Rao College of Life Sciences and Oceanography Shenzhen University Shenzhen, Guangdong, China

ISBN 978-981-99-4124-7 ISBN 978-981-99-4125-4 (eBook) https://doi.org/10.1007/978-981-99-4125-4 © The Editor(s) (if applicable) and The Author(s), under exclusive license to Springer Nature Singapore Pte Ltd. 2023 This work is subject to copyright. All rights are solely and exclusively licensed by the Publisher, whether the whole or part of the material is concerned, specifically the rights of translation, reprinting, reuse of illustrations, recitation, broadcasting, reproduction on microfilms or in any other physical way, and transmission or information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed. The use of general descriptive names, registered names, trademarks, service marks, etc. in this publication does not imply, even in the absence of a specific statement, that such names are exempt from the relevant protective laws and regulations and therefore free for general use. The publisher, the authors, and the editors are safe to assume that the advice and information in this book are believed to be true and accurate at the date of publication. Neither the publisher nor the authors or the editors give a warranty, expressed or implied, with respect to the material contained herein or for any errors or omissions that may have been made. The publisher remains neutral with regard to jurisdictional claims in published maps and institutional affiliations. This Springer imprint is published by the registered company Springer Nature Singapore Pte Ltd. The registered company address is: 152 Beach Road, #21-01/04 Gateway East, Singapore 189721, Singapore

Preface

Bicarbonate and carbon dioxide are two inorganic carbon sources that can be used by plants. The mutual transformation of the two inorganic carbon sources can be catalyzed by carbonic anhydrase. Plants evolved from aquatic to terrestrial. The utilization of root-derived bicarbonate by plants is the hinge of the coupling of photosynthesis in plants and karstification of carbonates. Although more direct evidence is needed, there is plenty of evidence to prove that bicarbonate photolysis and water photolysis play equally important roles in photosynthetic oxygen evolution. Therefore, the importance and rank of bicarbonate on the physiological functions of plants have been significantly improved, and the physiological effect of bicarbonate on plants is no less than that of carbon dioxide. Studies on the positive physiological effects of bicarbonate and the utilization of root-derived bicarbonate can provide theoretical support for the artificial design of photosynthetic reactors and the excavation of the carbon sink capacity of ecosystems. In this book, we will summarize and discuss inorganic carbon assimilation, focusing on different aspects related to bicarbonate utilization by plants, including (I) general introduction about past, present and future inorganic carbon assimilation, (II) the physiological effects of bicarbonate on plants, (III) the diversity, plasticity and roles of carbonic anhydrase in inorganic carbon utilization by plants, (IV) isotope technology in photosynthesis research and bidirectional isotope tracing culture technology applied in bicarbonate use by plants, and (V) root-derived bicarbonate use in karst habitats. This book can provide a new understanding of photosynthesis and a potential solution for the design of artificial photosynthetic reactors. It is a valuable resource for researchers and students in the fields of plant physiology, agronomy, ecology, paleontology and geochemistry. Guiyang, China Shenzhen, China

Yanyou Wu Sen Rao

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Contents

1 Past, Present and Future of Inorganic Carbon Assimilation . . . . . . . . 1.1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.2 Importance of Photosynthesis of Green Plants . . . . . . . . . . . . . . . . . . 1.3 The Discoveries of Photosynthesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.3.1 Early Studies on Photosynthesis . . . . . . . . . . . . . . . . . . . . . . . . 1.3.2 Modern Studies on Photosynthesis . . . . . . . . . . . . . . . . . . . . . . 1.4 Evolution of Photosynthesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.4.1 Anoxygenic and Oxygenic Photosynthesis . . . . . . . . . . . . . . . 1.4.2 Coevolution of Karstification and Photosynthesis . . . . . . . . . 1.4.3 The Compartmentalization in Photosynthesis . . . . . . . . . . . . . 1.4.4 Implication of Photosynthesis for Biodiversity and Environmental Issues . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2 Physiological Effects of Bicarbonate on Plants . . . . . . . . . . . . . . . . . . . . 2.1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2 Bicarbonate Dominates Photosynthetic Oxygen Evolution . . . . . . . . 2.2.1 Bicarbonate Photolysis, Bicarbonate Effect, and Stoichiometry of the Photo-Reaction and Dark Reaction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2.2 Bicarbonate Photolysis, Anaerobic Photosynthesis, and the Dole Effect . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.3 The Effect of Bicarbonate on Photosynthetic Carbon Assimilation of Higher Terrestrial Plants . . . . . . . . . . . . . . . . . . . . . . . 2.3.1 Multiple Effects of Bicarbonate on Photosynthetic Carbon Assimilation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.3.2 Differences Among Species in the Effects of Bicarbonate on Photosynthesis . . . . . . . . . . . . . . . . . . . . . . 2.3.3 The Role of Bicarbonate as an Alternative Carbon Source for Photosynthesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.3.4 Bicarbonate Roles in Stomatal Movement . . . . . . . . . . . . . . .

1 1 2 3 3 5 6 6 7 9 9 11 15 15 16

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2.3.5 The Role of Bicarbonate in Improving Glucose Metabolism and Stress Tolerance . . . . . . . . . . . . . . . . . . . . . . . 2.3.6 The Role of Bicarbonate in Inducing Chlorosis . . . . . . . . . . . 2.4 Effect of Bicarbonate on Inorganic Nitrogen Metabolism in Plants . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.5 Indirect Effects of Bicarbonate on the Growth and Development of Plants . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.6 The Whole Effect of Bicarbonate on Plant Growth and Development . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.7 Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3 The Diversity, Plasticity and Roles of Carbonic Anhydrase in Inorganic Carbon Utilization in Plants . . . . . . . . . . . . . . . . . . . . . . . . . 3.1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2 Distribution and Properties of Carbonic Anhydrase . . . . . . . . . . . . . . 3.2.1 Ubiquitous Carbonic Anhydrase . . . . . . . . . . . . . . . . . . . . . . . . 3.2.2 Properties of Carbonic Anhydrase . . . . . . . . . . . . . . . . . . . . . . 3.3 Biodiversity of Carbonic Anhydrase . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.3.1 Functional Diversity of Carbonic Anhydrase . . . . . . . . . . . . . 3.3.2 Convergent Evolution of Carbonic Anhydrase . . . . . . . . . . . . 3.3.3 Coordination Diversity of Metal Cofactors and Amino Acid Residues in Carbonic Anhydrase . . . . . . . . . . . . . . . . . . 3.3.4 Diversity of Response and Sensitivity to Inhibitors and Activators . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.4 Diversity of Carbonic Anhydrase in Plants . . . . . . . . . . . . . . . . . . . . . 3.5 Plasticity of Carbonic Anhydrase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.5.1 Response of Carbonic Anhydrase to Water . . . . . . . . . . . . . . . 3.5.2 Response of Carbonic Anhydrase to pH . . . . . . . . . . . . . . . . . 3.5.3 Response of Carbonic Anhydrase to Light Intensity . . . . . . . 3.5.4 Response of Carbonic Anhydrase to Anions . . . . . . . . . . . . . 3.5.5 Response of Carbonic Anhydrase to Cations . . . . . . . . . . . . . 3.5.6 Response of Carbonic Anhydrase to Plant Hormones . . . . . . 3.6 Role of Carbonic Anhydrase on Inorganic Carbon Assimilation in Plants . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.6.1 Unique Thylakoid Carbonic Anhydrase Versus Photosynthetic Oxygen Evolution . . . . . . . . . . . . . . . . . . . . . . 3.6.2 Direct Effect of Carbonic Anhydrase on Photosynthetic Inorganic Carbon Assimilation . . . . . . . . . 3.6.3 Indirect Effect of Carbonic Anhydrase on Photosynthetic Inorganic Carbon Assimilation . . . . . . . . . 3.7 Possible Significance of CA on the Origin and Evolution of Life . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.8 Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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4 Bidirectional Isotope Tracing Culture Technology and Bicarbonate Use by Plants . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.2 Radioactive Isotopic Tracing Technology in Photosynthesis Research . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.2.1 11 C as a Tracer to Study CO2 Fixation in Photosynthesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.2.2 14 C Tracing of the Pathways of CO2 Fixation in Photosynthesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.2.3 14 C Tracing of the Route of Root-Derived Bicarbonate . . . . 4.3 Stable Isotopic Tracing Technology in Photosynthesis Research . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.3.1 Isotopic Exchange and Carbon Isotope Discrimination in Photosynthesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.3.2 18 O Tracing Photosynthetic Oxygen Evolution . . . . . . . . . . . 4.3.3 18 O of Cellulose in Tree Rings Tracing Past Temperatures . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.3.4 13 C as a Tracer and δ13 C in Photosynthesis Research . . . . . . 4.4 Quantitative Determination of the Inorganic Carbon Source and Inorganic Carbon Use Pathways of Microalgae . . . . . . . . . . . . . . 4.4.1 Quantitative Determination of the Share of Different Carbon Sources Used by Microalgae . . . . . . . . . . . . . . . . . . . . 4.4.2 Quantify the Share of the Pathways of Different Inorganic Carbon Use by Microalgae . . . . . . . . . . . . . . . . . . . 4.4.3 Quantify the Direct Carbon Sink and Indirect Carbon Sink . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.5 Quantification of Bicarbonate Utilization by Terrestrial Plants . . . . . 4.5.1 Bidirectional Isotope Tracing Culture . . . . . . . . . . . . . . . . . . . 4.5.2 Quantifying the Daily Average Stable Carbon Isotope Composition of Atmospheric CO2 . . . . . . . . . . . . . . . . . . . . . . 4.5.3 Quantify Bicarbonate Uptake by Plants . . . . . . . . . . . . . . . . . . 4.5.4 Quantification of Root-Derived Bicarbonate Use and Total Photosynthetic Carbon Assimilation of Plants . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.6 Measurement of Total Inorganic Carbon Assimilation Capacity of Plants in Field Habitats . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.7 Conclusions and Outlook . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5 Root-Derived Inorganic Carbon Assimilation by Plants in Karst Environments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.2 How Plants Utilize Soil Dissolved Inorganic Carbon . . . . . . . . . . . . . 5.2.1 Bidirectional Carbon Flow at the Soil–Root Interface . . . . . .

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5.3

5.4

5.5

5.6

5.7

5.2.2 Using CO2 and Bicarbonate as Fertilizers: A Brief History . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.2.3 Root Uptake of Bicarbonate Ion . . . . . . . . . . . . . . . . . . . . . . . . 5.2.4 Anaplerotic Fixation in Roots . . . . . . . . . . . . . . . . . . . . . . . . . . 5.2.5 Transport of Root-Derived Inorganic Carbon in Xylem Sap . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.2.6 Assimilation of Xylem-Transported Carbon Sources Through Corticular Photosynthesis . . . . . . . . . . . . . . . . . . . . . 5.2.7 Assimilation of Xylem-Transported Carbon Sources via Leaf Photosynthesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Root-Derived Bicarbonate Assimilation by Plants Under Simulated Karst Environments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.3.1 Revisiting the Simulation Experiments . . . . . . . . . . . . . . . . . . 5.3.2 Bicarbonate Stress on Photosynthetic Inorganic Carbon Assimilation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.3.3 Interactive Effect of Bicarbonate Excess and Water Stress on Photosynthetic Inorganic Carbon Assimilation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.3.4 Stimulation of Bicarbonate on the Plants’ Total Carbon Gain and the Dynamics of Nonstructural Carbohydrates . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Characteristics of the Karst Environment . . . . . . . . . . . . . . . . . . . . . . . 5.4.1 Karst and Karstification . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.4.2 Karst Hydrochemical and Hydrodynamic Processes . . . . . . . 5.4.3 Karst Landform . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.4.4 Karst Soil . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.4.5 Karst Habitats and Vegetation . . . . . . . . . . . . . . . . . . . . . . . . . . Quantification of Bicarbonate Assimilation by Plants in Karst Habitats . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.5.1 What Can We Learn from Laboratory Experiments? . . . . . . . 5.5.2 Potential Carbon Sources for Leaf Photosynthesis in Karst Habitats . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.5.3 Theoretical Models to Estimate the Contribution of Soil DIC to Leaf Total Photosynthesis in Karst Habitats . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Isotopic Evidence for Plant Use of Soil DIC in Karst Environments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.6.1 Utilization of Soil DIC by Plants in Karst Habitats: Essential Questions to Be Addressed . . . . . . . . . . . . . . . . . . . . 5.6.2 Characteristics of Soil DIC and CO2 in Karst Habitats . . . . . 5.6.3 Disprepancy Between Predicted and Measured δ13 C of Newly Formed Photosynthates . . . . . . . . . . . . . . . . . . . . . . . 5.6.4 Relationship Between δA - δWSOM and f DIC_soil . . . . . . . . . . . Bicarbonate Assimilation in Karst Habitats . . . . . . . . . . . . . . . . . . . . .

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5.7.1 Species-Specific Induced Variation in Plant Use of Soil Bicarbonate in Karst Environments . . . . . . . . . . . . . . . 5.7.2 Spatial Variability in Root-Derived Inorganic Carbon Utilization by Plants in Karst Habitats . . . . . . . . . . . . . . . . . . . 5.7.3 Temporal Variability in Root-Derived Inorganic Carbon Utilization by Plants in Karst Habitats . . . . . . . . . . . . 5.7.4 Uncertainties Associated with the Estimation of Contribution of Soil DIC . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.8 Eco-Physiological and Biogeochemical Significance of Bicarbonate Assimilation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.8.1 Potential Strategy for Plants Adapting to Karst Environments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.8.2 Coupling of Photosynthesis and Karstification and Its Impact on the Terrestrial Carbon Sink . . . . . . . . . . . . . . . . . . . 5.9 Conclusions and Prospects . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Chapter 1

Past, Present and Future of Inorganic Carbon Assimilation

Abstract Photosynthesis is the most important biochemical reaction. It co-evolved and developed with Earth, driving the biogeochemical cycles of all elements on Earth. It is the only chemical process that can convert light energy into chemical energy and use light energy to supplement the Earth’s internal energy that is gradually depleted. This chapter introduces the discovery course of photosynthesis and pursues the trace of bicarbonate use by plants from these discoveries. The evolution of photosynthesis is explored, and the coupling process and development trend of photosynthesis and karstification are discussed. In addition, this chapter also discusses the current ecological environmental problems of related to photosynthesis and proposes the factors to be considered in the design of artificial photosynthesis systems. Keywords Inorganic carbon use · Photosynthesis · Evolution · Kastification–photosynthesis coupling · Bicarbonate photolysis · Artificial photosynthesis system

1.1 Introduction There are a variety of metabolic pathways in plants, including carbon metabolism, water metabolism, nitrogen metabolism, sulfur metabolism, phosphorus and other mineral nutrition metabolism. Carbon metabolism is the most important among the above metabolic pathways. This is because carbon is the main skeleton of all organic compounds, and more than 90% of the dry matter in plants is organic compounds. The assimilation of inorganic carbon is the basis of life survival and development in carbon metabolism. Life on the Earth would be impossible without inorganic carbon assimilation. Assimilation of inorganic carbon refers to the process of autotrophic organisms absorbing inorganic carbon and then reducing it into organic matter under the action of reductants. Inorganic carbon assimilation of autotrophic organisms includes bacterial chemosynthesis and photosynthesis of bacteria and green plants. Generally, it is believed that the photosynthesis of green plants is the most common and assimilates the largest amount of inorganic carbon. However, growing evidence demonstrates that © The Author(s), under exclusive license to Springer Nature Singapore Pte Ltd. 2023 Y. Wu and S. Rao, Root-Derived Bicarbonate Assimilation in Plants, https://doi.org/10.1007/978-981-99-4125-4_1

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the contribution of bacterial photosynthesis and chemosynthesis to the synthesis of organic matter cannot be underestimated, especially in some specific environments.

1.2 Importance of Photosynthesis of Green Plants It is so important that it is no exaggeration to say that plants, animals and human beings cannot survive, reproduce and develop, and the colorful and vibrant earth biosphere will become bald, gray and lifeless without the photosynthesis of plants. If the great potential of photosynthesis is not fully released, the urgent problems of food, resources, climate and environment encountered by human society will be difficult to solve. Generally, photosynthesis of green plants refers to the process by which plants assimilate inorganic carbon, produce organic matter, release oxygen, and convert light energy into chemical energy. The photosynthesis of green plants is the largest and sole biochemical reaction that can take place under normal temperature and pressure. The overall reaction for photosynthesis could be written as the following simple equation according to our recent research outcome (Wu 2023):   light H2 O + CO2 + H2 O → H+ + HCO3 − +H2 O −→(CH2 O) + O2 + H2 O (CH2 O) in the above equation represents carbohydrate. In this equation, inorganic carbon (CO2 and HCO3 − ) is in the most oxidized state of carbon, while (CH2 O) is in the relatively reduced state of carbon. After the reaction, inorganic carbon is reduced, and the hydroxy groups of water and bicarbonate are oxidized; oxygen in the hydroxy groups of water and bicarbonate is in a state of reduction, and O2 is in a state of oxidation. Therefore, photosynthesis is an oxidation–reduction reaction that fixes 114.3 kcal free energy from sunlight for 1 mol of carbon dioxide fixation (Krishnamurthy 1981). Each year, the radiant energy that reaches the earth equals approximately 5 × 1020 kcal. Photosynthesis captures approximately 1% of this huge supply of energy, using it to provide the energy that drives all life activities (Raven and Johnson 2002). The energy released by the burning of coal, firewood, gasoline, and by our bodies’ burning of all the food we eat-all, directly or indirectly, has been captured from sunlight by photosynthesis. The energy used by most living cells ultimately comes from the sun, captured by photosynthetic organisms through the process of photosynthesis. Photosynthesis provides food and oxygen for heterotrophic organisms. It is estimated that photosynthesis fixes 224.5 Pg carbon per year on Earth, of which terrestrial plants account for approximately 132.1 Pg, 59%, and aquatic plants account for approximately 92.4 Pg, 41% (Vitousek et al. 1986). Meanwhile, plants constantly absorb carbon dioxide and release oxygen during photosynthesis, in which the oxygen content is always maintained at 21% of the atmosphere. Oxygen provides conditions for aerobic respiration and accumulates and gradually forms the surface ozone

1.3 The Discoveries of Photosynthesis

3

layer of the atmosphere, which can absorb the strong ultraviolet radiation harmful to organisms in sunlight.

1.3 The Discoveries of Photosynthesis 1.3.1 Early Studies on Photosynthesis Currently, people already understand that photosynthesis is so-called that plants take simple, inorganic, raw materials-carbon dioxide from the atmosphere, and water and mineral salts from the soil, in the presence of sunlight, convert them into organic substances. However, before the seventeenth century, people generally believed the view that plants entirely derive their nutrition from humus in the soil under the influence of Aristotle. In 1648, a quantitative experiment of potted willow performed by Flemish physician Jan van Helmont (1579–1644) questioned this view. Van Helmont took a vessel in which he put 200 pounds of dried soil, which he watered with rainwater or distilled water, and he planted therein willow tree of the five pounds. Five years later, the tree weighed 169 pounds, and the dried soil remained the same 200 pounds. His willow experiment demonstrated that the increasing weight of approximately 164 pounds gained by the growing plant came from the water rather than the soil. Some years after Van Helmont carried out his experiment, Robert Boyle (1627–1691) performed similar experiments using squash and cucumber and demonstrated a conclusion similar to Van Helmont’s willow experiment (Krogmann 2005; Hill 2012). Although they did not realize that the organic substance was related to the atmosphere, they at least challenged the view of plant nutrition from the soil, opening the prologue of photosynthesis research. After decades, until 1727, Stephen Hales’s experiment of ‘pneumatic trough’ demonstrated that Van Helmont’s conception of the plant body as a structure built entirely of water was incomplete, and some of the plant’s nutrition was derived from ‘air’. Nevertheless, Hales’s views had little immediate impact on general conceptions of plant nutrition at that time (Krogmann 2005; Hill 2012). Until the 1770s, British minister Joseph Priestley (1733–1804) provided support for Hales’s view that air is important for plants. The discovery that plants absorb carbon dioxide from the air and produce oxygen into the atmosphere is widely credited to Joseph Priestley. In 1771, Priestley carried out an experiment using unique laboratory apparatus devised by himself. He found that the air inside an enclosed glass jar within a sprig of mint would neither extinguish a candle nor was it at all inconvenient to a mouse. Afterwards, he obtained similar results using other plants in place of mint. Consequently, he concluded that plants thrive in air made noxious by the respiration of animals, reverse the effects of breathing, and tend to maintain atmospheric purification. He proposed that plants can air “purify”. Nevertheless, he was not always able to duplicate his experiment that plants “improve” the air (Krogmann 2005; Hill 2012). Upon performing

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1 Past, Present and Future of Inorganic Carbon Assimilation

more than 500 experiments in less than 3 months’ time, Dutch physician Jan IngenHousz (1730–1799) found Priestley’s experiments not able to be duplicated because Priestley was unaware of the requirement for light during his experiments. Ingen-Housz carried out many experiments using Priestley’s laboratory apparatus modified by providing more intense illumination in 1796. He determined that the effect of the improving air would depend upon the greater or less exposure of the plant to the light and found that it is the sun’s light, not its heat, that brought out the effect (Reed 1950). Although considering that plants obtain their carbon mainly from carbon dioxide in the atmosphere rather than from the soil solution, he did not demonstrate the stoichiometric relationship between carbon dioxide absorbed and oxygen produced by plants under sunlight. In 1782, Jean Senebier (1742–1809) provided evidence that plants must have access to carbon dioxide to produce oxygen, demonstrated that the amount of oxygen produced by a plant is proportionally related to the amount of carbon dioxide absorbed, and concluded the importance of carbon dioxide in the photosynthesis of plants. In addition, he hypothesized that carbon dioxide dissolved in water is the source of the carbon in plants (Krogmann 2005; Hill 2012) (This is the earliest thought sprout that plants use root-derived bicarbonate!). Nevertheless, during his lifetime, Senebier was unable to consider the role of water in Van Helmont’s conception and the stoichiometric relationship between organic substances and water. Fortunately, his compatriot, Nicholas-Théodore de Saussure (1767–1845), took over the relay baton of photosynthesis research. De Saussure had a broad research field of vision. He comprehensively considered the various sources of plant nutrition from the atmosphere, water, and soil rather than aerial nutrition. He elaborated on quantitative experiments and made precise measurements of gas exchange. He found that growing plants produce more organic substances than can be accounted for by the amount of assimilated carbon dioxide and the production of oxygen and that the extra weight gain comes from the water. Therefore, he concluded that water is a reagent that directly participates in photosynthesis and that plants obtain all their carbon from carbon dioxide in the atmosphere rather than from soil (Krogmann 2005; Hill 2012). Although he, like Ingen-Housz or Senebier, holds that the oxygen released by plants from the carbon dioxide rather than from the water, he established the photosynthetic stoichiometric relationship among organic substances, oxygen, water and carbon dioxide. Ultimately, the chemical equation of photosynthesis can now be written as follows: CO2 + H2 O + light

green plants

−→

organic substance + O2

Afterwards, in 1845, German, a physician, Julius Robert Mayer (1814–1878), proposed that plants convert light energy into chemical energy during photosynthesis according to the First Law of Thermodynamics. In 1864, German plant physiologist and botanist Julius von Sachs (1832–1897) demonstrated that starch grains within leaves are the first visible product of photosynthesis and proved that chlorophyll is involved in photosynthesis. In 1905, an English plant physiologist, Frederick Frost

1.3 The Discoveries of Photosynthesis

5

Blackman (1866–1947), performed quantitative experiments on the rates of photosynthesis and proposed the concept of light-limited and dark-limited photosynthesis, which led to later Warburg’s finding of light and dark reactions in photosynthesis (Blackman and Matthaei 1905). In 1913, German chemist Richard Wilstatter (1872– 1942) ultimately demonstrated that chlorophyll in leaves plays an active role in photosynthesis through detailed chemical investigations on chlorophyll (Krogmann 2005). In 1924, J. H. Priestley discovered that plants produced cane sugar, a carbohydrate, during photosynthesis (Priestley 1924). Hereto, the chemical equation of photosynthesis could now be written as follows: chlorophyll

CO2 + H2 O + light −→ C(H2 O) + O2

1.3.2 Modern Studies on Photosynthesis What is oxygen of above equation evolved from water or carbon dioxide, Robert Hill (1899–1991) seems to have given a satisfactory answer. During the period of 1937–1939, Hill demonstrated that oxygen evolution and carbon dioxide fixation into carbohydrates seem to be separate processes according to his experiments on the chloroplast reaction (Hill 1937, 1939). In fact, Hill’s experiments only prove that photosynthetic oxygen evolution can be independent of the process of carbon dioxide fixation but do not prove that there is no internal relationship between carbon dioxide fixation and photosynthetic oxygen evolution. How did carbon dioxide incorporate into carbohydrates during photosynthesis? American scientist Melvin Calvin (1912–1997) and his coworkers provide a perfect solution. During the early 1950s, Melvin Calvin and his colleagues traced the path of carbon through different stages of photosynthesis using 14 C as a tracer. They found the first stable product of CO2 reduction, phosphoglyceraldehyde, and the acceptor of CO2 , ribulose bisphosphate, that there was a cycle to regenerate the acceptor (Calvin and Benson 1949; Bassham and Calvin 1962; Calvin 1962). In 1954, American scientist Daniel Arnon (1910–1994) and his coworkers demonstrated that light drives the synthesis of adenosine triphosphate (ATP) by isolated chloroplasts, which have the capacity for photosynthetic phosphorylation coupled with the reduction of ferricyanide. Even in the dark, chloroplasts can transform carbon dioxide into sugar. The light reaction produces assimilatory power to push the dark reaction, which uses assimilatory power to convert CO2 into carbohydrates (Arnon et al. 1954a, b). In 1960, Robin Hill and Fay Bendall proposed a ‘Z’-scheme to describe a detailed, explicit relation between two separate light reactions in photosynthesis (Hill and Bendall 1960). In the same year, Rajni Govindjee, Eugene Rabinowitch and Jan B. Thomas discovered the two-light effect of the Hill reaction in intact algal cells (Govindjee et al. 1960; Govindjee and Rabinowitch 1960). In 1961, Peter Mitchell (1920–1992) stated the chemiosmotic theory, in which a proton motive force couples

6

1 Past, Present and Future of Inorganic Carbon Assimilation

electron transfer to ATP synthesis in photosynthetic phosphorylation. During the period of 1965–1966, Hugo Kortschak, Hal Hatch, C. R. Slack and others discovered a new carboxylation reaction and pathway (hereafter called the C4 pathway) in photosynthesis (Kortschak et al. 1965; Hatch and Slack 1966). In 1971, Ogren and Bowes demonstrated that ribulose diphosphate carboxylase had the double feature of carboxylase and oxygenase and discovered that the enzyme can catalyze the oxygenation of ribulose-1,5-bisphosphate (RuBP) to produce phosphoglycolate, the first step in photorespiration (Ogren & Bowes 1971; Bowes et al. 1971). Subsequently, advances in the molecular mechanisms of photosynthesis were rapid, and we will not go into further detail here. Along the track of study in photosynthesis (Krogmann 2005; Hill 2012), it does not exclude that plants directly use bicarbonate as a carbon source. Not only rainwater or distilled water but also bicarbonate dissolved in it may nourish the willow in Van Helmont’s experiment (Vapaavuori and Pelkonen 1985; Vuorinen et al. 1992). Submerged leaves may absorb not only carbon dioxide but bicarbonate dissolved in water for photosynthesis in Ingen-Housz’s and Senebier’s experiments. In de Saussure’s experiment, the extra weight gained by plants may come from the water as well as bicarbonate dissolved in it. The bicarbonate effect in the Hill reaction was discovered by many scientists (Warburg and Krippahl 1958; Walker and Hill 1967; Walker et al. 1971; Stemler and Govindjee 1973; Wydrzynski and Govindjee 1975; Stemler 1980; Blubaugh and Govindjee 1986); it cannot be explained as oxygen originating from carbon dioxide but may be interpreted as oxygen originating from bicarbonate except water (Wu 2023). In the future, an increasing number of studies will hopefully confirm that plants can directly use bicarbonate as a substrate, not just water, and bicarbonate photolysis and water photolysis equally account for photosynthetic oxygen evolution.

1.4 Evolution of Photosynthesis 1.4.1 Anoxygenic and Oxygenic Photosynthesis According to Kluyver (1931), the most general form in all metabolic processes in organisms can be expressed as follows: H2 A + B → A + H2 B On the basis of this hypothesis, photosynthetic reactions can then be regarded as a generalized inorganic carbon assimilation process: light

H2 A + IC −→ 2A + CH2 O IC presented in the above formulation represents inorganic carbon, including carbon dioxide, bicarbonate, carbonic acids, and so on. The formulation implies that different

1.4 Evolution of Photosynthesis

7

photosynthetically active organisms require different final reductions. The anoxygenic photosynthesis of organisms such as Thiorhodaceae requires H2 S as a hydrogen donor for carbon assimilation (van Niel 1935). The equation of the inorganic carbon assimilation process in Thiorhodaceae can be expressed as follows: light

H2 S + IC −→ 2S + CH2 O However, the oxygenic photosynthetic activity as displayed by green plants has revealed that fundamentally, this process can be expressed by the equation: light

H2 O + IC −→ O2 + CH2 O Photosynthesis arose early in Earth’s history, perhaps before 3500 million years, and the earliest forms of photosynthetic life were almost certainly anoxygenic (Des Marais 2000). All current evidence from isotopes, signature molecules and fossils has demonstrated that anoxygenic photosynthesis occurred before the development of oxygenic photosynthesis (Blankenship 2010; Hohmann-Marriott and Blankenship 2011). The evolution of photosynthesis from anaerobic to aerobic organisms is involved in Earth’s atmosphere. Many workers have suggested that atmospheric oxygen first rose to appreciable levels at approximately 2000 million years, and the atmosphere in the early Archean contained little or no free oxygen (Kasting 1993). The anaerobic organisms thrived in a weakly reducing primitive atmosphere and resided in the biosphere of an atmosphere containing carbon dioxide and nitrogen and little or no free oxygen, where anoxygenic photosynthesis occurred. Oxygenic photosynthesis occurred when atmospheric oxygen levels rose and carbon dioxide levels declined.

1.4.2 Coevolution of Karstification and Photosynthesis Some species of rocks, such as silicate rock and carbonate rock, undergo weathering under the action of carbon dioxide and water. Silicate rock is first dissolved to form carbonate rock, which can be expressed in the following equation: CO2 + Ca(Mg)SiO3 + H2 O  Ca(Mg)CO3 + H2 SiO3 The carbonate rock then continued to undergo weathering, which can be expressed in the following equation:   CO2 + Ca(Mg) CO3 + H2 O  Ca2+ Mg2+ + 2HCO3 −

8

1 Past, Present and Future of Inorganic Carbon Assimilation

Therefore, the general equation of silicate rock dissolution can be expressed as follows:   2CO2 + Ca(Mg)SiO3 + 2H2 O  Ca2+ Mg2+ + 2HCO3 − +H2 SiO3 Both silicate rock and carbonate rock can be dissolved by consuming carbon dioxide and water and eventually form bicarbonate according to the equation of silicate or carbonate dissolution. The chemical dissolution of rocks, also called karstification, is a process for absorbing carbon dioxide on Earth, similar to photosynthesis. The interaction between karstification and photosynthesis has led to the evolution and development of the Earth’s biosphere. The composition of Earth’s atmosphere reflects the coupling of karstification and photosynthesis (Wu and Wu 2022). Higher levels of atmospheric CO2 have been observed in geological history during periods of lagging photosynthesis development. Stromatolites (a type of carbonate rock) formed by photosynthetic microorganisms occur in 3460 MA (Des Marais 2000). Before approximately 2000 MA, a small amount of photosynthetic organisms, cyanobacteria, resulted in little photosynthesis. However, karstification was extremely strong due to the high concentration of carbon dioxide. As a result, the concentration of CO2 in the atmosphere of Earth decreased rapidly, but the oxygen level was very low from 3500 to 2000 MA. From 2000 to 75 Ma, due to the continuous evolution and reproduction of plants, photosynthesis on Earth was constantly enhanced, and the coupling degree of karstification and photosynthesis was also constantly increased, resulting in the continuous reduction in carbon dioxide concentration and the continuous increase in oxygen in the atmosphere (Berner 1998; Moulton and Berner 1998; Knoll and Nowak 2017). Karstification and photosynthesis were completely coupled during the period from 75 Ma to Holocene (pre-Industrial Revolution), when carbon dioxide in the atmosphere remained at a constant level (Hart 1978; Kasting 1993; Des Marais 2000; Dismukes et al. 2001; Pagani et al. 2009; Halevy and Bachan 2017). After the Industrial Revolution, buried fossil fuels were exploited on a large scale, generating a significant quantity of CO2 in a short period of time, while plant adaptive evolution tended to lag behind, likely resulting in the decoupling of karstification and photosynthesis, which eventually resulted in a rise in global atmospheric CO2 . The atmospheric CO2 concentrations have improved by 50%, from 280 ppm just before the Industrial Revolution to 421 ppm presently. The global average temperature has climbed by approximately 1 °C since 1850–1900 as atmospheric CO2 concentrations have increased. The rise of global atmospheric carbon dioxide has become the most concerning problem today (Wu and Wu 2022). However, after the decoupling of karstification and photosynthesis, plants will naturally evolve new adaptive mechanisms and strategies to improve photosynthesis to adapt to the changing environment, thereby establishing a new coupling of karstification and photosynthesis. It takes a long time, even tens of thousands of years, which is too long for human beings. Therefore, reducing CO2 emissions from human activities and building an artificial photosynthetic system are the only ways to promote rapid coupling of karstification and photosynthesis.

1.4 Evolution of Photosynthesis

9

1.4.3 The Compartmentalization in Photosynthesis The evolution of photosynthesis is also reflected in the compartmentalization of the photoreaction with the dark reaction in photosynthesis. In anoxygenic photosynthetic bacteria, such as Rhodobacter sphaeroides, with weak cellular compartmentalization, accomplishing the connection between photo-reaction and CO2 fixation is the combined effect of carbonic anhydrase and phosphoenolpyruvate carboxylase (Park et al. 2017). In C3 plants, the light reaction and dark reaction in chloroplasts have temporary and spatial separation to a certain extent. The light reaction is the basis of the dark reaction. The light reaction takes place in the thylakoid membranes of chloroplasts, while the dark reaction takes place in the matrix of chloroplasts. Photolyzing bicarbonate releases oxygen and gathers carbon dioxide into the Calvin cycle, connecting the photo-reaction and inorganic carbon fixation. In C4 plants, a large temporary and spatial separation occurs in the light reaction and the dark reaction of photosynthesis. The light reaction takes place in the thylakoid membranes of mesophyll cells, while the dark reaction, the fixation of inorganic carbon, takes place in the chloroplasts of bundle sheath cells. The connecting light reaction and the fixation of carbon dioxide is the carboxylation reaction of phosphoenolpyruvate carboxylase combined with bicarbonate. Photosynthesis in crassulacean acid metabolism (CAM) plants has a day and night temporal separation in carbon fixation in addition to the same spatiotemporal separation between light reaction and dark reaction in C3 and C4 plants. Similarly, the carboxylation reaction of phosphoenolpyruvate carboxylase combined with bicarbonate connects the temporal gap between day and night in carbon fixation (Cushman 2001; Silvera et al. 2010).

1.4.4 Implication of Photosynthesis for Biodiversity and Environmental Issues There are multifarious light and dark reactions. The different combinations of light and dark reactions display different photosynthetic types. The diversity of photosynthesis incorporating the diversity of respiratory metabolism forms biodiversity on a biochemical basis. Although photosynthesis is diverse, the basic equation is similar. According to our viewpoint (Wu 2023), bicarbonate may be incorporated into the light reaction of photosynthesis and then broken to release oxygen and generate reducing equivalents. The equation of the light reaction in photosynthesis was written as follows: H2 O + H+ + HCO− 3 + 2ADP + 2Pi

+ 2NADP+ + hv → O2 + 2ATP + 2NADPH + 4H+ + CO2

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1 Past, Present and Future of Inorganic Carbon Assimilation

In the above equation, ADP denotes adenosine diphosphate, Pi denotes inorganic phosphate, NADP+ denotes oxidized nicotinamide adenine dinucleotide phosphate, and NADPH denotes reduced nicotinamide adenine dinucleotide phosphate. The equation of the dark reaction in photosynthesis was written as follows: 2ATP + 2NADPH + 4H+ + CO2 → CH2 O + 2ADP + 2Pi + 2NADP+ + H2 O The total equation of photosynthesis is as follows: H2 O + H+ + HCO3 − +hv → CH2 O + O2 + H2 O According to the total equation of photosynthesis, plants assimilate 1 mol of bicarbonate to return 1 mol water into the environment. Bicarbonate is then photosynthetically assimilated by plants and finally fixed in carbohydrates and releases water to the living environment. Therefore, we can say that photosynthesis is not only the driving force of the carbon cycle in the biosphere but also the driving force of the water cycle. Photosynthesis in photosynthetic organisms is broadly defined as a series of processes in which light energy is converted to chemical energy used for the biosynthesis of organic cell substances (Gest 1993). Carbon reduction depends not only on photosynthesis but also on the reduction of nitrogen, sulphur and iron (Hill and Scarisbrick 1940; Grant and Canvin 1970; de la Guardia and Alcántara 1996; Kopriva et al. 1999). During photosynthesis, light energy is converted into chemical energy as ATP concomitant with phosphate assimilation, resulting in inorganic carbon assimilation. The reducing equivalent generated by the light reaction competitively utilizes the reduction of bicarbonate, nitrate, sulfate and iron (Brunold and Suter 1984; Schmutz and Brunold 1984; Dofing et al. 1989; Koprivova et al. 2000). The reduction of 1 mol of nitrate or sulfate requires 8 mol of protons and electrons provided by 4 mol of bicarbonate photolysis or 2 mol of bicarbonate with water photolysis, respectively (Salsac et al.1987; Khan et al. 2010). In the biosphere, bicarbonate photolysis and water photolysis are hard currency of anion salt assimilation. Iron reduction and ATP formation as phosphate assimilation are hard currency of cation salt assimilation or chemical energy, respectively (Yi and Guerinot 1996). Therefore, photosynthesis drives the cycle of almost all elements, such as nitrogen, sulfur, phosphorus and iron, in the biosphere. The evolution and future of the environment depends on the evolution and development of photosynthesis in biosphere plants. At present, climatic and environmental problems are increasingly present in the biosphere with increasing human activities. These problems mainly include increasing emissions of greenhouse gases, global warming, acid rain, air, water and land pollution, and sharp reductions in biodiversity. Photosynthetic assimilation is a powerful weapon against changes in these climatic

References

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environments. Photosynthesis can not only consume water and absorb carbon dioxide but also reduce a more harmful greenhouse gas, carbonyl sulphide (COS) (Crane and Bar 1977). Plants not only directly assimilate hydrogen sulfide, sulfur dioxide, sulfate, nitrate, and phosphate to reduce air, water and soil pollution but also reduce the accumulation of some polluting elements in water media and soils by building themselves using the chemical energy generated by photosynthesis to obtain various metal elements (Sekiya et al. 1982; Alhendawi et al. 1997). Meanwhile, bicarbonate assimilation by plants results in increasing alkalinity neutralizing the acidity brought by acid rain. In addition, the various light and dark reactions as well as the changes adapted to the environment in photosynthesis, such as the formation of the C3 -C4 intermediate plant type (Monson and Moore 1989), increase the living space and survival chances, thus increasing biodiversity. At present, the current photosynthetic capacity of plants can no longer solve the practical problems of the ecological environment caused by human activities. Therefore, it is urgent for human beings to establish an artificial photosynthesis system to solve these complex ecological environmental problems. By coordinating light reactions and dark reactions, selecting proton and electron donors, designing various coupling reactions of light reactions and dark reactions, and making full use of bicarbonate photolysis and water photolysis, an efficient light energy converter (artificial photosynthesis system) can be built.

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Calvin M, Benson AA (1949) The path of carbon in photosynthesis IV: the identity and sequence of the intermediates in sucrose synthesis. Science 109(2824):140–142 Crane FL, Barr R (1977) Stimulation of photosynthesis by carbonyl compounds and chelators. Biochem Biophys Res Commun 74(4):1362–1368 Cushman JC (2001) Crassulacean acid metabolism. A plastic photosynthetic adaptation to arid environments. Plant Physiol 127(4):1439–1448 de la Guardia MD, Alcántara E. (1996). Ferric chelate reduction by sunflower (Helianthus annuus L.) leaves: influence of light, oxygen, iron-deficiency and leaf age. J Exp Bot 47(5): 669–675. Des Marais DJ (2000) When did photosynthesis emerge on Earth? Science 289(5485):1703–1705 Dismukes GC, Klimov VV, Baranov SV, Kozlov YN, DasGupta J, Tyryshkin A (2001) The origin of atmospheric oxygen on Earth: the innovation of oxygenic photosynthesis. Proc Natl Acad Sci USA 98(5):2170–2175 Dofing SM, Penas EJ, Maranville JW (1989) Effect of bicarbonate on iron reduction by soybean roots. J Plant Nutr 12(6):797–802 Gest H (1993) Photosynthetic and quasi-photosynthetic bacteria. FEMS Microbiol Lett 112(1):1–5 Govindjee, Rabinowitch E (1960) Two forms of chlorophyll a in vivo with distinct photochemical functions. Science 132(3423):355–356 Govindjee R, Thomas JB, Rabinowitch E (1960) “Second Emerson Effect” in the Hill reaction of chlorella cells with quinone as oxidant. Science 132(3424):421–421 Grant BR, Canvin DT (1970) The effect of nitrate and nitrite on oxygen evolution and carbon-dioxide assimilation and the reduction of nitrate and nitrite by intact chloroplasts. Planta 95(3):227–246 Halevy I, Bachan A (2017) The geologic history of seawater pH. Science 355(6329):1069–1071 Hart MH (1978) The evolution of the atmosphere of the Earth. Icarus 33:23–39 Hatch MD, Slack CR (1966) Photosynthesis in sugar cane leaves: a new carboxylation reaction and the pathway of sugar formation. Biochem J 101(1):103–111 Hill JF (2012) Early pioneers of photosynthesis research. Photosynthesis. Springer, Dordrecht, pp 771–800 Hill R (1937) Oxygen evolution by isolated chloroplasts. Nature 139:881–882 Hill R (1939) Oxygen produced by isolated chloroplasts. Proc R Soc Lond-B 127(847):192–210 Hill R, Bendall F (1960) Function of the cytochrome components in chloroplasts: a working hypothesis. Nature 186:136–137 Hill R, Scarisbrick R (1940) The reduction of ferric oxalate by isolated chloroplasts. Proc R Soc Lond-B 129(855):238–255 Hohmann-Marriott MF, Blankenship RE (2011) Evolution of photosynthesis. Annu Rev Plant Biol 62:515–548 Kasting JF (1993) Earth’s early atmosphere. Science 259(5097):920–926 Khan MS, Haas FH, Samami AA, Gholami AM, Bauer A, Fellenberg K, Reichelt M, H¯ansch R, Mendel RR, Meyer AJ, Wirtz M, Hell R (2010) Sulfite reductase defines a newly discovered bottleneck for assimilatory sulfate reduction and is essential for growth and development in Arabidopsis thaliana. Plant Cell 22(4):1216–1231 Kluyver AJ (1931) Chemical activities of micro-organisms. London University Press. Knoll AH, Nowak MA (2017) The timetable of evolution. Sci Adv 3:e1603076 Kopriva S, Muheim R, Koprivova A, Trachsel N, Catalano C, Suter M, Brunold C (1999) Light regulation of assimilatory sulphate reduction in Arabidopsis thaliana. Plant J 20(1):37–44 Koprivova A, Suter M, den CampRO BC, Kopriva S (2000) Regulation of sulfate assimilation by nitrogen in Arabidopsis. Plant Physiol 122(3):737–746 Kortschak HP, Hartt CE, Burr GO (1965) Carbon dioxide fixation in sugarcane leaves. Plant Physiol 40(2):209–213 Krishnamurthy S (1981) Hydrothermal vents and light-independent living systems. J Chem Educ 58(12):981–981 Krogmann D (2005) Discoveries in oxygenic photosynthesis (1727–2003): a perspective. Discoveries in photosynthesis. Springer, Dordrecht, pp 63–105

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Chapter 2

Physiological Effects of Bicarbonate on Plants

Abstract In this chapter, we focus on the positive physiological role of bicarbonate in the process of inorganic carbon assimilation. The significance of bicarbonate photolysis for photosynthetic oxygen evolution is discussed, the catalytic effect of inorganic carbon on photosynthetic oxygen evolution is explained, and the formation mechanism of the ‘bicarbonate effect’ and the ‘Dole effect’ is clarified. The effects of bicarbonate on the photosynthesis of different plants were analysed. In addition, this chapter also summarizes the carbon source function of bicarbonate and the regulation of bicarbonate on stomatal movement. The effect of bicarbonate on glucose metabolism was analysed. Finally, this chapter also analyses the overall effect of bicarbonate on plant growth and development. It is concluded that bicarbonate dominates photosynthetic oxygen evolution, coordinates the photo-reaction and dark reaction, as well as stomatal movement, acts as an alternative carbon source for photosynthesis, and improves the glucose metabolism and stress tolerance of plants. Based on the comprehensive impact of bicarbonate on many physiological and biochemical processes, bicarbonate is no less important to plants than carbon dioxide. Due to the transformation between bicarbonate and carbon dioxide, fully developing the positive role of bicarbonate in plants will help to realize carbon neutralization quickly and effectively. Keywords Inorganic carbon utilization · Bicarbonate photolysis · Bicarbonate effect · Photosynthesis · Alternative carbon source · Stomatal movement · Glucose metabolism

2.1 Introduction Carbon is one of the most basic and important elements of life. There are two main interchangeable forms of inorganic carbon in nature: carbon dioxide and bicarbonate. The evolution of the biosphere and organisms is involved in the biochemistry cycle of carbon (Kasting 1993; des Marais 2000; Dismukes et al. 2001). Inorganic carbon plays an important role in the growth, reproduction and evolution of organisms. Carbon and living substances composed of carbon perform all functions in living © The Author(s), under exclusive license to Springer Nature Singapore Pte Ltd. 2023 Y. Wu and S. Rao, Root-Derived Bicarbonate Assimilation in Plants, https://doi.org/10.1007/978-981-99-4125-4_2

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2 Physiological Effects of Bicarbonate on Plants

organisms. Therefore, bicarbonate, as the main form of inorganic carbon, must play an extremely important role, which is even no less than that of carbon dioxide in life. Bicarbonate, which is closely related to the origin of life and the evolution of the biosphere, plays an important role in the assimilation of inorganic carbon. Other physiological activities, such as the absorption and utilization of life elements, the transmission of genetic information, the transformation of energy and growth and development, are based on inorganic carbon assimilation. Bicarbonate is not only absorbed by roots from soil but also derived from the conversion of atmospheric CO2 diffusion through the cell wall into the cytosol, which dissolves in the cell wall or apoplast water. They can act alone or overlay on the growth and development of plants. Generally, low bicarbonate promotes and high bicarbonate inhibits the growth and development of plants.

2.2 Bicarbonate Dominates Photosynthetic Oxygen Evolution 2.2.1 Bicarbonate Photolysis, Bicarbonate Effect, and Stoichiometry of the Photo-Reaction and Dark Reaction 2.2.1.1

Photosynthetic Oxygen Evolution Versus Bicarbonate Effect

It is an indisputable fact that bicarbonate can greatly stimulate photosynthetic oxygen evolution. However, many heavy oxygen (18 O) isotope labelling experiments still make people generally believe that water is the sole substrate for photosyntheic oxygen evolution (Ruben et al. 1941; Stemler and Radmer 1975; Radmer and Ollinger 1980; Hillier et al. 2006). Recently, we found through comprehensive analysis of evidence from various aspects that bicarbonate can also be used as a substrate for photosynthetic oxygen release (Wu 2021a). Photosystem II (PSII) oxygen evolution is not only dependent on bicarbonate but also controlled by bicarbonate. Water photolysis and bicarbonate photolysis each account for half of photosynthetic oxygen evolution (Wu 2023) (Fig. 2.1). The preceding viewpoint that bicarbonate photolysis plays a controlling role in photosynthetic oxygen evolution of the joint efforts of bicarbonate and water perfectly explains the two outstanding biological issues. The first is that bicarbonate is more important than water in photosynthetic oxygen evolution (bicarbonate effect), and the second is that plants can precisely control photosynthetic oxygen evolution and CO2 assimilation according to the stoichiometric relationship of 1:1 (mol/mol). The reaction equation of photosynthetic oxygen evolution, 2H2 O + CO2 → H2 O + H+ − + + HCO− 3 → O2 + 4e + 4H + CO2 , shows that CO2 clearly acts as a part-time ‘catalyst’ to accelerate photosynthetic oxygen evolution in photosynthetic organisms (bicarbonate effect), which Warburg and Krippahl referred to as “activated CO2 ”

2.2 Bicarbonate Dominates Photosynthetic Oxygen Evolution

17

Fig. 2.1 Bicarbonate dominates photosynthetic oxygen evolution. Bicarbonate photolysis is the precondition of water photolysis. Bicarbonate photolysis and water photolysis are carried out in a 1:1 (mol/mol) stoichiometric relationship (from Wu 2023)

(Warburg and Krippahl 1958). Bicarbonate-mediated CO2 formation was found on both the donor and acceptor sides of PSII, confirming the role of CO2 as a concurrent “catalyst” and the validity of the above reaction equation (Shevela et al. 2020). The catalytic principle of the catalyst is to reduce the activation energy. Thermodynamic evidence also supports the catalytic role of inorganic carbon in photosynthetic oxygen evolution. The standard free energy variance was 37.3 kcal/mol for water photolysis, 24.8 kcal/mol for bicarbonate photolysis and 31.1 kcal/mol for the entire process of photosynthetic oxygen evolution (Dismukes et al. 2001). Therefore, photosynthetic oxygen evolution dominated by bicarbonate photolysis plays two important roles in the life activities of plants. One is to accelerate the process of photosynthetic oxygen release, and the other is to concentrate carbon dioxide for ribulose-1,5-bisphosphonate carbonoxylase/oxygenase (Rubisco) of C3 species or phosphoenolpyruvate carbonoxylase (PEPC) of C4 species (Wu 2021b).

18

2 Physiological Effects of Bicarbonate on Plants

Fig. 2.2 The coordination between light reaction and dark reaction of photosynthesis by photosynthetic oxygen evolution from plants (from Wu 2023). Note ➀(➁) is bicarbonate photolysis (water photolysis). ➀ is equal to ➁.➂(➃) is carbon similation (carbon dissimilation) (water photolysis). ➂ is equal to ➃

2.2.1.2

Photosynthetic Oxygen Evolution Versus Stoichiometry of the Photoreaction-Dark Reaction

It is obvious that water photolysis cannot provide a carbon source for carbon dioxide assimilation. However, the simultaneous even photolysis of bicarbonate and water ensures that oxygen and CO2 are released in equal amounts, which precisely regulates the stoichiometric ratio between the photo-reaction and dark reaction from the beginning of photosynthesis and presets a 1:1 stoichiometric relationship between assimilation and dissimilation from the headstream of the carbon biogeochemical cycle (Fig. 2.2, from Wu 2023).

2.2.2 Bicarbonate Photolysis, Anaerobic Photosynthesis, and the Dole Effect 2.2.2.1

Photosynthetic Oxygen Evolution Versus Anoxygenic Photosynthesis

Simultaneously, the above viewpoint can also explain sulfid photolysis in anaerobic photosynthetic bacteria during photosynthesis, as well as the Dole effect, which is nearly 24‰ more enriched in 18 O of atmospheric O2 than seawater. When both inorganic carbon and water are present, anoxygenic photosynthetic microorganisms such as sulfur bacteria do not photolyze either water or bicarbonate. They live in an acidic environment where inorganic carbon exists as carbon dioxide rather than bicarbonate. Carbon dioxide itself cannot be used as a substrate for photolysis to yield oxygen, and only bicarbonate can be used as a substrate for photolysis. Without the photolysis of bicarbonate, the precondition of water photolysis will be

2.3 The Effect of Bicarbonate on Photosynthetic Carbon Assimilation …

19

lost. Therefore, these bacteria can only photolyze acidic substances (such as hydrogen sulfide) without oxygen evolution (Cohen et al. 1975).

2.2.2.2

Photosynthetic Oxygen Evolution Versus Dole Effect

The Dole effect is the most powerful counter evidence that photosynthetic oxygen release comes from water photolysis by plants. Assuming that the oxygen in the atmosphere only comes from the water photolysis of photosynthetic organisms, there ought to be no Dole effect in that the 18 O content of atmospheric O2 is almost 24‰ higher than that of seawater. We calculate the enrichment of 18 O in atmospheric O2 according to bicarbonate photolysis and water photolysis to release oxygen equivalently. The enrichment of 18 O in atmospheric O2 relative to water and bicarbonate (0.2043) is vastly near the actual content of 18 O in the atmosphere (0.2041) (Wu 2023). This shows that the oxygen in the atmosphere comes evenly from water and bicarbonate in water and proves that the view that half of photosynthetic oxygen evolution originated from water photolysis and half from bicarbonate photolysis is true and reliable.

2.3 The Effect of Bicarbonate on Photosynthetic Carbon Assimilation of Higher Terrestrial Plants 2.3.1 Multiple Effects of Bicarbonate on Photosynthetic Carbon Assimilation Bicarbonate in the soil has both negative and positive effects on the carbon assimilation and growth of terrestrial plants. On the one hand, excessive bicarbonate may produce ion toxicity; lead to a high-pH environment and physiological drought; affect the expression of genes and protein synthesis; promote stomatal closure; inhibit the activities of key enzymes; change the synthesis and transport of metabolites, electrolyte balance and other physiological and biochemical activities; and hinder the normal growth and metabolic carbon assimilation of plants. On the other hand, suitable bicarbonate may become the carbon source of carbon assimilation, support the function and stability of PSII, provide protons and electrons, serve as substrates for some carboxylases such as acetyl-CoA carboxylase (ACCase, EC.6.4.1.2) and phosphoenolpyruvate carboxylase (PEPC, EC 4.1.1.31), stimulate stomatal opening, enhance drought resistance, and benefit other physiological and biochemical functions. Bicarbonate has a complex effect on photosynthetic carbon assimilation, which varies with the concentrations of bicarbonate, plant species and growing environments.

20

2 Physiological Effects of Bicarbonate on Plants

2.3.2 Differences Among Species in the Effects of Bicarbonate on Photosynthesis 2.3.2.1

Net Photosynthetic Rate

The photosynthetic response to bicarbonate varied with concentration, plant species and treatment duration. Four plant species (two Moraceae plant species, Broussonetia papyrifera (L.) Vent. (Bp) and Morus alba L. (Ma); two Cruciferae plant species, Orychophragmus violaceus (Ov) and Brassica napus (Bn)), were cultured under different bicarbonate levels (BC-0, BC-1, BC-5, and BC-10) in modified Hoagland culture solution (the pH was adjusted to 8.0 ± 0.2) with normal Zn levels or no Zn for 50 days. BC-0, BC-1, BC-5, and BC-10 were the treatments with bicarbonate added at 0, 1, 5, and 10 mM L−1 sodium bicarbonate, respectively. Photosynthetic parameters, such as net photosynthetic rate (Pn), transpiration (E), stomatal conductance (gs), and intercellular CO2 concentration (Ci), were measured using a portable LI-6400XT photosynthesis system (LI-COR Inc., Lincoln, NE, USA). The fourth youngest fully expanded leaf from the top was used for measurement between 9:00 and 11:00 a.m. The CO2 concentration, photosynthetically active radiation, and temperature during measurement were 380 µmol mol−1 , 600 µmol m−2 s−1 and 25 °C, respectively. Table 2.1 shows the differential response of the net photosynthetic rate in four plant species under different bicarbonate levels in the presence of Zn. From Table 2.1, in the presence of Zn, the Pn of all plant species increased with the duration of treatments except BC-10 for Bp and Ma. The Pn in the short duration of treatment had little change with the concentration of bicarbonate added to the culture solution. However, the response of Pn in the long duration of treatment by bicarbonate was different in the four plant species. The Pn on Day 50 had the largest difference among bicarbonate levels. The suitable concentration of bicarbonate for Pn in Bp, Ov and Bn was 5 mM L−1 and that for Pn in Ma was 1 mM L−1 . However, in the absence of Zn, bicarbonate restrained the photosynthetic carbon assimilation of all species. Table 2.2 shows the differential response of the net photosynthetic rate in four plant species under different bicarbonate levels in the absence of Zn. From Table 2.2, the Pn of all species significantly decreased with the addition of bicarbonate and duration of treatment. The Pn on Day 50 had the largest decrease under 10 mM L−1 bicarbonate added to the culture solution.

2.3.2.2

Water-Use Efficiency

Table 2.3 shows the differential response of the water-use efficiency (WUE) in four plant species under different bicarbonate levels in the presence of Zn. In the presence of Zn, the response of Pn in the short duration of treatment by bicarbonate was different in the four plant species. The WUE of all plant species increased with the duration of treatments except BC-10 for two Moraceae plant species. However, the WUE on Day 50 had the largest difference among bicarbonate levels. The suitable

2.3 The Effect of Bicarbonate on Photosynthetic Carbon Assimilation …

21

Table 2.1 The net photosynthetic rate (Pn, µmol m–2 s–1 ) in four plant species under different bicarbonate levels in the presence of Zn. The data were determined on days 10 (Day 10), 20 (Day 20), 30 (Day 30), 40 (Day 40), and 50 (Day 50). The values in brackets represent the percentage compared to the control (BC-0) Plant species Bp

Ma

Ov

Bn

Treatment

Day 10

Day 20

Day 30

Day 40

Day 50

BC-0

7.55 (100)

7.89 (100)

7.98 (100)

8.46 (100)

9.21 (100)

BC-1

8.16 (108)

8.36 (106)

9.32 (117)

10.45(124)

11.10(121)

BC-5

8.93 (118)

9.47 (120)

9.93 (124)

11.10(131)

13.29(144)

BC-10

7.67 (102)

6.88 (87)

6.35 (80)

5.97 (71)

4.87 (53)

BC-0

6.59 (100)

6.63 (100)

6.80 (100)

7.00 (100)

7.88 (100)

BC-1

6.77 (103)

7.86 (119)

7.97 (117)

8.95 (128)

10.76(137)

BC-5

7.01 (106)

7.22 (109)

7.78 (114)

8.21 (117)

9.32 (118)

BC-10

7.67 (116)

5.97 (90)

5.97 (88)

5.20 (74)

3.63 (46)

BC-0

7.01 (100)

7.96 (100)

8.37 (100)

9.01 (100)

9.44 (100)

BC-1

7.41 (106)

8.19 (103)

9.58 (114)

10.29(114)

12.45(132)

BC-5

7.77 (111)

8.75 (110)

10.78(129)

11.43(127)

13.34(141)

BC-10

6.91 (99)

8.20 (103)

8.52 (102)

9.43 (105)

9.82 (104)

BC-0

5.70 (100)

6.96 (100)

7.57 (100)

8.48 (100)

8.93 (100)

BC-1

6.12 (107)

7.41 (106)

8.24 (109)

9.37 (110)

10.63(119)

BC-5

6.30 (111)

7.79 (112)

9.38 (124)

10.62(125)

12.00(134)

BC-10

6.17 (108)

7.62 (109)

8.27 (109)

8.97 (106)

8.01 (90)

concentration of bicarbonate for the WUE of Bp, Ov and Bn was 5 mM L−1 and that for the WUE of Ma was 1 mM L−1 . However, in the absence of Zn, bicarbonate decreased the WUE of all species except on Day 10. Table 2.4 shows the differential response of WUE in four plant species under different bicarbonate levels in the absence of Zn. As shown in Table 2.4, the WUE of all species significantly decreased with the addition of bicarbonate and duration of treatment except on Day 10. Similarly, the WUE on Day 50 had the largest decrease under 10 mM L−1 bicarbonate added to the culture solution. Therefore, we can conclude that bicarbonate also inhibits the WUE of plants in the absence of Zn.

2.3.2.3

Chlorophyll Fluorescence

Table 2.5 shows the response of the PSII photochemical efficiency (Fv /Fm ) in four plant species under different bicarbonate levels in the presence of Zn. In the presence of Zn, Fv /Fm was not affected by bicarbonate in the four plant species except BC-10 for Bp on days 30, 40, and 50 and for Bn on Day 50. Therefore, we can conclude that bicarbonate had little effect on the Fv /Fm of plants in the presence of Zn.

22

2 Physiological Effects of Bicarbonate on Plants

Table 2.2 The net photosynthetic rate (Pn, µmol m–2 s–1 ) in four plant species under different bicarbonate levels in the absence of Zn. The data were determined on days 10 (Day 10), 20 (Day 20), 30 (Day 30), 40 (Day 40), and 50 (Day 50). The values in brackets represent the percentage compared to the control (BC-0) Plant species Bp

Ma

Ov

Bn

Treatment

Day 10

Day 20

Day 30

Day 40

Day 50

BC-0

8.79 (100)

7.50 (100)

7.08 (100)

5.57 (100)

4.67 (100)

BC-1

7.75 (88)

6.90 (92)

5.06 (72)

4.87 (87)

3.88 (83)

BC-5

7.03 (80)

6.45 (86)

6.30 (89)

4.17 (75)

3.57 (76)

BC-10

7.19 (82)

5.47 (73)

4.87 (69)

2.96 (53)

2.65 (57)

BC-0

6.73 (100)

6.70 (100)

6.40 (100)

6.23 (100)

5.08 (100)

BC-1

6.45 (96)

5.98 (89)

4.75 (74)

4.17 (67)

3.13 (62)

BC-5

5.50 (82)

4.84 (72)

3.96 (62)

3.58 (57)

2.87 (56)

BC-10

5.45 (81)

3.70 (55)

3.82 (60)

2.73 (44)

1.66 (33)

BC-0

13.34(100)

12.00(100)

10.29(100)

8.37 (100)

6.12 (100)

BC-1

12.94 (97)

11.53 (96)

9.01 (88)

7.01 (84)

5.20(85)

BC-5

12.77 (96)

11.19 (93)

8.75 (85)

6.38 (76)

4.88(80)

BC-10

12.45 (93)

10.63(89)

7.47(73)

5.77 (69)

3.45 (56)

BC-0

7.45 (100)

6.55 (100)

6.09 (100)

4.70 (100)

3.31 (100)

BC-1

7.41 (99)

6.17 (94)

5.24 (86)

4.13 (88)

2.60 (78)

BC-5

7.35 (99)

6.03 (92)

4.94 (81)

3.63 (77)

2.19 (66)

BC-10

6.55 (88)

5.58 (85)

4.37 (72)

2.24 (48)

1.39 (42)

Table 2.6 shows the differential response of Fv /Fm in four plant species under different bicarbonate levels in the absence of Zn. From Table 2.6, in the absence of Zn, the Fv /Fm of all four plant species in the short duration of treatment on days 10 and 20 had little change with the concentration of bicarbonate added to the culture solution except for Ma. However, during the long duration of treatment, bicarbonate decreased the Fv /Fm of all four plant species. Table 2.7 shows the differential response of the PSII electron transport rate (ETR) in four plant species under different bicarbonate levels in the presence of Zn. From Table 2.7, in the presence of Zn, the ETR in Bp, Ma and Bn had little change with the duration of treatments and concentration of bicarbonate added to the culture solution except for BC-10. However, the response of ETR in Ov differed in the duration of treatments and concentration of bicarbonate. The ETR of Ov on Day 10, Day 20, and Day 30 slightly increased with the concentration of bicarbonate added to the culture solution except for BC-10, and that over a long duration of treatment significantly increased under BC-1 and BC-5. However, in the absence of Zn, bicarbonate restrains the ETR of all species. Table 2.8 shows the differential response of the net photosynthetic rate in four plant species under different bicarbonate levels in the absence of Zn. From Table 2.8, the ETR of all species significantly decreased with the addition of bicarbonate and duration of treatment. The ETR on Day 50 had the largest decrease under 10 mM

2.3 The Effect of Bicarbonate on Photosynthetic Carbon Assimilation …

23

Table 2.3 The water-use efficiency (WUE, µmol(CO2 ) mol–1 (H2 O)) in four plant species under different bicarbonate levels in the presence of Zn. The data were determined on days 10 (Day 10), 20 (Day 20), 30 (Day 30), 40 (Day 40), and 50 (Day 50). The values in brackets represent the percentage compared to the control (BC-0) Plant species Bp

Ma

Ov

Bn

Treatment

Day 10

Day 20

Day 30

Day 40

Day 50

BC-0

4.94 (100)

5.34 (100)

5.54 (100)

5.99 (100)

6.81 (100)

BC-1

4.71 (96)

4.78 (90)

6.81 (123)

7.35 (123)

8.70 (128)

BC-5

5.44 (110)

6.34 (119)

7.10 (128)

8.25 (138)

10.54(155)

BC-10

5.54 (112)

4.40 (82)

3.90 (70)

3.31 (55)

2.66 (39)

BC-0

4.04 (100)

4.29 (100)

4.51 (100)

5.51 (100)

5.82 (100)

BC-1

4.26 (105)

5.92 (138)

6.74 (149)

7.60 (138)

8.03 (138)

BC-5

4.31 (107)

5.99 (140)

6.53 (145)

7.10 (129)

7.53 (129)

BC-10

4.65 (115)

3.24 (76)

2.91 (65)

2.47 (45)

2.01 (35)

BC-0

6.19 (100)

6.83 (100)

8.23 (100)

8.74 (100)

9.63 (100)

BC-1

6.83 (110)

7.69 (113)

8.49 (103)

9.17 (105)

11.30(117)

BC-5

7.62 (123)

8.69 (127)

9.14 (111)

11.28(129)

12.80(133)

BC-10

7.42 (120)

8.46 (124)

8.93 (108)

9.31 (107)

9.89 (103)

BC-0

6.10 (100)

7.03 (100)

7.79 (100)

8.24 (100)

8.93 (100)

BC-1

6.14 (101)

7.25 (103)

7.81 (100)

8.74 (106)

10.04(112)

BC-5

7.16 (117)

8.16 (116)

9.63 (124)

10.65(129)

11.18(125)

BC-10

7.04 (115)

7.98 (113)

8.32 (107)

7.90 (96)

7.71 (86)

L− concentration bicarbonate added to the culture solution. Bp decreased ETR more than Ma, and Ov decreased ETR more than Bn under the treatment with the same concentration of bicarbonate. This result indicated that the ETR of Bp and Ov with Zn deficiency was more sensitive to high concentrations of bicarbonate than that of Ma and Bn.

2.3.2.4

Chlorophyll Content

Photosynthesis is closely related to the content of chlorophyll, and bicarbonate is bound to affect the photosynthetic efficiency of unit chlorophyll. Table 2.9 shows the differential response of the chlorophyll (Chl) content in four plant species under different bicarbonate levels in the presence of Zn. From Table 2.9, in the presence of Zn, bicarbonate slightly increased the Chl content in Bp and Ma with a short duration of treatment or under low concentrations of bicarbonate added to the culture solution. The Chl content in Bp and Ma on Day 50 had the largest decline under BC-10. The Chl content of Ov decreased with the concentration of bicarbonate added to the culture solution. The Chl content of Bn changed little with the duration of treatments and the concentration of bicarbonate added to the culture solution, except for BC-10.

24

2 Physiological Effects of Bicarbonate on Plants

Table 2.4 The water-use efficiency in four plant species under different bicarbonate levels in the absence of Zn. The data were determined on days 10 (Day 10), 20 (Day 20), 30 (Day 30), 40 (Day 40), and 50 (Day 50). The values in brackets represent the percentage compared to the control (BC-0) Plant species Bp

Ma

Ov

Bn

Treatment

Day 10

Day 20

Day 30

Day 40

Day 50

BC-0

4.45 (100)

4.26 (100)

3.90 (100)

3.68 (100)

3.38 (100)

BC-1

4.52 (102)

4.11 (97)

3.44 (88)

3.13 (85)

2.91 (86)

BC-5

4.26 (96)

3.89 (91)

2.93 (75)

2.47 (67)

2.04 (60)

BC-10

4.51 (101)

3.32 (78)

2.47 (63)

1.92 (52)

1.49 (44)

BC-0

4.81 (100)

4.42 (100)

3.90 (100)

3.13 (100)

2.93 (100)

BC-1

4.91 (102)

3.76 (85)

3.57 (91)

3.31 (106)

2.27 (77)

BC-5

3.90 (81)

2.93 (66)

1.55 (40)

1.45 (46)

1.47 (50)

BC-10

3.44 (72)

2.02 (46)

1.78 (46)

1.16 (37)

0.90 (31)

BC-0

11.32(100)

8.16 (100)

7.55 (100)

5.70 (100)

5.13 (100)

BC-1

11.09(98)

8.17 (101)

6.64 (88)

5.25 (92)

3.95(77)

BC-5

9.89 (87)

7.98 (98)

5.38 (71)

4.04 (71)

3.41(66)

BC-10

8.41(74)

7.28 (89)

5.05 (67)

3.61(63)

3.01(59)

BC-0

7.98 (100)

7.11 (100)

6.24 (100)

5.74 (100)

4.15 (100)

BC-1

8.32 (104)

6.92 (97)

5.71 (91)

4.15 (72)

3.12 (75)

BC-5

7.81 (98)

6.51 (92)

5.76 (92)

5.23 (91)

2.92 (70)

BC-10

7.66 (96)

5.33 (75)

4.91 (79)

3.81 (66)

1.88 (45)

Table 2.10 shows the differential response of the chlorophyll (Chl) content in four plant species under different bicarbonate levels in the absence of Zn. From Table 2.10, in the absence of Zn, bicarbonate increased the Chl content in Bp and Ma with a short duration of treatment or under a long concentration of bicarbonate added to the culture solution. The Chl content in Bp and Ma on Day 50 had the largest decline under BC-10. The Chl content of Ov and Bn decreased with the concentration of bicarbonate added to the culture solution.

2.3.2.5

The Specific Net Photosynthetic Rate

The photosynthetic efficiency per unit chlorophyll is called the specific net photosynthetic rate (SPn). Compared with the net photosynthetic rate, SPn can reflect the photosynthetic capacity of plants better than the net photosynthetic rate. Table 2.11 shows the differential response of SPn in four plant species under different bicarbonate levels in the presence of Zn. From Table 2.11, in the presence of Zn, SPn of Bp increased with the concentration of bicarbonate added to the culture solution except for BC-10. BC-1 significantly increased and BC-10 greatly decreased SPn for Ma. The suitable concentration of bicarbonate for SPn in Bp was 5 mM L−1 and that for SPn in Ma was 1 mM L−1 . Bicarbonate significantly increased SPn and stimulated

2.3 The Effect of Bicarbonate on Photosynthetic Carbon Assimilation …

25

Table 2.5 The PSII photochemical efficiency (Fv /Fm ) in four plant species under different bicarbonate levels in the presence of Zn. The data were determined on days 10 (Day 10), 20 (Day 20), 30 (Day 30), 40 (Day 40), and 50 (Day 50); the values in brackets represent the percentage compared to the control (BC-0) Plant species Bp

Ma

Ov

Bn

Treatment

Day 10

Day 20

Day 30

Day 40

Day 50

BC-0

0.754(100)

0.764(100)

0.779(100)

0.787(100)

0.788(100)

BC-1

0.765(101)

0.770(101)

0.782(100)

0.788(100)

0.792(100)

BC-5

0.768(102)

0.772(101)

0.779(100)

0.785(100)

0.788(100)

BC-10

0.755(100)

0.771(101)

0.700(90)

0.750(95)

0.743(94)

BC-0

0.727(100)

0.729(100)

0.727(100)

0.734(100)

0.737(100)

BC-1

0.726(100)

0.722(99)

0.731(101)

0.739(101)

0.742(101)

BC-5

0.728(100)

0.729(100)

0.731(101)

0.737(100)

0.740(100)

BC-10

0.728(100)

0.736(101)

0.730(100)

0.727(99)

0.718(97)

BC-0

0.658(100)

0.673(100)

0.704(100)

0.721(100)

0.724(100)

BC-1

0.678(103)

0.684(102)

0.711(101)

0.724(100)

0.729(101)

BC-5

0.681(103)

0.695(103)

0.710(101)

0.715(99)

0.730(101)

BC-10

0.669(102)

0.690(103)

0.708(101)

0.731(99)

0.720(101)

BC-0

0.688(100)

0.703(100)

0.707(100)

0.717(100)

0.724(100)

BC-1

0.698(101)

0.706(100)

0.712(101)

0.721(101)

0.725(100)

BC-5

0.703(102)

0.712(101)

0.716(101)

0.721(101)

0.749(103)

BC-10

0.686(100)

0.718(102)

0.712(101)

0.702(98)

0.679(94)

photosynthesis in Ov and Bn. The suitable concentration of bicarbonate for SPn in Ov and Bn was 5 mM L−1 in the short duration of treatment and that in the long duration of treatment was 10 mM L−1 . Table 2.12 shows the differential response of SPn in four plant species under different bicarbonate levels in the absence of Zn. From Table 2.12, in the absence of Zn, SPn of all four plant species had similar responses compared with Pn except Ov and Bn in the short duration of treatment on Day 10 and Day 20. Bicarbonate can significantly increase SPn and stimulate photosynthesis for Ov and Bn on Day 10. To summarize, a low bicarbonate increase and high bicarbonate decrease photosynthesis and water-use efficiency of plants in the presence of Zn. However, in the absence of Zn, bicarbonate significantly decreases the photosynthesis and wateruse efficiency of plants. In the presence of Zn, bicarbonate had little effect on PSII photochemical efficiency and significantly changed the electron transport of PSII. However, in the absence of Zn, bicarbonate inhibits the electron transport of PSII. The above effect of bicarbonate on photosynthetic carbon assimilation of higher terrestrial plants is related to the following mechanisms and functions. On the one hand, bicarbonate may be involved in photosynthetic oxygen evolution, providing (accepting) protons and electrons for PSII, and carbon dioxide for Rubisco (ribose-1,5-bisphosphonate carboxylase/oxygenase) (Shevela et al. 2020;

26

2 Physiological Effects of Bicarbonate on Plants

Table 2.6 The PSII photochemical efficiency (Fv /Fm ) in four plant species under different bicarbonate levels in the absence of Zn. The data were determined on days 10 (Day 10), 20 (Day 20), 30 (Day 30), 40 (Day 40), and 50 (Day 50); the values in brackets represent the percentage compared to the control (BC-0) Plant species Bp

Ma

Ov

Bn

Treatment

Day 10

Day 20

Day 30

Day 40

Day 50

BC-0

0.731(100)

0.729(100)

0.727(100)

0.727(100)

0.726(100)

BC-1

0.724(99)

0.718(98)

0.712(98)

0.706(97)

0.702(97)

BC-5

0.705(96)

0.717(98)

0.618(85)

0.687(95)

0.674(93)

BC-10

0.700(96)

0.715(98)

0.600(83)

0.679(93)

0.659(91)

BC-0

0.717(100)

0.697(100)

0.691(100)

0.678(100)

0.667(100)

BC-1

0.711(99)

0.712(102)

0.693(100)

0.591(87)

0.640(96)

BC-5

0.718(100)

0.690(99)

0.681(99)

0.508(75)

0.599(90)

BC-10

0.676(94)

0.636(91)

0.572(83)

0.517(76)

0.521(78)

BC-0

0.764(100)

0.751(100)

0.744(100)

0.743(100)

0.734(100)

BC-1

0.749(98)

0.743(99)

0.739(99)

0.734(99)

0.725(99)

BC-5

0.748(98)

0.743(99)

0.733(99)

0.709(95)

0.695(95)

BC-10

0.745(98)

0.717(96)

0.713(96)

0.677(91)

0.640(87)

BC-0

0.769(100)

0.767(100)

0.761(100)

0.759(100)

0.754(100)

BC-1

0.773(100)

0.763(100)

0.754(99)

0.752(99)

0.733(99)

BC-5

0.768(100)

0.758(99)

0.749(98)

0.743(98)

0.735(98)

BC-10

0.755(98)

0.746(97)

0.745(98)

0.728(96)

0.724(96)

Wu 2021a,b, 2023; Shitov 2022); on the other hand, bicarbonate may be involved in bicarbonate-mediated stomatal movement and other positive and negative effects.

2.3.3 The Role of Bicarbonate as an Alternative Carbon Source for Photosynthesis Various adverse conditions, including excessive bicarbonate, may lead to a decrease in stomatal conductance or stomatal closure of plant leaves, which seriously limits atmospheric carbon dioxide into the inorganic carbon pool of photosynthetic cells, hinders the photosynthesis of plants, makes it difficult to dissipate the light energy absorbed by plant leaves, and causes damage to photosynthetic organs. However, plants have an alternative solution to overcome this dilemma. Dissolved inorganic carbon (DIC), including bicarbonate and carbon oxide, obtained from rhizospheric soils can be used to replenish the inorganic carbon pool (IC pool) to fulfil photosynthetic carbon assimilation, eliminate the “idling” phenomenon of “photosynthetic machines”, alleviate the situation of inorganic carbon sources and water shortages under stomatal closure, promote stomatal opening, and improve the utilization of

2.3 The Effect of Bicarbonate on Photosynthetic Carbon Assimilation …

27

Table 2.7 The PSII electron transport rate (ETR) in four plant species under different bicarbonate levels in the presence of Zn. The data were determined on days 10 (Day 10), 20 (Day 20), 30 (Day 30), 40 (Day 40), and 50 (Day 50); the values in brackets represent the percentage compared to the control (BC-0) Plant species Bp

Ma

Ov

Bn

Treatment

Day 10

Day 20

Day 30

Day 40

Day 50

BC-0

28.33(100)

30.38(100)

34.90(100)

36.44(100)

40.27(100)

BC-1

28.43(100)

31.75(105)

33.23(95)

38.03(104)

40.50(101)

BC-5

29.33(104)

32.75(108)

36.87(105)

39.23(108)

41.70(104)

BC-10

29.17(103)

31.00(102)

33.40(95)

35.18(97)

31.03(77)

BC-0

26.65(100)

28.47(100)

29.53(100)

32.58(100)

34.47(100)

BC-1

26.98(101)

29.23(103)

30.30(103)

34.28(105)

36.78(107)

BC-5

27.87(105)

29.55(104)

30.60(104)

33.13(102)

38.00(110)

BC-10

26.27(99)

28.50(100)

33.35(113)

30.48(94)

24.72(72)

BC-0

20.76(100)

24.55(100)

27.60(100)

28.06(100)

32.05(100)

BC-1

22.42(108)

25.50(104)

32.24(117)

33.50(119)

43.85(137)

BC-5

24.92(120)

26.08(106)

28.80(104)

38.75(138)

41.18(128)

BC-10

19.54(94)

22.15(90)

23.90(87)

29.43(106)

33.06(103)

BC-0

28.50(100)

29.48(100)

32.66(100)

35.45(100)

43.03(100)

BC-1

29.08(102)

30.18(102)

33.45(102)

39.73(112)

46.48(108)

BC-5

29.88(105)

34.90(118)

35.85(110)

38.34(108)

47.53(110)

BC-10

28.42(100)

30.30(103)

32.36(99)

37.73(106)

34.30(80)

atmospheric carbon dioxide by plants (Wu et al. 2018; Banerjee et al. 2019). The alternative utilization of atmospheric carbon dioxide and rhizosphere inorganic carbon sources by plants is shown in Fig. 2.3. The details about bicarbonate utilization by plants are shown in Chapter 5. Here, we display the utilization of bicarbonate by plants with an example as follows. We studied the response of two Moraceae plant species, Broussonetia papyrifera (L.) Vent. and Morus alba L. to excessive bicarbonate in net photosynthetic rate (determined by Li-6400 portable photosynthesis measurement system) and bicarbonate-use rate, chlorophyll fluorescence, and water-use efficiency. The bicarbonate-use capacity of the plants was quantified using bidirectional isotope labelling tracer technology (see Chapter 4) and comparing the compositions of hydrogen isotopes (Wu and Xing 2012). The response of these two Moraceae plant species to excessive bicarbonate in the photosynthetic assimilation of atmospheric carbon dioxide and rhizosphere inorganic carbon added in culture solution in the form of bicarbonate is shown in Table 2.13. Excessive bicarbonate significantly decreases the net CO2 assimilation rate (An), water-use efficiency (WUE), and stomatal conductance (gs ) of the two species, and B. papyrifera shows a greater decrease in these parameters. The total photosynthetic rate (Pn ' ) includes the net CO2 assimilation rate and the bicarbonate-use rate (BU). Excessive bicarbonate also inhibits the bicarbonate-use rate and total photosynthetic

28

2 Physiological Effects of Bicarbonate on Plants

Table 2.8 The PSII electron transport rate (ETR) in four plant species under different bicarbonate levels in the absence of Zn. The data were determined on days 10 (Day 10), 20 (Day 20), 30 (Day 30), 40 (Day 40), and 50 (Day 50); the values in brackets represent the percentage compared to the control (BC-0) Plant species Bp

Ma

Ov

Bn

Treatment

Day 10

Day 20

Day 30

Day 40

Day 50

BC-0

34.38(100)

33.10(100)

26.53(100)

24.75(100)

22.80(100)

BC-1

33.80(98)

26.27(79)

24.40(92)

22.55(91)

20.53(90)

BC-5

29.57(86)

26.65(81)

23.43(88)

21.50(87)

18.70(82)

BC-10

29.05(84)

25.88(78)

23.30(88)

19.33(78)

15.58(68)

BC-0

28.80(100)

26.53(100)

23.33(100)

21.48(100)

18.58(100)

BC-1

28.10(98)

25.28(95)

22.57(97)

21.67(101)

16.62(89)

BC-5

27.05(94)

24.37(92)

22.41(96)

19.78(92)

15.53(84)

BC-10

27.20(94)

23.03(87)

21.38(92)

18.14(84)

14.85(80)

BC-0

40.92(100)

27.62(100)

24.38(100)

20.52(100)

18.87(100)

BC-1

38.25(93)

27.04(98)

22.43(92)

18.70(91)

16.37(87)

BC-5

31.98(78)

23.63(86)

20.48(84)

17.48(85)

14.60(77)

BC-10

28.82(70)

23.32(84)

19.56(80)

15.60(76)

12.27(65)

BC-0

21.76(100)

19.53(100)

18.12(100)

15.48(100)

14.68(100)

BC-1

21.55(99)

18.70(96)

17.54(97)

14.33(93)

13.07(89)

BC-5

21.60(99)

17.20(88)

16.90(93)

13.40(87)

11.70(80)

BC-10

19.90(91)

15.47(79)

12.70(70)

11.28(73)

10.73(73)

rate. Both M. alba and B. papyrifera under control conditions had a greater BU than under bicarbonate treatment. B. papyrifera had a greater BU than M. alba under the same treatment during the same period, and no BU was observed in M. alba under the bicarbonate treatment on day 20. However, B. papyrifera had a great BU even on day 20 under bicarbonate treatment. Compared with the control plants, both M. alba and B. papyrifera did not show marked differences in the Fv /Fm and ϕp values under bicarbonate treatment on day 10. However, on day 20, the Fv /Fm values in M. alba under bicarbonate treatment were significantly higher than those in the control plants, whereas the ϕp values in B. papyrifera under bicarbonate treatment were lower than those in the control plants. It also demonstrated that the short-term effect of bicarbonate treatment on photosynthesis did not involve any damage to the PSII reaction centers (Wu and Xing 2012). Previously, the amount of bicarbonate utilized by plants was usually unclear because of limited measurement methods. However, we can observe that both B. papyrifera and M. alba have a considerable share of bicarbonate use from the above examples. Under the stress of excessive bicarbonate, stomatal conductance decreased significantly, inducing a decline in the net CO2 assimilation rate in B. papyrifera and M. alba. Moreover, the PSII reaction centers were not damaged owing to high bicarbonate use capacity to reduce the bicarbonate content in B. papyrifera but were

2.3 The Effect of Bicarbonate on Photosynthetic Carbon Assimilation …

29

Table 2.9 The chlorophyll (Chl) content in four plant species under different bicarbonate levels in the presence of Zn. The Chl content was determined using SPAD-502 readings (Konica Minolta Sensing Inc., Osaka, Japan). The data were determined on days 10 (Day 10), 20 (Day 20), 30 (Day 30), 40 (Day 40), and 50 (Day 50); the values in brackets represent the percentage compared to the control (BC-0) Plant species

Treatment

Day 10

Day 20

Day 30

Day 40

Bp

BC-0

38.08(100)

41.30(100)

41.75(100)

43.16(100)

45.23(100)

BC-1

41.02(108)

43.20(105)

45.20(108)

46.59(108)

48.56(107)

Ma

Ov

Bn

Day 50

BC-5

43.15(113)

44.48(108)

46.11(110)

42.33(98)

39.22(87)

BC-10

40.33(106)

42.60(103)

37.02(89)

35.12(81)

30.59(68)

BC-0

35.88(100)

39.88(100)

40.21(100)

40.47(100)

40.91(100)

BC-1

36.87(103)

40.94(103)

40.06(100)

40.40(100)

41.08(100)

BC-5

41.44(115)

43.67(110)

45.22(112)

43.81(108)

45.16(110)

BC-10

38.21(107)

41.52(104)

43.05(107)

33.63(83)

31.00(75)

BC-0

43.61(100)

45.38(100)

47.11(100)

48.88(100)

50.72(100)

BC-1

43.17(99)

43.45(96)

44.41(94)

48.53(99)

50.11(99)

BC-5

34.8(80)

40.89(90)

42.29(90)

46.58(95)

49.58(98)

BC-10

32.2(74)

36.84(81)

41.68(88)

37.55(77)

34.03(67)

BC-0

36.01(100)

36.44(100)

37.30(100)

40.05(100)

42.7(100)

BC-1

34.71(96)

35.75(98)

37.20(100)

39.79(99)

44.18(103)

BC-5

32.18(89)

35.69(98)

36.39(98)

38.84(97)

42.51(100)

BC-10

34.54(96)

37.69(103)

33.58(90)

30.44(76)

27.75(65)

damaged owing to no or little bicarbonate use to accumulate bicarbonate gradually in M. alba (Wu and Xing 2012). In fact, the bicarbonate-use capacity of plants is species-specific and depends on the environment. Orychophragmus violaceus L. and Brassica juncea L. are C3 annual herbaceous plants of Cruciferae. They were cultured in Hoagland nutrient solution with the addition of three bicarbonate levels (5, 10 and 15 mM NaHCO3 ) (no adjustment of pH). The proportions of bicarbonate utilized by O. violaceus were 5.28% at 5 mM bicarbonate, 13.27% at 10 mM bicarbonate and 17.31% at 15 mM bicarbonate. Meanwhile, the proportions of bicarbonate utilized by B. juncea were 5.05% at 5 mM bicarbonate, 5.21% at 10 mM bicarbonate and 5.45% at 15 mM bicarbonate (Hang and Wu 2016). Euphorbia lathyris L. is a C4 annual herbaceous plant of the Euphorbiaceae family. E. lathyris, O. violaceus, and B. juncea were grown in substrate with a 1:1 quartz sand to vermiculite mixture (m/m) that was fertilized with modified Hoagland’s nutrient solution (pH 8.1 ± 0.1). Four drought treatments were applied: well-watered (control, 20% moisture), mild drought stress (D1, 17% moisture), moderate drought stress (D2, 14% moisture), and severe drought stress (D3, 11% moisture). After the 180-day growth phase, the share of bicarbonate use in leaves of the three species was different: 9.44% for E. lathyris, 11.45% for O. violaceus,

30

2 Physiological Effects of Bicarbonate on Plants

Table 2.10 The chlorophyll (Chl) content in four plant species under different bicarbonate levels in the absence of Zn. The Chl content was determined using SPAD-502 readings (Konica Minolta Sensing Inc., Osaka, Japan). The data were determined on days 10 (Day 10), 20 (Day 20), 30 (Day 30), 40 (Day 40), and 50 (Day 50); the values in brackets represent the percentage compared to the control (BC-0) Plant species

Treatment

Day 10

Day 20

Day 30

Day 40

Bp

BC-0

39.90(100)

40.70(100)

40.87(100)

41.27(100)

40.66(100)

BC-1

45.70(115)

43.78(108)

42.17(103)

41.07(100)

40.27(99)

Ma

Ov

Bn

Day 50

BC-5

45.13(113)

44.72(110)

42.37(104)

39.60(96)

37.48(92)

BC-10

41.63(104)

41.52(102)

39.00(95)

36.12(98)

32.11(79)

BC-0

34.15(100)

37.06(100)

39.18(100)

39.36(100)

37.88(100)

BC-1

43.18(126)

43.06(116)

41.33(105)

40.92(104)

38.11(101)

BC-5

39.33(115)

40.02(108)

41.86(107)

35.71(91)

30.44(80)

BC-10

42.02(123)

41.21(111)

31.98(82)

27.96(71)

24.84(66)

BC-0

48.40(100)

34.15(100)

32.17(100)

30.30(100)

29.10(100)

BC-1

40.40(83)

32.80(96)

29.55(92)

28.30(93)

25.53(88)

BC-5

43.30(89)

31.75(93)

28.50(89)

27.07(89)

24.87(85)

BC-10

32.90(68)

30.60(90)

26.50(82)

25.70(85)

24.60(85)

BC-0

40.47(100)

38.53(100)

35.65(100)

34.20(100)

32.90(100)

BC-1

39.33(97)

36.75(95)

32.65(92)

31.35(92)

30.15(92)

BC-5

36.13(89)

32.35(84)

31.95(90)

30.45(89)

28.65(87)

BC-10

32.80(81)

30.40(79)

26.57(75)

25.57(75)

21.26(65)

and 10.39% for B. juncea. After a 21-day drought stress phase, the highest share of bicarbonate use was reached for each species: 26.95% for E. lathyris in D3, 26.72% for O. violaceus in D3, and 22.10% for B. juncea in D2 (Wang et al. 2017). Bicarbonate use in response to various adversities is a manifestation of biodiversity, which is closely related to the adaptability of plants to the growth environment. The efficient utilization of bicarbonate by plants plays the role of "killing two birds with one stone" in coping with various adversities. On the one hand, it improves water use efficiency and reduces the water demand of plants; on the other hand, it turns waste (bicarbonate) into wealth (photosynthate) and increases the productivity of plants (Wu et al. 2018).

2.3.4 Bicarbonate Roles in Stomatal Movement 2.3.4.1

Bicarbonate-Mediated Stomatal Movement

Stomata in terrestrial plants, which are formed from two specialized cells in the epidermis (guard cells), are small adjustable pores (Franks & Farqhuar, 2007).

2.3 The Effect of Bicarbonate on Photosynthetic Carbon Assimilation …

31

Table 2.11 The specific net photosynthetic rate (SPn) in four plant species under different bicarbonate levels in the presence of Zn. The data were determined on days 10 (Day 10), 20 (Day 20), 30 (Day 30), 40 (Day 40), and 50 (Day 50). The values in brackets represent the percentage compared to the control (BC-0) Plant species Bp

Ma

Ov

Bn

Treatment

Day 10

Day 20

Day 30

Day 40

Day 50

BC-0

0.198(100)

0.191(100)

0.191(100)

0.196(100)

0.204(100)

BC-1

0.199(100)

0.194(101)

0.206(108)

0.224(114)

0.229(112)

BC-5

0.207(105)

0.213(111)

0.215(113)

0.262(134)

0.339(166)

BC-10

0.190(96)

0.162(85)

0.172(90)

0.170(87)

0.159(78)

BC-0

0.184(100)

0.166(100)

0.169(100)

0.173(100)

0.193(100)

BC-1

0.184(100)

0.192(116)

0.199(118)

0.221(128)

0.262(136)

BC-5

0.169(92)

0.165(100)

0.172(102)

0.187(108)

0.206(107)

BC-10

0.201(109)

0.144(87)

0.139(82)

0.155(89)

0.117(61)

BC-0

0.161(100)

0.175(100)

0.178(100)

0.184(100)

0.186(100)

BC-1

0.172(107)

0.188(108)

0.216(121)

0.212(115)

0.248(134)

BC-5

0.223(139)

0.214(122)

0.255(143)

0.245(133)

0.269(145)

BC-10

0.215(133)

0.223(127)

0.204(115)

0.251(136)

0.289(155)

BC-0

0.158(100)

0.191(100)

0.203(100)

0.212(100)

0.209(100)

BC-1

0.176(112)

0.207(108)

0.221(109)

0.236(111)

0.241(115)

BC-5

0.196(124)

0.218(114)

0.258(127)

0.273(129)

0.282(135)

BC-10

0.179(113)

0.202(106)

0.246(121)

0.295(139)

0.289(138)

Guard cells must operate to ensure an appropriate balance between CO2 uptake for photosynthesis and water loss and ultimately plant water use efficiency. Stomatal opening and closure in terrestrial plants is related to the concentration of inorganic carbon in the environment where plants grow. Low bicarbonate-induced stomatal opening (Mrinalini et al. 1982) and high bicarbonate-mediated stomatal closure (Kolla and Raghavendra 2007) are well known. Increasing evidence shows that stomatal movement is related to anion channels and the enzymes using bicarbonate as a substrate, carbonic anhydrase and phosphoenolpyruvate carboxylase in the guard cells of leaves.

2.3.4.2

CA-Mediated Signals in the Control of Stomatal Movement

Guard cells are the gateway for CO2 , water, and pathogens, while mesophyll cells mainly function in photosynthesis. The different responses of guard cells and mesophyll cells to bicarbonate result in their different functions. Mesophyll cells increased, but guard cells decreased, amino acids, phenylpropanoids, redox metabolites, auxins and cytokinins upon elevated CO2 through supplementation with bicarbonate (Misra et al. 2015). Carbonic anhydrase has been linked to stomatal regulation in terrestrial plants (Sharma et al. 1996; Stimler et al. 2012). Zn-deficient leaves had a stomatal

32

2 Physiological Effects of Bicarbonate on Plants

Table 2.12 The specific net photosynthetic rate (SPn) in four plant species under different bicarbonate levels in the absence of Zn. The data were determined on days 10 (Day 10), 20 (Day 20), 30 (Day 30), 40 (Day 40), and 50 (Day 50). The values in brackets represent the percentage compared to the control (BC-0) Plant species Bp

Ma

Ov

Bn

Treatment

Day 10

Day 20

Day 30

Day 40

Day 50

BC-0

0.220(100)

0.184(100)

0.173(100)

0.135(100)

0.115(100)

BC-1

0.170(77)

0.158(86)

0.120(69)

0.119(88)

0.096(84)

BC-5

0.156(71)

0.144(78)

0.149(86)

0.105(78)

0.095(83)

BC-10

0.173(79)

0.132(72)

0.125(72)

0.082(60)

0.083(72)

BC-0

0.197(100)

0.181(100)

0.163(100)

0.158(100)

0.134(100)

BC-1

0.149(76)

0.139(77)

0.115(71)

0.102(64)

0.082(61)

BC-5

0.140(71)

0.121(67)

0.095(58)

0.100(63)

0.094(70)

BC-10

0.130(60)

0.090(50)

0.119(73)

0.098(62)

0.067(50)

BC-0

0.276(100)

0.351(100)

0.320(100)

0.276(100)

0.210(100)

BC-1

0.320(116)

0.352(100)

0.305(95)

0.248(90)

0.204(97)

BC-5

0.295(107)

0.352(100)

0.307(96)

0.236(85)

0.196(93)

BC-10

0.378(137)

0.347(99)

0.282(88)

0.225(81)

0.140(67)

BC-0

0.184(100)

0.170(100)

0.171(100)

0.137(100)

0.101(100)

BC-1

0.188(102)

0.168(99)

0.160(94)

0.132(96)

0.086(85)

BC-5

0.203(111)

0.186(110)

0.155(90)

0.119(87)

0.076(76)

BC-10

0.200(108)

0.184(108)

0.164(96)

0.087(64)

0.066(65)

Fig. 2.3 Diagrammatic sketch of alternative utilization of atmospheric carbon dioxide and rhizosphere inorganic carbon sources by plants. CH2 O, carbohydrate; RuBP, ribulose-1,5-bisphosphate; Rubisco, ribulose-1,5-bisphosphate carboxylase/oxygenase; PGA, 3-phosphoglycerate; PP, P-type H+ -ATPase; PS II, photosystem II; AE, anion exchanger; IC, inorganic carbon; TR, transporter DIC, dissolved inorganic carbon; ctCA, cytosolic CA; pCA, periplasmic CA; tCA, thylakoid CA; pmCA, cytoplasmic membrane CA

2.3 The Effect of Bicarbonate on Photosynthetic Carbon Assimilation …

33

Table 2.13 The net CO2 -assimilation rate (An, µmol m–2 s–1 ), bicarbonate-use rate (BU, µmol m–2 s–1 ), total photosynthetic rate (Pn ' , µmol m–2 s–1 ), stomatal conductance (gs , mol(H2 O)m–2 s –1 ), water-use efficiency (WUE, µmol(CO2) mol–1 (H2 O)), maximal PSII photochemical efficiency (Fv /Fm ) and photochemical efficiency of open PSII (ϕp ) in the two species under different treatments. The means (SE) (n = 5, 9) followed by different letters in the same parameter of Moraceae plants differ significantly at p ≤ 0.05, according to one-way ANOVA and t test. Control- the treatment without bicarbonate added, BT- the treatment with bicarbonate added to 10 mM/L sodium bicarbonate in the modified Hoagland culture solution (the pH was adjusted to 8.1 ± 0.5, and the concentration of phosphate was 0.25 mM/L rather than 1 mM/ L) for 20 days. The experiments were conducted in a growth chamber under a 12-h photoperiod, 300 µmol m–2 s–1 PPFD, a day/night temperature cycle of 28/20 °C, and 60% relative air humidity. The CO2 concentration, photosynthetically active radiation, and temperature during measurement were 400 µmol mol−1 , 300 µmol m−2 s−1 and 30 °C, respectively. The data were determined on days 10 (Day 10) and 20 (Day 20) (Wu and Xing 2012) Treatments

M. alba Day 10

B. papyrifera Day 20

Day 10

Day 20

Mean (SE)

Mean (SE)

Mean (SE)

Mean (SE)

Control

An

4.08 (0.39)c

4.89 (0.21)b

5.91 (0.34)a

6.13 (0.02)a

BT

An

1.82 (0.32)e

2.55 (0.18)d

2.49 (0.14)de

1.91 (0.10)de

Control

BU

0.73

0.88

2.54

2.64

BT

BU

0.15

0

0.97

0.74

Control

Pn '

4.81

5.77

8.45

8.77

BT

Pn '

1.97

2.55

3.46

2.65

Control

WUE

5.70 (0.48)bc

4.02 (0.43)c

13.14 (1.16)a

WUE

2.81

(0.60)cd

2.68(0.45)cd

1.56

Control

gs

0.05(0.004)b

0.07(0.007)a

0.04(0.005)bc

0.06(0.005)ab

BT

gs

0.03(0.002)c

BT

(0.14)d

6.91 (0.42)b 2.95 (0.39)cd

0.05(0.006)b

0.02(0.002)c

0.03 (0.003)c

Fv /Fm

0.76

(0.01)ab

0.74(0.03)b

0.77(0.02)ab

0.81(0.00)a

BT

Fv /Fm

0.77(0.01)ab

0.79

(0.01)a

0.78(0.01)ab

0.78(0.01)ab

Control

ϕp

0.40 (0.01)c

0.38(0.03)c

0.47(0.04)bc

0.58(0.02)a

BT

ϕp

0.44(0.02)c

0.44(0.01)c

0.53(0.01)ab

0.50(0.00)b

Control

opening smaller than the controls owing to the drastic decrease in carbonic anhydrase activity in the guard cells. Decreased water loss from the leaves of Zn-deficient plants was primarily due to decreased stomatal transpiration (Sharma et al. 1996; Hu et al. 2015). CA-mediated signals play a significant role not only in the control of stomatal movement (Kolbe et al. 2018; Hu et al. 2015) but also in stomatal development in some species (Engineer et al. 2014).

34

2 Physiological Effects of Bicarbonate on Plants

Fig. 2.4 A simplified model illustrating the functions of recently identified genes and mechanisms in guard cell bicarbonate-mediated stomatal movements according to previous works (Park et al. 2009; Kim et al. 2010; Xue et al. 2011; Tian et al. 2015; Engineer et al. 2016). In this model, the HT1 protein kinase, PP2Cs and ABCB14 proteins function as negative regulators (yellow), and CA1 and CA4, RCH1, OST1, GCA2, and SLAC1 function as positive mediators (green) of bicarbonatemediated stomatal closing. Abbreviations: ABA, abscisic acid; HT1, high leaf temperature kinase; GCA2, growth controlled by abscisic acid 2; CA, carbonic anhydrase; SLAC1, slow anion channel 1. RHC1, Resistance to high CO2 ; OST1, Stomatal open 1 protein kinase; PYR, Pyrabactin resistance; RCAR, Regulator component of ABA receptors; PP2Cs, protein phosphatase 2C; ABCB14, ABC malate uptake transporter (ABC Transporter B Family Member 14)

2.3.4.3

The Role of Bicarbonate in the Guard Cell Signal Transduction Network

Bicarbonate plays a central role in the guard cell signal transduction network during stomatal movement (Fig. 2.4). It can function as a small molecule activator of S-type anion channels, promote S-type anion currents, boost the intracellular Ca2+ sensitivity of S-type anion channel activation, and lead to ionic efflux, thus triggering stomatal closure (Xue et al. 2011; Kollist et al. 2014).

2.3.4.4

The Role of OST1 in the Guard Cell Signal Transduction Network

Stomatal open 1 protein kinase (OST1) is located in a central position in the guard cell signal transduction network. It is a central integrator of abscisic acid (ABA) and CO2 signal transduction in the regulation of SLAC1 (slow anion channel 1) in guard cells (Tian et al. 2015). The convergence point of CO2 and ABA signaling may be OST1, the ABA signal connects to OST1 through ABA receptors and PP2Cs (protein

2.3 The Effect of Bicarbonate on Photosynthetic Carbon Assimilation …

35

phosphatase 2C), and the CO2 signal connects to OST1 through carbonic anhydrase and RCH1 (resistant to high CO2 ) (Engineer et al. 2016). CO2 and ABA signaling synergistically mediate stomatal closing by converging pathways (Engineer et al. 2016).

2.3.4.5

The Role of HT1 in Bicarbonate-Mediated Stomatal Closing

HT1 (high leaf temperature kinase) is a major negative regulator of bicarbonatemediated stomatal closure. It phosphorylates and inactivates OST1 and prevents OST1-induced activation of SLAC1. However, RCH1 linked to high bicarbonate removes the inhibitory effect of HT1 on SLAC1 activation by OST1 (Tian et al. 2015). PP2Cs are major negative regulators of ABA-mediated stomatal closure. Perception of ABA signaling by the PYR (Pyrabactin resistance)/RCAR (Regulator component of ABA receptors) proteins shuts down negative regulation of ABA signaling by PP2Cs (Kim et al. 2010).

2.3.4.6

The Role of PEPC in the Regulation of Stomatal Movement

Phosphoenolpyruvate (PEP) carboxylase plays an important role in the regulation of stomatal movement (Thorpe 1983; Asai et al. 2000; Lawson et al. 2014). The function of bicarbonate-stimulated movement suggested the involvement of PEP carboxylase using bicarbonate as a substrate (Raschke 1975). Malate, which is synthesized by the reduction of oxaloacetate formed from PEP carboxylation, may be a major regulator of bicarbonate-mediated stomatal movement; meanwhile, PEP carboxylation is a significant source of malate in guard cells (Das and Raghavendra 1974; Pearson and Milthorpe 1974; Thorpe 1983). Extracellular malate (in the guard cell wall) from both guard cells and mesophyll cells eventually enhances anion channel activity and ABA- and CO2 -induced stomatal closure (Hedrich and Marten 1993; Hedrich et al. 1994), and malate reuptake into guard cells by a plasma membrane ABC malate uptake transporter, ABCB14 (ABC Transporter B Family Member 14), negatively regulates stomatal closure (Kim et al. 2010). Meanwhile, PEP carboxylation activates H+ -ATPases in the plasma membrane of guard cells for stomatal opening (Assmann et al. 1985; Shimazaki et al. 1986). Malate synthesis, mediated by PEP carboxylase, and reuptake in guard cells control stomatal movement. When malate reuptake is dominant, stomata open; otherwise, they close.

2.3.4.7

The Impact of Bicarbonate on Stomatal Aperture

Bicarbonate has a complex impact on stomatal aperture through carbonic anhydrase and phosphoenolpyruvate carboxylase, even as an inorganic carbon source. Table 2.14 shows the differential response of stomatal conductance in two Moraceae plant species under different bicarbonate levels in the presence and absence of Zn.

36

2 Physiological Effects of Bicarbonate on Plants

From Table 2.14, on Day 10, in the presence of Zn, BC-10 had the greatest stomatal conductance, and BC-5 had the smallest stomatal conductance; however, in the absence of Zn, a high bicarbonate level decreased the stomatal conductance among all treatments. It can be inferred that in the presence of Zn, excessive bicarbonate seriously inhibits carbonic anhydrase leading to stomatal opening and displaying the maximum stomatal conductance; however, in the absence of Zn, the activity of carbonic anhydrase in plants also decreased significantly (Rengel 1995; Pandey et al. 2002), which led to an increase in stomatal opening, but the stomatal aperture gradually decreased with increasing bicarbonate. Finally, stomatal conductance may decrease with increasing bicarbonate. Whether in the presence or absence of Zn, on Day 20 and Day 30, low bicarbonate-induced stomatal opening and high bicarbonate-mediated stomatal closure were also found in all treatments except on Day 30 for Bp in the absence of Zn. Bicarbonate positively regulates stomatal closure through SLAC1, and PEP carboxylation positively regulates stomatal opening by activating H+ -ATPases in the plasma membrane and absorbing malate by ABCB14 in guard cells. Stomatal conductance is dependent on the counterbalance of bicarbonate regulating opening and closing (Lawson 2009). The interaction of PEP carbonylation, activation of H+ -ATPases, malate synthesis and reuptake can be evaluated by measuring stomatal conductance under different bicarbonate levels. Table 2.14 Stomatal conductance (gs , mol(H2 O)m–2 s –1 ) in two plant species under different bicarbonate levels in the presence and absence of Zn. The data were determined on days 10 (Day 10), 20 (Day 20), and 30 (Day 30). The values in brackets represent the percentage compared to the control (BC-0) (the experimental conditions and process are the same as stated in Sect. 2.3.2.1) Plant species

Treatment

Bp

+Zn

Ma

Bp

Ma

+Zn

−Zn

−Zn

BC-0

Day 10

Day 20

Day 30

0.152(100)

0.118(100)

0.155(100)

BC-1

0.131(86)

0.106(90)

0.199(128)

BC-5

0.112(74)

0.185(157)

0.194(125)

BC-10

0.182(120)

0.100(85)

0.076(49)

BC-0

0.098(100)

0.112(100)

0.103(100)

BC-1

0.128(131)

0.151(135)

0.126(122)

BC-5

0.087(89)

0.102(91)

0.115(112)

BC-10

0.166(169)

0.097(87)

0.075(73)

BC-0

0.119(100)

0.103(100)

0.126(100)

BC-1

0.134(113)

0.121(117)

0.072(57)

BC-5

0.097(82)

0.122(118)

0.080(63)

BC-10

0.072(61)

0.102(99)

0.090(71)

BC-0

0.172(100)

0.146(100)

0.090(100)

BC-1

0.155(99)

0.114(78)

0.112(124)

BC-5

0.076(44)

0.155(106)

0.127(141)

BC-10

0.086(50)

0.053(36)

0.046(51)

2.3 The Effect of Bicarbonate on Photosynthetic Carbon Assimilation …

2.3.4.8

37

Bicarbonate Versus SLC1

Figure 2.4 shows that OST1 and SLAC1 are two important nodes in the network regulating anion homeostasis in guard cells and stochastic movement. SLC1 is involved not only in the metabolism and utilization of inorganic carbon but also in the metabolism of inorganic nitrogen and reactive oxygen species production. Inorganic carbon fixation of mesophyll cells and guard cells decreases the concentration of intercellular inorganic carbon to inactivate SLC1, while photophosphorylation also inactivates SLC1 by activating H+ -ATPases to stimulate stomatal opening (Lawson 2009). Bicarbonate is closely related to nitrogen metabolism by regulating SLC1. First, SLC1 is a nitrate-selective anion channel that affects nitrate absorption and metabolism (Sun et al. 2015). Second, bicarbonate can stimulate an increase in nitric oxide (Kolla and Raghavendra 2007), a signaling molecule involved in the regulation of stomatal aperture, which regulates K+ and Cl− channels in guard cells (GarciaMata et al. 2003; Zhang et al. 2016). Nitric oxide can activate S-type anion channels as secondary messengers (Vahisalu et al. 2010). There are at least two enzymes that can mediate nitric oxide synthesis: nitric oxide synthase (NOS), which catalyzes the synthesis of nitric oxide from L-arginine (Kolla and Raghavendra 2007), and nitrate reductase (NR) (Desikan et al. 2002; Kaiser et al. 2002; Meyer et al. 2005; Neill et al. 2008). Therefore, bicarbonate promotes inorganic nitrogen assimilation in plants not only by providing electrons, protons and reducing power for nitrate reduction in mesophyll cells (see Sect. 2.2, Lu et al. 2018) but also by changing nitrate uptake and utilization through SLC1 in guard cells.

2.3.4.9

The Role of ROS in the Regulation of Stomatal Closure

Reactive oxygen species (ROS) are also involved in the regulation of stomatal closure in plants (Mori et al. 2001; Hetherington and Woodward 2003; Song et al. 2014). Arabidopsis OST1 protein kinase could act upstream of reactive oxygen species production (Mustilli et al. 2002), which mediates the activation of S-type anion channels (Vahisalu et al. 2010). The production of superoxide or hydrogen peroxide is catalyzed by NADPH oxidase, and OST1 acts upstream of NADPH oxidase, which triggers ABA to activate Ca2+ channels in the plasma membrane, whereas Ca2+ mediated stomatal closure acts on osmotic regulation of guard cells through SLAC1 as a positive mediator (Murata et al., 2001; Ma et al. 2009; Park et al. 2009; Shi et al. 2015). Bicarbonate can also stimulate an increase in hydrogen peroxide (Kolla et al 2007), which regulates K+ and Cl− channels in guard cells (Zhang et al. 2016). Similarly, hydrogen peroxide is a crucial signaling molecule between OST1 and SLAC1 in bicarbonate-mediated stomatal closure (Desikan et al. 2004; Shi et al. 2015). It can be seen from the above statement that the reactive oxygen species and stomatal aperture decreases with the increase in bicarbonate; the more reactive oxygen species are produced, the greater the damage to the photosynthetic organ in plant leaves. Obviously, bicarbonate-mediated stomatal closure protects the plant

38

2 Physiological Effects of Bicarbonate on Plants

(photosynthetic organs) from water loss at the expense of the decreasing in carbon assimilation.

2.3.5 The Role of Bicarbonate in Improving Glucose Metabolism and Stress Tolerance In fact, a specific concentration of bicarbonate has a positive effect on scavenging reactive oxygen species to protect photosynthetic organs during bicarbonatemediated stomatal closure, when reactive oxygen species are produced. Plants adapt to various stresses through glucose metabolism regulation at the transcriptional and posttranslational levels. The pentose phosphate (PPP) and glycolic pathways (EMP) are the two most important and central glucose metabolic pathways in plants. PPP is an important source of reducing equivalents in the form of NADPH for biosynthetic processes such as the assimilation of inorganic nitrogen and fatty acid synthesis and modulates the redox state of cells to protect against oxidative stress. This pathway is also the source of biosynthetic precursors for the synthesis of nucleotides, aromatic amino acids, phenylpropanoids and their derivatives. EMP is a fermentative pathway of respiration that plays the primary role in providing NADH, ATP and other precursor metabolites for biomass production to meet the energy demands of metabolic activities (Yao and Wu 2016). Under stress conditions, plants alter their glucose metabolic pathway in response to environmental changes and maintain physiological function. Phosphofructokinases (PFK) and glucose-6-phosphate dehydrogenase (G6PDH) are the rate-limiting enzymes of the EMP and PPP, respectively. Many studies have indicated that glucose metabolism is switched from the EMP to PPP, and stress-induced signaling molecules such as ABA, H2 O2 , and NO are simultaneously produced by plants under stress. Salt stress caused H2 O2 production and stimulated an increase in the activities of G6PDH and antioxidative enzymes, including superoxide dismutase (SOD), peroxidase (POD), catalase (CAT), and ascorbate peroxidase (APX), in red kidney bean (Phaseolus vulgaris L.) roots under salt stress (100 mM NaCl), and exogenous H2 O2 also enhanced the activities of antioxidative enzymes as well as G6PDH (Liu et al. 2012). Although the expression of the G6PDH gene in wheat roots was stimulated by 0.15 M NaCl treatment, it did not respond to abscisic acid treatment (Nemoto and Sasakuma 2000). However, drought could trigger rapid H2 O2 and ABA accumulation and cause a marked increase in the total and cytosolic G6PDH activities in soybean roots, while exogenous H2 O2 or ABA treatment could enhance the total and cytosolic G6PDH activities (Wang et al. 2016). Meanwhile, exogenous H2 O2 could stimulate NO accumulation. NO positively regulates the transcription of genes encoding cytosolic G6PD under drought stress in soybean roots (Wang et al. 2020). Similarly, G6PDH in leaves plays a pivotal role in improving stress tolerance in plants. Zhao et al. (2015) demonstrated that G6PDH activity decreased but alternative

2.3 The Effect of Bicarbonate on Photosynthetic Carbon Assimilation …

39

pathway (AP) capacity increased, while hydrogen peroxide (H2 O2 ) accumulated when highland barley was exposed to UV-B radiation. Landi et al. (2016) found a significant increase in the total activity of G6PDH and proline synthesis and the accumulation of ascorbate peroxidase in tomato leaves exposed to short- and longterm drought stress, which could be strictly related to the ABA and PP2C signaling cascade. Yao and Wu (2016) found an activating effect on G6PDH activity and an inhibitory effect on PFK activity, an increase in the activities of peroxidase (POD), superoxide dismutase (SOD), catalase (CAT) and the proline content at first, and then a decrease in Morus alba leaves under drought induced by polyethylene glycol 6000 (PEG6000) and bicarbonate stress. Ribulose-1,5-bishphosphate carboxylase/oxygenase (Rubisco) is a key enzyme in photosynthetic inorganic carbon assimilation in plants. Ribose-1,5-diphosphate (RuBP) is the receptor of the photosynthetic carbon cycle combined with CO2 , and its production and depletion also determine the photosynthetic rate of plants. RuBP is consumed by the glycolate pathway in photorespiration, and its production and regeneration are completed through PPP (Andersson 2008). Therefore, Rubisco and PPP coordinately regulate RuBP to modulate photosynthesis in plants. Rubisco is significantly influenced by adversity. Drought or long-term water stress could significantly inhibit Rubisco activity in wheat, tobacco and subterranean clover (Demirevska et al. 2009; Medrano et al. 1997; Parry et al. 2002); however, Rubisco increased in birch (Betula pendula) under drought stress (Pääkkönen et al. 1998). Rubisco in cotton and tobacco leaves was rapidly inactivated, and photosynthesis was inhibited by heat stress (Salvucci and Crafts-Brandner 2004a,b). High light stress led to a decrease in carboxylase activity in Rubisco (Siedlecka and Krupa 2004). Low light stress could act downstream of the Rubisco gene during cucumber (Cucumis sativus L.) leaf development (Sun et al. 2014). Bicarbonate, which could induce Fe deficiency in plants, reduced the leaf Rubisco activity and RuBP carboxylation capacity to diminish photosynthetic competence (Msilini et al. 2009). Moreover, proline, which was stimulated by various stresses, could inhibit the activity of Rubisco carboxylase, even when the concentration was as low as 100 mM, and the inhibition increased with increasing proline concentration (Sivakumar et al. 1998). Therefore, the decrease in Rubisco activity under stress may be partly related to the accumulation of osmolytes such as proline and betaine. Specific concentrations of bicarbonate could provide carbon sources and activate the pentose phosphate pathway and ribulose-1,5-bishphosphate carboxylase/ oxygenase (Rubisco) to counteract the negative influence in plants under stress. Recently, we examined the effects of different bicarbonate concentrations (0, 3 mM, 6 mM, and 9 mM) with simulated mild drought stress (induced by 50 g L−1 PEG 6000) on growth, photosynthetic traits, glucose metabolism, Rubisco and radicalscavenging activity (Yao and Wu 2021). Six-week-old paper mulberry (Broussonetia papyrifera L.) seedlings were used for the following treatments: (i) addition of 50 g L−1 PEG 6000 in 1/2 Hoagland nutrient solution to simulate mild drought stress. (ii) Bicarbonate treatment was performed by adding one of four levels of bicarbonate [0 (T1 ), 3 (T2 ), 6 (T3 ), or 9 (T4 ) mM as NaHCO3 ] to simulate mild drought stress, and a control solution was also used. The solution pH was adjusted to 7.8 in all treatments

40

2 Physiological Effects of Bicarbonate on Plants

by the addition of NaOH, and solutions were replaced every day after the imposition of stress treatments. The results obtained from the above experiment show that growth and photosynthesis can be recovered by 3 mM bicarbonate (T2 treatment), and high concentrations of bicarbonate (6, 9 mM) inhibit the growth and reduce the photosynthetic assimilation ability of paper mulberry seedlings. Except for the T2 treatment, the ABA content increased, and the stomatal conductance decreased with increasing bicarbonate. This result demonstrated that the positive role of 3 mM bicarbonate can partly offset the negative effects of stress imposed on plants. Glucose metabolism differs in bicarbonate concentrations. Paper mulberry seedlings under 3 mM bicarbonate with simulated mild drought stress showed the highest total activity of glucose catabolism enzymes and G6PDH activity; similarly, high concentrations of bicarbonate (6, 9 mM) decreased the total activity of glucose catabolism enzymes and G6PDH activity. Moreover, the proportion of glucose metabolism through PPP increased with increasing concentrations of bicarbonate, and T4 treatments (9 mM bicarbonate) at 10 d were the exception. Under the control, T1, T2 T3 and T4 treatments at 10 d, the proportion of glucose metabolism through PPP was stable at approximately 32%, 56%, 57%, 63% and 44% of the total amount of glucose catabolism, respectively. This result indicated that the switch from the EMP to PPP of glucose metabolism can be superimposed on simulated mild drought and excessive bicarbonate stress (Yao and Wu 2021). Our study also showed that specific concentrations of bicarbonate could increase Rubisco activities. Although simulated mild drought stress significantly suppressed Rubisco activity in paper mulberry, Rubisco activities can be recovered by 3 mM bicarbonate (T2 treatment), and a high concentration of bicarbonate (9 mM) inhibits Rubisco activity. Excessive bicarbonate stress can strengthen the inhibition of Rubisco activity by simulated mild drought (Yao and Wu 2021). The reactive oxygen species scavenging ability of plants also differs in bicarbonate concentrations. The reactive oxygen species scavenging ability increased with increasing bicarbonate at 1 d, and there was a significant difference among treatments. Bicarbonate (3 mM) increased the reactive oxygen species scavenging ability all the time and maintained a maximum at 10 d; however, 9 mM bicarbonate decreased the reactive oxygen species scavenging ability during the later stage of treatment. Therefore, plants under 3 mM bicarbonate with simulated mild drought stress showed the best reactive oxygen species scavenging ability (Yao and Wu 2021). Obviously, adversity had negative effects on plant physiological activities and growth, and a high concentration of bicarbonate had a superposition effect on these negative effects. However, specific concentrations of bicarbonate had positive effects on photosynthesis, glucose metabolism, Rubisco activation, the switch from the EMP to PPP of glucose metabolism and reactive oxygen species scavenging ability, which improved the growth of plants grown in the environment with the concentration of bicarbonate. Table 2.1 shows that the concentration of bicarbonate, which is suitable for plant growth, has interspecific differences. Photosynthate provides metabolic substrates, osmolytes and soluble sugar for plants. A specific concentration of bicarbonate (3 mM) increased the photosynthetic capacity, including net CO2 assimilation and bicarbonate-use, which promoted

2.3 The Effect of Bicarbonate on Photosynthetic Carbon Assimilation …

41

Fig. 2.5 Schematic representation of specific concentrations of bicarbonate improving growth, photosynthesis, glucose metabolism and stress tolerance. An, net CO2 assimilation rate; BU, bicarbonate-use rate; gs , stomatal conductance; IC, inorganic carbon; PPP, pentose phosphate pathway; EMP, glycolic pathway

total sugar metabolism, including PPP and EMP. Meanwhile, a specific concentration of bicarbonate also had an activating effect on G6PDH and an inhibitory effect on PFK activity in paper mulberry leaves. Enhanced glycolysis provided more ATP, NADH and other precursor metabolites for biomass production to meet the energy demands of metabolic activities. G6PDH is the key regulatory enzyme of the PPP, which controls the flow of carbon and produces NADPH. The stimulation of PPP, on the one hand, increases the efficiency of antioxidant enzymes to strengthen reactive oxygen species scavenging ability and restrain photosynthetic system damage by membrane-lipid peroxidation; on the other hand, it enhances the production of ribulose-5-phosphate to improve the regeneration ability of RuBP and Rubisco activity in the Calvin-Benson cycle, thereby generating a virtuous cycle, which provides more ATP, NADH and other precursor metabolites for biomass production to meet the energy demands of metabolic activities (Schnarrenberger et al. 1973; Sharkey and Weise 2016). Therefore, the specific concentration of bicarbonate in the rhizosphere of plants under drought stress mainly acts as an alternative carbon source, stimulates glucose metabolism, increases the switch from the EMP to PPP of glucose metabolism, and has a positive effect on the recovery of growth and photosynthesis (Fig. 2.5). Bicarbonate by spraying on leaves can also restore the photosynthetic capacity of plants under drought conditions (Burbulis et al. 2017).

2.3.6 The Role of Bicarbonate in Inducing Chlorosis Bicarbonate could buffer soil solution, which is maintained in a weak alkali environment, markedly reduce Fe solubility and availability, inhibit the induction of root Fe (III) chelate, prevent the transport of Fe from root to shoot, and cause the alkalinisation of the leaf apoplast, which results in reduced Fe uptake by mesophyll cells, ultimately resulting in chlorosis symptomatology. Bicarbonate at high concentrations makes iron (Fe) not easily available to roots and induces iron deficiency and

42

2 Physiological Effects of Bicarbonate on Plants

chlorosis, which is called Fe deficiency-induced chlorosis or Fe-deficiency chlorosis (Zhou et al. 1984; Lucena 2000; Bertoni et al. 1992; Msilini et al. 2009). Fe-deficiency chlorosis induced by bicarbonate is a common physiological symptomatology in plants. It was found to be widespread in some plants, such as soybean (Glycine max (L.) Merr. (Dofing et al.1989), tomato (Lycopersicon esculentum Mill.)(Buckhout, et al. 1989), sunflower (Helianthus annuus L.) (Kolesch et al. 1984), Chrysanthemum morifolium (Ram.) (Rutland and Bukovac 1971), pear (Pyrus communis L.), quince (Cydonia oblonga Mill.) (Donnini et al. 2009), pea (Pisum sativum L.) (Jelali et al. 2011), barley (Hordeum vulgare L.), sorghum (Sorghum bicolor L.), and maize (Zea mays L.) (Alhendawi et al. 1997). However, some plants present adaptive responses to Fe deficiency when they suffer bicarbonate-induced Fe-deficiency chlorosis. There are two strategies for plants to adapt to Fe deficiency: Strategy I is adopted by most plant species, including dicotyledonous and nongrammaceous monocotyledonous species, and Strategy II is adopted by a few plant species from graminaceous monocots (Jolley et al. 1996). The Strategy II plants could release phytosiderophores to solubilize Fe in response to Fe-deficiency stress (Marschner et al. 1987; Jolley et al. 1996). Strategy I is the most common adaptive mechanism adopted by most plant species in response to Fe deficiency. Therefore, we solely discuss the adaptive biochemical mechanisms of the iron uptake efficiency of strategy I plants in this section. Fe3+ uptake rates were several orders of magnitude less than those for ferric chelate reduction, and plants assimilated Fe2+ , which was from the reduction of Fe3+ by root Fe (III) chelate reductase (Eckhardt and Buckhout 2000). Ferric reduction, prior to Fe2+ uptake, is thought to be an obligatory step in iron uptake as well as the primary factor in making iron available for absorption by plants (Yi and Guerinot 1996). The response to Fe deficiency of Strategy I plants includes stimulation of Fe(III)-chelate reductase and H+ -ATPase, which lower the rhizosphere pH and increase the capacity to reduce ferric to ferrous forms, make available Fe2+ in the soil solution, and regulate cation absorption (Dofing et al.1989; Yi and Guerinot 1996; Msilini et al. 2009; Donnini et al. 2009; Martinez-Cuenca et al. 2013); an increase in phosphoenolpyruvate carboxylase (PEPC) activities of roots, which is receptor of bicarbonate, regulate pH and sustain ATP and NAD(P)H demands by enhancing catabolic carbohydrate flux ((De Nisi and Zocchi 2000); and a decrease in Rubisco of leaves (Msilini et al. 2009). In addition, Fe deficiency stimulates an accumulation of organic acids in roots, mainly citrate and malate, which is mainly involved in Fe transport to shoots as Fe citrate in xylem sap (Abadía et al. 2002; Stephan 2002). However, the biochemical response to bicarbonate-induced iron deficiency was dependent on plant species and genotypes. Gao and Shi (2007) demonstrated that the Fe reduction capacity and quality of released hydrogen ions from roots increased only in Fe-resistant peanut cultivars under Fe deficiency stress. Donnini et al. (2009) found significant differences between Pyrus communis L. cv. Conference and Cydonia oblonga Mill. BA29 and MA clones in response to chlorosis. Bicarbonate-induced Fe deficiency increased ferric chelate reductase (FC-R) activity in cv. Conference and BA29, while FC-R activity was not significant for MA, and PEPC activity was enhanced only in pear rather than quince. Jelali et al. (2011) demonstrated

2.4 Effect of Bicarbonate on Inorganic Nitrogen Metabolism in Plants

43

a higher accumulation of potassium in two tolerant pea cultivars, Kelvedon and Douce, whereas no significant accumulation was recorded in the content of this nutrient in leaves of the susceptible cultivar, Lincoln. Mohamed et al. (2013) found, by comparing the biochemical responses of two Lactuca sativa varieties (Romaine and Vista) to bicarbonate-induced iron deficiency, that PEPC activity was enhanced only in the Vista variety. Gama et al. (2020) assessed the effect of bicarbonate on root FC-R activity and the genetic expression of the calmodulin-regulated Ca2+ -ATPase pump (ACA gene) of carob-tree seedlings. They found that root FC-R activity and ACA gene expression were not enhanced under Fe deficiency induced by bicarbonate. Iron deficiency symptoms occur prior to leaf chlorosis, and bicarbonate-induced Fe-deficiency chlorosis can be divided into bicarbonate-induced Fe-deficiency and Fe deficiency-induced chlorosis stages (Gruber and Kosegarten 2002). Therefore, the emergence time and degree of chlorosis are different owing to the differential response to Fe deficiency induced by bicarbonate among plant species. Two Moraceae plant species, B. papyrifera and M. alba, were more resistant to Fe deficiency induced by bicarbonate than two Cruciferae plant species, O. violaceus and B. napus. Bicarbonate stimulated or restrained chlorophyll synthesis to some extent in the two Moraceae plant species during the early or later stages of treatment, respectively (see Table 2.9). Ferric reduction was also affected by biological and abiotic factors. Older leaves reduced Fe(III)-chelate at a lower rate than did young leaves (de la Guardia and Alcantara 1996). Ferric reduction was stimulated by light but inhibited by O2 (de la Guardia and Alcantara 1996). Hypoxia suppresses the expression of several Fe acquisition genes in Fe-deficient Arabidopsis, cucumber and pea plants (García et al. 2014). Moreover, iron deficiency decreased catalase but increased peroxidase and superoxide dismutase in response to the decrease in hydroxyl radicals and the increase in superoxide anions in leaves of young Brassica napus plants (Tewari et al. 2013), while antioxidant enzymes such as superoxide dismutase decreased ferric chelate reduction (de la Guardia and Alcantara 1996; Valipour et al. 2020). Oxidative stress and antioxidant responses to bicarbonate-induced iron deficiency were also dependent on plant species and genotypes. In the leaves of sensitive quince seedlings, bicarbonate-induced iron deficiency decreased ascorbate peroxidase, catalase, and guaiacol peroxidase activities, which were not reduced in tolerant hawthorn rootstock (Valipour et al. 2020).

2.4 Effect of Bicarbonate on Inorganic Nitrogen Metabolism in Plants Bicarbonate also indirectly affects the inorganic carbon assimilation of plants by coordinating the contribution of ammonium nitrogen and nitrate nitrogen. The effect of bicarbonate on the inorganic nitrogen metabolism of plants has speciesspecific characteristics. Xia and Wu (2022a) studied the conjugation effects of

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bicarbonate and ammonium/nitrate nitrogen on inorganic nitrogen metabolism in Orychophragmus violaceus and Brassica napus. The results showed that bicarbonate and ammonium nitrogen jointly promoted ammonium nitrogen use in B. napus rather than in O. violaceus. Bicarbonate and ammonium nitrogen conjugates decreased the nitrate contribution in O. violaceus and the accumulation of total inorganic nitrogen in B. napus (Xia and Wu 2022a). Xia and Wu (2022a, b) also studied the responses of inorganic nitrogen assimilation of O. violaceus and B. napus to different bicarbonate levels (1, 5 and 15 mM NaHCO3 added to the culture solution). The results showed that the increased bicarbonate concentration improved the activity of glutamate synthase and nitrate reductase in O. violaceus more than in B. napus. The highest activities of glutamate synthase and nitrate reductase were assessed in O. violaceus at the middle level of bicarbonate, whereas a decline in the activities of these enzymes was observed in B. napus. Moreover, at the heaviest bicarbonate level, the decline in the activities of glutamate synthase and nitrate reductase was less in O. violaceus than in B. napus (Xia and Wu 2022b). At the middle bicarbonate level, bicarbonate significantly improved the nitrate contribution, resulting in an increase in nitrogen utilization in O. violaceus compared to B. napus. However, at the heaviest level of bicarbonate, the nitrate/ammonium contribution and total nitrogen assimilation were suppressed in B. napus. However, in O. violaceus, nitrate utilization was reduced, and ammonium utilization was increased (Xia and Wu 2022b).

2.5 Indirect Effects of Bicarbonate on the Growth and Development of Plants Bicarbonate can indirectly influence the growth and development of plants by changing the osmotic potential to induce salt stress and increasing the pH of the soil solution to produce alkali stress. The stresses induced by neutral salts such as NaCl and Na2 SO4 are different from those induced by NaHCO3 . These stresses are actually two distinct kinds of stresses: the former is called salt stress, and the latter is called alkali stress (Shi and Sheng 2005). Salt stress increases osmotic potential and disrupts ion homeostasis in plant cells. However, although alkali stress exerts the same negative effects, its adverse impact is further intensified when it is combined with a high pH value (Guo et al. 2017). Bie et al. (2004) investigated the influences of salinity on growth, gas exchange and distribution of mineral composition in plants. They demonstrated that leaf growth, shoot dry weight, photosynthetic rate and stomatal conductance decreased with increasing concentrations of Na2 SO4 or NaHCO3 . Both NaHCO3 and Na2 SO4 stress can cause chlorosis of lettuce leaves, which appeared at NaHCO3 concentrations above 5 mM and at Na2 SO4 concentrations above 40 mM. Shi and Sheng (2005)

2.5 Indirect Effects of Bicarbonate on the Growth and Development of Plants

45

observed the effect of alkalinity and salinity on the growth and physiological parameters of sunflower seedlings and demonstrated that the relative growth rate, leaf area, and K+ content decreased with increasing salinity and pH. The electrolyte leakage rate, K+ content, citric acid content, and proline content increased with increasing salinity and pH. This result indicated that the deleterious effects of excessive NaHCO3 were significantly greater than those of a high pH value or salinity alone. Yang et al. (2009) compared the effects of salt stress and alkali stress on the growth, photosynthesis, solute accumulation, and ion balance of barley plants. They demonstrated that the relative growth rate, net photosynthetic rate, stomatal conductance, and water content of barley decreased with increased salinity, and the reduction under salt stresses (1:1 molar ratio of NaCl to Na2 SO4 ) was lower than that under alkali stresses (1:1 molar ratio of NaHCO3 to Na2 CO3 ). Barley accumulated more inorganic ions other than organic acids to balance the massive influx of cations under salt stresses. However, alkali stresses might enhance organic acid synthesis to maintain intracellular ion balance and stable pH. Zhang and Mu (2009) also observed the response of alkali stress (9:1 molar ratio of NaHCO3 : Na2 CO3 , pH 8.71–8.89) and salt stress (9:1 molar ratio of NaCl: Na2 SO4 , pH 6.44–6.65) on the germination, growth, photosynthesis, ionic balance and activity of antioxidant enzymes of Lathyrus quinquenervius. Alkali stress had greater inhibitory effects on germination, growth, photosynthesis and root system activity than salt stress. Meanwhile, Lathyrus quinquenervius may enhance organic acid synthesis, H2 O2 and malondialdehyde content, resulting in severe intracellular oxidative stress under alkali stress. Moreover, salt stress and low alkali stress slightly increased the activities of ascorbate peroxidase and superoxide dismutase but did not affect catalase activity. However, strong alkali stress significantly increased the activities of ascorbate peroxidase and superoxide dismutase and decreased catalase activity. It can be seen from the above that the effects of alkali stress on germination, growth, photosynthesis, ionic balance and activity of antioxidant enzymes are greater than those of salt stress. In fact, both salt stress and alkali stress can influence gene expression, saccharide metabolism and protein synthesis in plants (García et al. 2014; Debouba et al. 2013; Yin et al. 2017). Moreover, the effects of alkali stress on gene expression and other metabolic processes surpass those of salt stress (Zhu et al. 2012; Chen et al. 2015). Therefore, it can be speculated that plants need more positive effects of bicarbonate and consume more energy to offset the massive deleterious effects of alkali stresses.

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2.6 The Whole Effect of Bicarbonate on Plant Growth and Development In fact, the physiological and biochemical effects of bicarbonate on plants are entirely synergistic rather than from a single profile. Comprehensive and synergistic responses to bicarbonate in plants are involved in the conversion of inorganic carbon to organic carbon, the accumulation of organic acids, and the harvest, transfer and flow of chemical energy, as illustrated in Fig. 2.6. First, to date, bicarbonate channels or transporters have been less characterized in higher terrestrial plants, but the possibility of membrane transport by HCO− 3 transporters or anion channels cannot be excluded (Price et al. 2004; Poschenrieder et al. 2018). However, most bicarbonate enters the cytosol mediated by fast transformation between bicarbonate and CO2 by periplasmic CA and cytoplasmic membrane CA, as well as transmembrane diffusion of CO2 . Dissolved inorganic carbon in the cytosol in roots was transferred to the shoots and leaves via apoplast and symport transport. Bicarbonate in roots mediated the expression of several Fe acquisition genes as a signal molecule. Bicarbonate in leaves, on the one hand, provides protons, electrons, and carbon dioxide for photosynthesis as a direct substrate of photosynthetic oxygen evolution; on the other hand, it mediates stomatal movements as a signal molecule. Second, bicarbonate use by plant roots is carried out via the carboxylation of phosphoenolpyruvate catalysed by phosphoenolpyruvate carboxylase in the cytosol (E.C. 4.1.1.31), which generate oxaloacetate and are partially rapidly transferred to the shoots and leaves; the rest remain in the roots (Vapaavuori and Pelkonen 1985; Bialczyk and Lechowski 1992). Subsequently, oxaloacetate is reduced to malate by malate dehydrogenase, and malate is converted into citrate by citrate synthase (Andaluz et al. 2002). Malate and citrate in roots, on the one hand, may lower the rhizosphere pH and regulate Fe uptake; on the other hand, they may be incorporated into the tricarboxylic acid cycle and nonstructural carbohydrate pools (Rao et al. 2019). The decarboxylation of organic acids, the reduction of oxaloacetate, and the malate-oxaloacetate shuttle between the cytosol and mitochondria in leaves may provide carbon sources for refixing inorganic carbon and enhancing catabolic carbohydrate flux, thereby stimulating glucose metabolism and increasing the switch from the EMP to PPP of glucose metabolism. The stimulation of PPP not only strengthens the reactive oxygen species scavenging ability but also improves the regeneration ability of RuBP. Meanwhile, malate in leaves may enhance anion channel activity and activate H+ -ATPases, eventually controlling stomatal movement. Finally, the harvest, transfer and flow of chemical energy in plants in response to bicarbonate also form a unity. Although the increased production of organic acids to reduce pH and the increased synthesis of plant hormones, DNA, RNA, proteins and other substances to cope with bicarbonate stress all need energy, these energies can be supplemented by bicarbonate harvesting light energy from photosynthesis. Different organs, tissues, cells and organelles of plants respond to bicarbonate differently and have different energy requirements. Energy among organs, tissues, cells and organelles can be transferred through the malate-oxaloacetate shuttle (Giersch 1982;

2.6 The Whole Effect of Bicarbonate on Plant Growth and Development

47

Fig. 2.6 Hypothetical schematic model of the comprehensive and synergistic responses to bicarbonate in plants. EMP, glycolic pathway; IC, inorganic carbon; PEPC, phosphoenolpyruvate carboxylase; pCA, periplasmic CA; pmCA, cytoplasmic membrane CA; PPP, pentose phosphate pathway; ROS, reactive oxygen species; RuBP, ribose-1,5-diphosphate; TCA cycle, tricarboxylic acid cycle

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Zoglowek et al. 1988; Heineke et al. 1991; Gardeström and Igamberdiev 2016). Eventually, energy can flow smoothly from chloroplasts to various organelles, tissues and organs. Therefore, assimilating bicarbonate from photosynthesis is the determinant of plant adaptation to bicarbonate stress.

2.7 Conclusion Bicarbonate and carbon dioxide are two interchangeable inorganic carbon forms in nature. On the one hand, bicarbonate controls photosynthetic oxygen evolution, coordinates photo-reaction and dark reaction, and can also be used as an alternative inorganic carbon source when the supply of carbon dioxide is limited. On the other hand, bicarbonate can also regulate stomatal movement to coordinate the supply of water and inorganic carbon. Meanwhile, bicarbonate can also regulate glucose metabolism, promote the conversion of plants from glycolysis to the pentose phosphate pathway, and increase the stress resistance of plants. The adaptability of plants to bicarbonate is species specific. Finally, we can conclude that bicarbonate has no less physiological effect on plants than carbon dioxide. The roles of bicarbonate and carbon dioxide in plants are difficult to distinguish. In the future, we can discuss the differential effects of carbon dioxide and bicarbonate from more physiological and biochemical aspects and make full use of the positive effects of bicarbonate to serve the goal of carbon neutralization.

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Murata Y, Pei ZM, Mori IC, Schroeder J (2001) Abscisic acid activation of plasma membrane Ca2+ channels in guard cells requires cytosolic NAD(P)H and is differentially disrupted upstream and downstream of reactive oxygen species production in abi1-1 and abi2-1 protein phosphatase 2C mutants. Plant Cell 13:2513–2523 Neill S, Barros R, Bright J, Desikan R, Hancock J, Harrison J, Morris P, Ribeiro D, Wilson I (2008) Nitric oxide, stomatal closure, and abiotic stress. J Exp Bot 59:165–176 Nemoto Y, Sasakuma T (2000) Specific expression of glucose-6-phosphate dehydrogenase (G6PDH) gene by salt stress in wheat (Triticum aestivum L.). Plant Sci 158(1–2):53–60 Pandey N, Pathak GC, Singh AK, Sharma CP (2002) Enzymic changes in response to zinc nutrition. J Plant Physiol 159(10):1151–1153 Parry MAJ, Andralojc PJ, Khan S, Lea PJ, Keys AJ (2002) Rubisco activity: effects of drought stress. Ann Bot-London 89(7):833–839 Pääkkönen E, Vahala J, Pohjola M, Holopainen T, Kärenlampi L (1998) Physiological, stomatal and ultrastructural ozone responses in birch (Betula pendula Roth.) are modified by water stress. Plant Cell Environ 21(7): 671–684 Pearson CJ, Milthorpe FL (1974) Structure, carbon dioxide fixation and metabolism of stomata. Aus J Plant Physiol 1:221–236 Radmer R, Ollinger O (1980) Isotopic composition of photosynthetic O2 flash yields in the presence of H2 18 O and HC18 O3 − . FEBS Lett 110(1):57–61 Rao S, Wu Y, Wang R (2019) Bicarbonate stimulates nonstructural carbohydrate pools of Camptotheca acuminata. Physiol Plant 165(4):780–789 Raschke K (1975) Stomatal action. Ann Rev. Plant Physiol 26:309–340 Rengel Z (1995) Carbonic anhydrase activity in leaves of wheat genotypes differing in Zn efficiency. J Plant Physiol 147(2):251–256 Ruben S, Randall M, Kamen M, Hyde JL (1941) Heavy oxygen (O18 ) as a tracer in the study of photosynthesis. J Am Chem Soc 63(3):877–879 Rutland RB, Bukovac MJ (1971) The effect of calcium bicarbonate on iron absorption and distribution by Chrysanthemum morifolium (Ram.). Plant Soil 35(1–3): 225–236 Salvucci ME, Crafts-Brandner SJ (2004a) Mechanism for deactivation of Rubisco under moderate heat stress. Physiol Plant 122(4):513–519 Salvucci ME, Crafts-Brandner SJ (2004b) Inhibition of photosynthesis by heat stress: the activation state of Rubisco as a limiting factor in photosynthesis. Physiol Plant 120(2):179–186 Schnarrenberger C, Oeser A, Tolbert NE (1973) Two isoenzymes each of glucose-6-phosphate dehydrogenase and 6-phosphogluconate dehydrogenase in spinach leaves. Arch Biochem Biophys 154(1):438–448 Sharkey TD, Weise SE (2016) The glucose 6-phosphate shunt around the Calvin-Benson cycle. J Exp Bot 67(14):4067–4077 Sharma PN, Kumar N, Bisht SS (1996) Guard cell carbonic anhydrase activity and stomatal opening in zinc deficient faba bean, vicia faba L. Indian J Exp Biol 34(6):560–564 Shevela D, Do HN, Fantuzzi A, Rutherford AW, Messinger J (2020) Bicarbonate-mediated CO2 formation on both sides of Photosystem II. Biochemistry 59(26):2442–2449 Shitov AV (2022) An Insight into the bicarbonate effect in photosystem II through the prism of the JIP Test. Photochem 2:779–797 Shi D, Sheng Y (2005) Effect of various salt–alkaline mixed stress conditions on sunflower seedlings and analysis of their stress factors. Environ Exp Bot 54(1):8–21 Shi K, Li X, Zhang H, Zhang G, Liu Y, Zhou Y, Xia X, Chen Z, Yu J (2015) Guard cell hydrogen peroxide and nitric oxide mediate elevated CO2 -induced stomatal movement in tomato. New Phytol 208(2):342–353 Shimazaki K, Iino M, Zeiger E (1986) Blue light-dependent proton extrusion by guard-cell protoplasts of Vicia faba. Nature 319:324–326 Siedlecka A, Krupa Z (2004) Rubisco activity maintenance in environmental stress conditions-how many strategies. Cellular Mol Biol Lett 9:56–57

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Sivakumar P, Sharmila P, Saradhi PP (1998) Proline suppresses Rubisco activity in higher plants. Biochem Biophys Res Commun 252(2):428–432 Stemler A, Radmer R (1975) Source of photosynthetic oxygen in bicarbonate-stimulated Hill reaction. Science 190(4213):457–458 Stimler K, Berry JA, Yakir D (2012) Effects of carbonyl sulfide and carbonic anhydrase on stomatal conductance. Plant Physiol 158(1):524–530 Stephan UW (2002) Intra- and intercellular iron traffificking and subcellular compartmentation within roots. Plant Soil 241:19–25 Song YW, Miao YC, Song CP (2014) Behind the scenes: the roles of reactive oxygen species in guard cells. New Phytol 201:1121–1140 Sun JL, Sui XL, Huang HY, Wang SH, Wei YX, Zhang ZX (2014) Low light stress down-regulated Rubisco gene expression and photosynthetic capacity during Cucumber (Cucumis sativus L.) leaf development. J Integr Agr 13(5): 997–1007 Sun SJ, Qi GN, Gao QF, Wang HQ, Yao FY, Hussain J, Wang YF (2015) Protein kinase OsSAPK8 functions as an essential activator of S-type anion channel OsSLAC1, which is nitrate-selective in rice. Planta 243(2):489–500 Tewari RK, Hadacek F, Sassmann S, Lang I (2013) Iron deprivation-induced reactive oxygen species generation leads to non-autolytic PCD in Brassica napus leaves. Environ Exp Bot 91:74–83 Thorpe N (1983) The role of phosphoenolpyruvate carboxylase in the guard cell of Commelina cyanea. Plant Sci Lett 30(3):331–338 Tian W, Hou C, Ren Z, Pan JJ, Zhang H, Bai F, Zhang P, Zhu H, He Y, Luo S, Li L, Luan S (2015) A molecular pathway for CO2 response in Arabidopsis guard cells. Nature Commun 6:6057 Vahisalu T, Puzõrjova I, Brosché M, Valk E, Lepiku M, Moldau H, Pechter P, WangYS LO, Salojärvi J, Loog M, Kangasjärvi J, Kollist H (2010) Ozone-triggered rapid stomatal response involves the production of reactive oxygen species, and is controlled by SLAC1 and OST1. Plant J 62(3):442–453 Valipour M, Baninasab B, Khoshgoftarmanesh AH, Gholami M (2020) Oxidative stress and antioxidant responses to direct and bicarbonate-induced iron deficiency in two quince rootstocks. Sci Hortic-Amsterdam 261:108933 Vapaavuori EM, Pelkonen P (1985) HCO3 - uptake through the roots and its effect on the productivity of willow cuttings. Plant Cell Environ 8:531–544 Warburg O, Krippahl G (1958) Hill-reaktionen. Z Naturforsch B 13(8):509–514 Wang H, Yang L, Li Y, Hou J, Huang J, Liang W (2016) Involvement of ABA-and H2 O2 -dependent cytosolic glucose-6-phosphate dehydrogenase in maintaining redox homeostasis in soybean roots under drought stress. Plant Physiol Biochem 107:126–136 Wang R, Wu Y, Xing D, Hang H, Xie X, Yang X, Zhang K, Rao S (2017) Biomass production of three biofuel energy plants’ use of a new carbon resource by carbonic anhydrase in sSimulated karst soils: Mechanism and capacity. Energies 10(9):1370–1384 Wang X, Ruan M, Wan Q, He W, Yang L, Liu X, He L, Yan L, Bi Y (2020) Nitric oxide and hydrogen peroxide increase glucose-6-phosphate dehydrogenase activities and expression upon drought stress in soybean roots. Plant Cell Rep 39(1):63–73 Wu Y (2021a) Is bicarbonate directly used as substrate to participate in photosynthetic oxygen evolution. Acta Geochim 40(4):650–658 Wu Y (2021b) Bicarbonate use and carbon dioxide concentrating mechanisms in photosynthetic organisms. Acta Geochim 40(5):846–853 Wu Y (2023) Combined effect of bicarbonate and water in photosynthetic oxygen evolution and carbon neutralits. Acta Geochim 42(1):77–88 https://doi.org/10.1007/s11631-022-00580-9 Wu YY, Xing DK (2012) Effect of bicarbonate treatment on photosynthetic assimilation of inorganic carbon in two plant species of Moraceae. Photosynthetica 50(4):587–594 Wu YY, Xing DK, Hang HT, Zhao K (2018) The adaptive mechanism of the karst-adaptable plants. Principles and technology of determination on plants’ adaptation to karst environment. Science Press, Beijing, pp 1–88

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Xia A, Wu Y (2022a) Joint interactions of carbon and nitrogen metabolism dominated by bicarbonate and nitrogen in Orychophragmus violaceus and Brassica napus under simulated karst habitats. BMC Plant Biol 22:264 Xia A, Wu Y (2022b) Differential responses of nitrate/ammonium use to bicarbonate supply in two Brassicaceae species under simulated karst habitat. Agronomy 12:2080 Xue S, Hu H, Ries A, Merilo E, Kollist H, Schroeder JI (2011) Central functions of bicarbonate in S-type anion channel activation and OST1 protein kinase in CO2 signal transduction in guard cell. Embo J 30(8):1645–1658 Yang CW, Xu HH, Wang LL, Liu J, Shi DC, Wang DL (2009) Comparative effects of salt-stress and alkali-stress on the growth, photosynthesis, solute accumulation, and ion balance of barley plants. Photosynthetica 47(1):79–86 Yao K, Wu YY (2016) Phosphofructokinase and glucose-6-phosphate dehydrogenase in response to drought and bicarbonate stress at transcriptional and functional levels in mulberry. Russ J Plant Physiol 63(2):235–242 Yao K, Wu YY (2021) Rhizospheric bicarbonate improves glucose metabolism and stress tolerance of Broussonetia papyrifera L. seedlings under simulated drought stress. Russ J Plant Physiol 68(1):126–135 Yi Y, Guerinot ML (1996) Genetic evidence that induction of root Fe (III) chelate reductase activity is necessary for iron uptake under iron deficiency. Plant J 10(5):835–844 Yin Z, Balmant K, Geng S, Zhu N, Zhang T, Dufresne C, Dai S, Chen S (2017) Bicarbonate induced redox proteome changes in Arabidopsis suspension cells. Front Plant Sci 8:58 Zhang A, Ren HM, Tan YQ, Qi GN, Yao FY, Wu GL, Yang LW, Hussain J, Sun SJ, Wang YF (2016) S-type anion channels SLAC1 and SLAH3 function as essential negative regulators of inward K+ channels and stomatal opening in Arabidopsis. Plant Cell 28(4):949–965 Zhang JT, Mu CS (2009) Effects of saline and alkaline stresses on the germination, growth, photosynthesis, ionic balance and anti-oxidant system in an alkali-tolerant leguminous forage Lathyrus quinquenervius. Soil Sci Plant Nutr 55(5):685–697 Zhao C, Wang X, Wang X, Wu K, Li P, Chang N, Wang J, Wang F, Li J, Bi Y (2015) Glucose-6phosphate dehydrogenase and alternative oxidase are involved in the cross tolerance of highland barley to salt stress and UV-B radiation. J Plant Physiol 181:83–95 Zhou HJ, Korcak RF, Fan F, Faust M (1984) The effect of bicarbonate induced Fe chlorosis on mineral content and Ca45 uptake of apple seedlings. J Plant Nutr 7(9):1355–1364 Zhu D, Cai H, Luo X, Bai X, Deyholos MK, Chen Q, Chen C, Ji W, Zhu Y (2012) Over-expression of a novel JAZ family gene from Glycine soja, increases salt and alkali stress tolerance. Biochem Bioph Res Co 426:273–279 Zoglowek C, Krömer S, Heldt HW (1988) Oxaloacetate and malate transport by plant mitochondria. Plant Physiol 87(1):109–115

Chapter 3

The Diversity, Plasticity and Roles of Carbonic Anhydrase in Inorganic Carbon Utilization in Plants

Abstract Carbonic anhydrase (CA, EC4.2.1.1) is ubiquitous in almost all types of organisms in the soil and water medium and on the surface of rock. Carbonic anhydrase has many isoenzymes, which are divided into at least nine genetic families and designated α-, β-, γ-, δ-, ζ-, ε-, η-, θ-, and ι-CA. Different isoenzymes of CA have different amino acid sequences and different metal cofactors. The structures of CAs and the metal coordination patterns of their active centers are diverse. Inhibition constants of inorganic anions, inhibitors and organic acids against different classes of CA isozymes and the activation profile of CAs with amino acids and amines were documented. The number of genes encoding CA in different plant species is listed. This chapter summarizes that the differential expression of genes encoding different CA isoenzymes resulted in the functional diversity of CA. Wide differences in the activation or inhibition effects of organic and inorganic substances on different carbonic anhydrase isoenzymes resulted in each well-organized carbon metabolism pathway under normal physiological conditions. Unique thylakoid carbonic anhydrase is closely related to photosynthetic oxygen evolution. PS II may be a multienzyme complex with oxygen evolution and CA functions. Carbonic anhydrase not only directly controls photosynthetic oxygen release and CO2 and bicarbonate utilization by plants but also indirectly affects the inorganic carbon assimilation of plants by regulating water and inorganic nutrition metabolism. The plasticity of carbonic anhydrase determines the adaptability of plants to the environment. Plants can carry out efficient inorganic carbon assimilation by regulating the diversity and plasticity of carbonic anhydrase in the case of insufficient inorganic carbon sources. Finally, we also elaborated on the possible significance of Mn-substituted CA in the origin and evolution of life. Keywords Carbonic anhydrase · Bicarbonate utilization · Isoenzyme · Biodiversity · Plasticity · Carbon sources · Photosynthetic assimilation

© The Author(s), under exclusive license to Springer Nature Singapore Pte Ltd. 2023 Y. Wu and S. Rao, Root-Derived Bicarbonate Assimilation in Plants, https://doi.org/10.1007/978-981-99-4125-4_3

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3 The Diversity, Plasticity and Roles of Carbonic Anhydrase in Inorganic …

3.1 Introduction Carbonic anhydrase (CA, EC4.2.1.1) is one of the key enzymes using bicarbonate as a substrate. The catalytic mechanism, gene expression and functions of carbonic anhydrase in various organelles of plant cells have been well reviewed (Prince and Woolley 1973; Badger and Price 1994; Rudenko et al. 2015; DiMario et al. 2017a, b, 2018). In this chapter, we will focus on the diversity, plasticity and roles of carbon anhydrase in inorganic carbon utilization in plants and discuss how the utilization of inorganic carbon in plants can cope with the complex and changing biological and abiotic environment through the action of carbonic anhydrase.

3.2 Distribution and Properties of Carbonic Anhydrase 3.2.1 Ubiquitous Carbonic Anhydrase Since carbonic anhydrase was purified from bovine red blood corpuscles (erythrocytes) in 1933 (Meldrum and Roughton 1933), it has been widely studied by biologists. Bradfield (1947) determined the activity of CA in many plants (Bradfield 1947). Afterwards, scientists proved that CA is ubiquitous and plays a very important role in the fixation of CO2 in plants (Badger and Price 1994). In prokaryotes, CA was first identified in Neisseria sicca in 1963 (Veitch and Blankenship 1963). To date, CA has been found in many Archaea and anaerobes (Alber and Ferry 1994; Smith and Ferry 2000; Ferry 2013) and even in fungi (Schlicker et al 2009; Capasso and Supuran 2016). Seemingly, carbonic anhydrase is ubiquitous in almost all types of organisms in the soil, water medium, and on the surface of rock (Bradfield 1947; Veitch and Blankenship 1963; Badger and Price 1994; Smith and Ferry 2000; Supuran 2008a, b; Gilmour and Perry 2009; Capasso and Supuran 2015, 2016; Emino˘glu et al. 2016). Carbonic anhydrase also exists widely in various organelles. According to the location of CAs in cells, carbonic anhydrases can be divided into two categories: external and internal carbonic anhydrases. External CAs, which are linked to the cell surface by metal ions, are distributed on the plasma membrane (called membranebound CA) and in the periplasmic space (called periplasmic CA). Internal CA is distributed in the inner plasma membrane. According to the organelle location, carbonic anhydrase can be divided into chloroplast, mitochondrial, and cytosolic carbonic anhydrase (Badger and Price 1994; Moroney et al. 2001; Bertucc et al. 2011). Even in the carboxysomes of chemoautotrophs and cyanobacteria, CA, which is called carboxysomal CA, is also found (Cannon et al. 2010). Moreover, chloroplast CA can be divided into chloroplast cytosolic CA, chloroplast stroma CA and thylakoid CA (Badger and Price 1994; Moroney et al. 2001).

3.3 Biodiversity of Carbonic Anhydrase

57

3.2.2 Properties of Carbonic Anhydrase The carbonic anhydrase of bovine erythrocytes is a zinc-containing metalloenzyme with a molecular weight of approximately 30,000 Da. Its structure consists of a coiled protein chain and a zinc (II) ion, which is in a distorted tetrahedral coordination geometry (Kannan et al. 1977; Vedani et al. 1989). CA catalyzes the reversible conversion between CO2 and HCO3 − (Prince and Woolley 1973). During dehydration, HCO3 − reacts with Zn-H2 O in the active center of the enzyme, which makes the reaction (Eq. (3.1)) move quickly to the right, while during hydration, CO2 reacts with Zn-OH in the active center, which makes the reaction move quickly to the left (Eq. (3.1)). + HCO− 3 + H ↔ CO2 + H2 O

(3.1)

Compared with 1 Min in the absence of CA, the equilibrium of the above reaction (Eq. (3.1)) takes only 10–6 s in the presence of CA. Moreover, carbonic anhydrase catalyzes the hydration of aldehydes, the hydrolysis of esters, and other reactions whose common factor is attack by nucleophilic oxygen (Prince and Woolley 1973).

3.3 Biodiversity of Carbonic Anhydrase 3.3.1 Functional Diversity of Carbonic Anhydrase Carbonic anhydrase plays various physiological functions in organisms and has high biological significance. CA can promote respiration to generate energy by removing CO2 /HCO3 − (Henry 1996). It can also convert CO2 into HCO3 − and remove CO2 from the decarboxylation reaction, thus stimulating the decarboxylation reaction (Moroney et al. 2001). CA catalyzes the reversible hydration of CO2 and promotes the diffusion of CO2 to Rubisco while providing protons for photophosphorylation during photosynthesis in plants (Graham and Reed 1971). It is also essential for respiration, acid–base homeostasis, CO2 /HCO3 − transfer, ion transport, biosynthesis and calcification (Coulson and Herbert 1984; Tashian, 1989; Badger and Price 1994; Müller et al. 2014). CA can also catalyze other reactions, such as the hydration of acetaldehyde, the hydrolysis of carboxylic acid esters and halogen derivatives, the methanogenesis of Methanosarcina thermophila, the decomposition of cyanate into ammonia by some bacteria, and the absorption of carbonyl sulfide (COS) by some plants and bacteria (Pocker and Meany 1965; Kozliak et al. 1995; ProtoschillKrebs et al. 1996; Innocenti et al. 2004a, b; Stimler et al. 2012; Ogawa et al. 2016). Phosphate, sulfate, phenolic, sulfonate, and carboxylic acid esters can also be used as substrates of CA (Cabiscol and Levine 1996; Freskgard et al. 1992; Tanc et al. 2015). In the future, an increasing number of physiological functions and actions of carbonic anhydrase are being discovered.

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3.3.2 Convergent Evolution of Carbonic Anhydrase The structure and function of organisms adapt to each other. The diverse functions of carbonic anhydrase are closely related to its structure of diversification. To date, at least nine CA genetic families, designated α-, β-, γ-, δ-, ζ-, ε-,η-, θ-, and ι-CA, have different amino acid sequences, no significant sequence consistency, and evolve independently (So et al. 2004; Del Prete et al. 2014a, b, c; Supuran 2016; Jensen et al. 2019). Therefore, carbonic anhydrase is an excellent example of the convergent evolution of catalytic function. The α-CAs evolved from a common primitive gene 500–600 million years ago. It is a well-known CA that is mainly present in protozoa, many Gram-negative bacteria, algae, the cytoplasm of green plants, and vertebrates (Hewett-Emmett and Tashian 1996). The β-CAs are mainly present in the chloroplasts of higher plants of monoas well as di-cotyledons and algae and play an essential role in the acquisition of CO2 and the maintenance of CO2 concentration during photosynthesis (Fukuzawa et al. 1992; Eriksson et al. 1996; Dimario et al. 2017a, b); meanwhile, β-CAs are also found in both Gram-negative and Gram-positive bacteria, many fungi and some Archaea (Gotz et al. 1999; Smith et al. 1999; Supuran 2008a). γ-CAs were first found in Methanosarcina thermophila in 1994 (Alber and Ferry 1994). At present, the gene encoding the γ-CA structural protein is widely found in Archaea, cyanobacteria, most types of bacteria, fungi and plants (Supuran 2008a, b; Del Prete et al. 2016a, b, c; DiMario et al. 2018). Six other carbonic anhydrase genetic families have also been found in addition to the three common carbonic anhydrase classes mentioned above. δ- and ζ-CAs are mainly found in marine diatoms (Roberts et al. 1997; Lane et al. 2005; Xu et al. 2008; Del Prete et al. 2014a). ε-CAs are mainly present in cyanobacteria and some chemoautotrophic bacteria (So and Espie 2005; So et al. 2004), whereas η-CAs are present in protozoa (Del Prete et al. 2014b). θ-CAs have recently been found in the diatom Phaeodactylum tricornutum (Kikutani et al. 2016) and the chlorophyte Chlamydomonas reinhardtii (Jin et al. 2016). ι-CAs have recently been found in the marine diatom Thalassiosira pseudonana (Jensen et al. 2019) and the Gram-negative bacterium Burkholderia territorii (Del Prete et al. 2020).

3.3.3 Coordination Diversity of Metal Cofactors and Amino Acid Residues in Carbonic Anhydrase The CAs are metalloenzymes that are catalytically effective only with one metal ion bound within the active site cavity. The active center normally comprises M(II) ions in a tetrahedral geometry, with three amino acid residues as ligands, in addition to a water molecule/hydroxide ion coordinating the metal (Supuran 2016). However, the species of metal ions and their coordination of amino acid residues are different among carbonic anhydrase genetic families.

3.3 Biodiversity of Carbonic Anhydrase

59

α-CAs, which have been widely studied, are normally monomers and rarely dimers, and their molecular weight is approximately 30 kDa. The active site of α-CAs consists of a Zn(II) coordinated by three histidine residues and a water molecule or hydroxide ion (Moroney et al. 2001), whereas Co(II) may substitute for Zn(II) in many α-CAs without a significant loss of catalytic activity (Supuran 2016). The molecular weight of β-CAs is 100–200 kDa, and β-CAs are composed of dimers, tetramers or octamers of 23–25 kDa subunits. The crystal structure shows that Zn(II) in its active site is linked with two conserved cysteine residues and one conserved histidine residue (Bracey et al. 1994; Rowlett et al. 1994). γ-CAs, which may have evolved from 3 to 4 billion years ago, have a left-handed parallel β-helix folded structure and are homotrimers containing three metal atoms, and their molecular weight is approximately 69 kDa. The metal ion of the active center is Fe (II), which is coordinated by three histidine residues and a water molecule or hydroxide ion (Hewett-Emmett and Tashian 1996; Tripp et al. 2004), but Zn (II) and Co (II) can replace Fe (II) to perform physiological reaction catalysis (MA et al. 2015). δ-CAs, with a molecular weight of approximately 27 kDa, have an active site similar to that of α-CAs. The metal ion of the active center is Cd(II) or Zn(II), which is coordinated by three histidine residues and a water molecule or hydroxide ion (Roberts 1997; Xu et al. 2008). The ζ-CA is a structural mimic of a functional β-CA dimer, with a molecular weight of 69 kDa. Its active site probably has a roughly tetrahedral geometry, and the metal ion is bound by two or more thiolates, as in the zinc-containing β-CAs in higher plants. The metal ion of the active center is Cd(II) or Zn(II), which is coordinated by two cysteines, a histidine and a water molecule or hydroxide ion (Lane et al. 2005; Xu et al. 2008). The predicted molecular masses of the ε-CA polypeptides range from 55.2 to 63.4 kDa. The metal ion of the active center is Zn(II), which is coordinated by three histidine residues or a combination of histidine, cysteine, and sometimes aspartate (So and Espie 2005; So et al. 2004). The 3D fold of the η-CAs is similar to that of the δ-CAs, but the pattern of Zn(II) coordination seems to be completely different from that of the δ-CAs, with two histidine and one glutamine residue in addition to the water molecule/hydroxide ion binding the Zn(II) (Simone et al. 2015; Del Prete et al. 2016b). θ-CAs have a predicted molecular mass of approximately 85 kDa. At least, Zn(II) was probably one of the metal ions coordinated by amino acid residues in the active center (Kikutani et al. 2016). However, in ι-CAs, Mn(II) is probably a metal ion that is coordinated by two histidine, one glutamate, and aspartate residues (Jensen et al. 2019).

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3 The Diversity, Plasticity and Roles of Carbonic Anhydrase in Inorganic …

3.3.4 Diversity of Response and Sensitivity to Inhibitors and Activators 3.3.4.1

The Inhibition Pattern of Halides Against CAs

It can be seen from the above that the structures of CAs and the metal coordination patterns of their active centers are diverse. Therefore, the binding behavior of different inorganic and organic inhibitors against CAs varied. Inhibition constants, which were obtained under the same conditions using recombinant highly purified enzymes and a stopped flow assay monitoring the hydration of CO2 to bicarbonate, can reflect the diversity of response and sensitivity to inhibitors (Simone and Supuran 2012). The inhibition pattern of the CAs shown in Table 3.1 with halides demonstrates how differently these isozymes interact with these inhibitors. As shown in Table 3.1, the inhibitory effects of different halides on different types of CA isoenzymes are different, and these differences can reach 6 orders of magnitude. The inhibition effects of the same halide on different types of CA isoenzymes are not the same, and these differences reach 5 to 6 orders of magnitude. The inhibition effects of different halides on the same type of CA isoenzyme are also different, and these differences are relatively small; the majority of these differences are less than an order of magnitude, and the largest difference is less than 3 orders of magnitude (except for h CAI). This shows that different halides or even the same halide have different inhibitory effects on different types or the same type of CA isoenzyme.

3.3.4.2

The Inhibition Pattern of Inorganic Anions (Except for Halides) Against CAs

The inhibition pattern of the CAs shown in Table 3.2 with HCO3 − , CO3 2− , NO3 − , NO2 − , HSO3 − , and SO4 2− demonstrates how differently these isozymes interact with these inhibitors. As shown in Table 3.2, the inhibitory effects of different nutritional anions and even the same nutritional anions on different types or the same type of CA isoenzymes are different, and these differences can reach 5 orders of magnitude. This shows that different nutritional anions or even the same nutritional anion have different inhibitory effects on different types or the same type of CA isoenzymes.

3.3.4.3

The Inhibition Pattern of CA Inhibitors and Organic Acids Against CAs

The inhibition pattern of the CAs shown in Table 3.3 with acetozolamide (AZ), ethoxzolamide (EZ), oxalate, malate, and citrate demonstrates how differently these isozymes interact with these inhibitors.

3.3 Biodiversity of Carbonic Anhydrase

61

Table 3.1 Inhibition constants (mM) of halides against different classes of CA isozymes for the CO2 hydration reaction at 20°C Cl−

Br−

>300

6

4

0.3

Simone and Supuran (2012)

Human

>300

200

63

26

Simone and Supuran (2012)

h CAIII

Human

78.5

0.98

0.96

0.9

Simone and Supuran (2012)

α

h CAIV

Human

0.07

0.09

0.09

0.08

Simone and Supuran (2012)

α

h CAVA

Human

241

156

50

25

Simone and Supuran (2012)

α

h CAVB

Human

11

43

72

71

Simone and Supuran (2012)

α

h CAVI

Human

0.6

0.72

0.73

0.81

Simone and Supuran (2012)

α

h CAVII

Human

1.24

1.84

1.06

0.25

Simone and Supuran (2012)

α

h CAIX

Human

48

33

16

7

Simone and Supuran (2012)

α

h CAXII

Human

0.56

73

82

215

Simone and Supuran (2012)

α

m CAXIII

Murine

3

138

45

5.4

Simone and Supuran (2012)

α

h CAXIV

Human

37

0.77

0.77

0.78

Simone and Supuran (2012)

α

m CAXV

Murine

0.054

0.085

0.104

0.288

Simone and Supuran (2012)

α

SspCA

Sulfurihydrogenibium yellowstonensis

41.7

8.30

49.0

0.86

Vullo et al. (2014a)

α

SazCA

Sulfurihydrogenibium azorense

0.98

0.85

0.94

0.87

Vullo et al. (2012)

Classes

Isozymes

Source

α

h CAI

Human

α

h CAII

α

F−

I−

References

(continued)

62

3 The Diversity, Plasticity and Roles of Carbonic Anhydrase in Inorganic …

Table 3.1 (continued) F−

Cl−

Br−

I−

Classes

Isozymes

Source

α

TcCA

Trypanosoma cruzi

0.90

0.81

0.73

0.044

Del Prete et al. (2014a)

References

α

LjCAA1

Lotus japonicus

0.73

0.87

41.1

76.9

Vullo et al. (2014b)

α

LjCAA2

Lotus japonicus

7.3

8.7

6.4

7.6

Vullo et al. (2014b)

α

SpiCA1

Stylophora pistillata

0.62

0.50

0.0097

α

SpiCA2

Stylophora pistillata

0.92

0.53

0.96

33.0

Del Prete et al. (2018)

α

SpiCA3

Stylophora pistillata

0.48

0.51

0.23

0.56

Del Prete et al. (2018)

β

Cab

Methanobacterium thermoautotrophicum

>1000

152

42.1

132

Zimmerman and Supuran (2007)

β

PCA

Streptococcus pneumoniae

0.85

0.052

0.046

0.054

Simone and Supuran (2012)

β

stCA1

Salmonella typhimurium

0.97

0.77

0.92

0.8

Simone and Supuran (2012)

β

stCA2

Salmonella typhimurium

0.63

0.49

0.68

0.64

Simone and Supuran (2012)

β

Can2

Cryptococcus neoformans

086

0.92

1.00

1.11

Simone and Supuran (2012)

β

caNce103

Candida albicans

0.69

0.85

0.94

1.4

Simone and Supuran (2012)

β

scCA

Saccharomyces cerevisiae

2.85

0.85

0.0108

β

cgNce103

Candida glabrata

0.36

0.58

27

42.4

Simone and Supuran (2012)

β

DmBCA

Drosophila melanogaster

0.80

0.97

1.04

1.18

Simone and Supuran (2012)

β

CAS1

Sordaria macrospora

>100

9.2

9.3

8.6

Vullo et al. (2020)

0.0090 Del Prete et al. (2018)

0.0103 Simone and Supuran (2012)

(continued)

3.3 Biodiversity of Carbonic Anhydrase

63

Table 3.1 (continued) F−

Cl−

Br−

I−

Classes

Isozymes

Source

β

CAS2

Sordaria macrospora

>100

>100

>100

7.7

Vullo et al. (2020)

References

β

CAS3

Sordaria macrospora

>100

>100

>100

9.9

Vullo et al. (2020)

β

HpyCA

Helicobacter pylori

0.67

0.56

0.38

0.63

Vullo et al. (2014c)

β

CahB1

Coleofasciculus chthonoplastes

0.53

0.74

0.63

0.66

Vullo et al. (2014c)

β

PgiCAb

Porphyromonas gingivalis

7.8

7.5

15.9

21.4

Vullo et al. (2014c)

β

FbiCA 1

Flaveria bidentis

0.71

0.74

0.67

0.71

Vullo et al. (2014c)

β

GsaCAβ

Gyrodactylus salaris

5.5

3.3

8.2

>50

Aspatwar et al. (2022)

γ

Zn-Cam

Methanosarcina thermophila

>200

>200

160

160

Zimmerman and Supuran (2007)

γ

Co-Cam

Methanosarcina thermophila

>200

>200

22.2

5.3

Zimmerman and Supuran (2007)

γ

PgiCAa

Porphyromonas gingivalis

0.95

0.94

0.92

8.7

Del Prete et al. (2017)

γ

NcoCA

Nostoc commune

>100

8.7

9.3

8.9

Del Prete et al. (2017)

γ

BpsγCA

Burkholderia pseudomallei

>100

>100

>100

>100

Del Prete et al. (2017)

γ

PhaCAγ

Pseudoalteromonas haloplanktis

>100

>100

8.7

>100

Del Prete et al. (2017)

ζ

Cd-Rl

Thalassiosira weissflflogii

0.53

0.76

0.85

1.12

Simone and Supuran (2012)

ζ

Zn-Rl

Thalassiosira weissflflogii

0.36

0.41

0.53

0.61

Simone and Supuran (2012)

δ

TweCA

Thalassiosira weissflflogii

5.8

>200

8.3

1.9

Del Prete et al. (2014a)

η

PfCA

Plasmodium falciparum

5.78

9.76

6.05

2.74

Del Prete et al. (2014b)

TcCA

LjCAA1

LjCAA2

SpiCA1

α

α

SazCA

α

α

SspCA

α

α

h CAXIV

m CAXV

α

m CAXIII

α

α

h CAIX

h CAXII

α

h CAVII

α

α

h CAVB

h CAVI

α

h CAVA

α

α

h CAIII

h CAIV

α

α

h CAI

h CAII

α

α

Isozymes

Classes

Stylophora pistillata

Lotus japonicus

Lotus japonicus

Trypanosoma cruzi

Sulfurihydrogenibium azorense

Sulfurihydrogenibium yellowstonensis

Murine

Human

Murine

Human

Human

Human

Human

Human

Human

Human

Human

Human

Human

Source

0.45

3.9

5.4

0.58

15.70

33.2

0.008

1.1

140

0.75

13

0.16

0.8

0.71

82

6.6

0.74

85

12

HCO3 −

0.01

6.1

0.45

0.69

7.60

39.3

0.106

0.98

5.5

0.64

29

0.27

0.69

0.93

95

5.7

0.01

73

15

CO3 2−

0.56

7.0

5.2

0.77

0.76

0.86

0.068

0.81

36

79

46

0.19

0.76

0.72

16

58

117

35

7

NO3 −

0.77

3.7

4.1

0.70

0.58

0.48

0.061

0.92

12.6

94

42

1.78

0.82

0.8

16

31

53

63

8.4

NO2 −

0.41

7.4

5.3

0.91

0.95

21.1

0.01

0.76

75

0.84

75

7.3

14.2

0.72

65

13.2

1.06

89

18

HSO3 −

0.91

51.9

7.1

6.9

10.0

0.82

0.01

0.01

>200

0.77

>200

1.38

9.9

0.83

1.17

9.0

1.00

>200

63

SO4 2−

Del Prete et al. (2018)

Vullo et al. (2014b)

Vullo et al. (2014b)

Del Prete et al. (2014c)

Vullo et al. (2012)

Vullo et al. (2014a)

(continued)

Simone and Supuran (2012)

Simone and Supuran (2012)

Simone and Supuran (2012)

Simone and Supuran (2012)

Simone and Supuran (2012)

Simone and Supuran (2012)

Simone and Supuran (2012)

Simone and Supuran (2012)

Simone and Supuran (2012)

Simone and Supuran (2012)

Simone and Supuran (2012)

Simone and Supuran (2012)

Simone and Supuran (2012)

References

Table 3.2 Inhibition constants (mM) of HCO3 − , CO3 2− , NO3 − , NO2 − , HSO3 − , and SO4 2− against different classes of CA isozymes for the CO2 hydration reaction at 20°C

64 3 The Diversity, Plasticity and Roles of Carbonic Anhydrase in Inorganic …

Can2

caNce103

scCA

β

β

β

CAS1

CAS2

CAS3

HpyCA

β

β

β

β

cgNce103

stCA2

β

DmBCA

stCA1

β

β

PCA

β

β

SpiCA3

Cab

α

SpiCA2

α

β

Isozymes

Classes

Table 3.2 (continued)

Helicobacter pylori

Sordaria macrospora

Sordaria macrospora

Sordaria macrospora

Drosophila melanogaster

Candida glabrata

Saccharomyces cerevisiae

Candida albicans

Cryptococcus neoformans

Salmonella typhimurium

Salmonella typhimurium

Streptococcus pneumoniae

Methanobacterium thermoautotrophicum

Stylophora pistillata

Stylophora pistillata

Source

0.50

3.4

5.5

6.5

26.9

0.086

0.78

0.62

0.75

27.9

0.64

0.33

44.9

0.40

7.81

HCO3 −

0.42

8

8.8

>100

0.86

0.31

0.76

0.01

0.6

6.53

0.54

0.53

9.6

5.66

0.24

CO3 2−

0.78

8.5

>100

>100

43.7

0.097

13.9

0.69

0.92

0.27

0.50

0.39

7.8

12.8

0.99

NO3 −

0.67

8.3

>100

>100

28.6

0.088

0.46

0.53

0.96

0.32

0.70

0.66

44.8

0.45

3.15

NO2 −

0.63

>100

7.3

3.3

1.29

0.1

0.33

0.54

0.71

0.68

0.93

0.57

45.1

5.20

0.43

HSO3 −

0.57

>100

4.8

>100

1.36

0.58

0.58

14.15

0.86

0.71

0.94

4.15

950

0.61

0.33

SO4 2− References

Vullo et al. (2014c)

Vullo et al. (2020)

Vullo et al. (2020)

Vullo et al. (2020)

(continued)

Simone and Supuran (2012)

Simone and Supuran (2012)

Simone and Supuran (2012)

Simone and Supuran (2012)

Simone and Supuran (2012)

Simone and Supuran (2012)

Simone and Supuran (2012)

Simone and Supuran (2012)

Zimmerman and Supuran (2007)

Del Prete et al. (2018)

Del Prete et al. (2018)

3.3 Biodiversity of Carbonic Anhydrase 65

FbiCA 1

GsaCAβ

Zn-Cam

Co-Cam

PgiCAa

NcoCA

BpsγCA

PhaCAγ

Cd-Rl

Zn-Rl

TweCA

PfCA

γ

γ

γ

γ

γ

γ

ζ

ζ

δ

η

PgiCAb

β

β

CahB1

β

β

Isozymes

Classes

Table 3.2 (continued)

Plasmodium falciparum

Thalassiosira weissflflogii

Thalassiosira weissflflogii

Thalassiosira weissflflogii

Pseudoalteromonas haloplanktis

Burkholderia pseudomallei

Nostoc commune

Porphyromonas gingivalis

Methanosarcina thermophila

Methanosarcina thermophila

Gyrodactylus salaris

Flaveria bidentis

Porphyromonas gingivalis

Coleofasciculus chthonoplastes

Source

0.78

0.89

0.10

0.12

4.2

6.2

4.5

0.96

0.10

42

>50

0.66

7.3

0.61

HCO3 −

0.90

2.5

0.11

0.13

8.3

2.5

2.4

0.89

0.009

6.7

>50

0.84

3.7

0.68

CO3 2−

0.66

0.97

0.21

0.82

24.1

5.6

27.6

8.5

0.09

36.5

>50

0.78

>200

1.86

NO3 −

2.46

3.1

0.58

0.88

6.8

6.7

8.8

3.1

7.3

6.8

9.1

0.57

7.8

1.38

NO2 −

>100

8.2

0.34

0.63

>100

24.1

14.9

9.3

1.8

11.7

6.2

55.3

>200

0.51

HSO3 −

9.2

>200

0.24

0.48

>100

>100

>100

8.7

> 200

>200

>50

0.62

>200

7.9

SO4 2− References

Del Prete et al. (2014b)

Del Prete et al. (2014a)

Simone and Supuran (2012)

Simone and Supuran (2012)

Del Prete et al. (2017)

Del Prete et al. (2017)

Del Prete et al. (2017)

Del Prete et al. (2017)

Zimmerman and Supuran (2007)

Zimmerman and Supuran (2007)

Aspatwar et al. (2022)

Vullo et al. (2014c)

Vullo et al. (2014c)

Vullo et al. (2014c)

66 3 The Diversity, Plasticity and Roles of Carbonic Anhydrase in Inorganic …

Isozymes

h CAI

h CAII

h CAIV

h CAVA

h CAIX

SazCA

Cab

Can2

caNce103

CAS1

CAS2

CAS3

CahB1

FbiCA 1

GsaCAβ

Zn-Cam

Co-Cam

PgiCAa

Classes

α

α

α

α

α

α

β

β

β

β

β

β

β

β

β

γ

γ

γ

Porphyromonas gingivalis

Methanosarcina thermophila

Methanosarcina thermophila

Gyrodactylus salaris

Flaveria bidentis

Coleofasciculus chthonoplastes

Sordaria macrospora

Sordaria macrospora

Sordaria macrospora

Candida albicans

Cryptococcus neoformans

Methanobacterium thermoautotrophicum

Sulfurihydrogenibium azorense

Human

Human

Human

Human

Human

Source

324

1.43

0.063

460

27

76

94 ± 3.0

816

445

10.5

12.1

0.9

12

250

AZ 8

25

0.74

0.20

95 ± 2.8

3170

440

5.35

EZ

37.7

67.7

3280

2230

0.99

990

oxalate

87.7

899

7420

2490

53.7

1870

malate

39.1

88.6

4930

1670

0.99

2160

citrate

Del Prete et al. (2017) (continued)

Zimmerman and Supuran (2007)

Zimmerman and Supuran (2007)

Aspatwar et al. (2022)

Vullo et al. (2014c)

Vullo et al. (2014c)

Vullo et al. (2020)

Vullo et al. (2020)

Vullo et al. (2020)

Simone and Supuran (2012)

Simone and Supuran (2012)

Zimmerman and Supuran (2007)

Vullo et al. (2012)

Simone and Supuran (2012)

Simone and Supuran (2012)

Simone and Supuran (2012)

Simone and Supuran (2012)

Simone and Supuran (2012)

References

Table 3.3 Inhibition constants (nM) of acetozolamide (AZ), ethoxzolamide (EZ), oxalate, malate, and citrate against different classes of CA isozymes for the CO2 hydration reaction at 20°C

3.3 Biodiversity of Carbonic Anhydrase 67

Isozymes

Cd-Rl

Zn-Rl

TweCA

PfCA

Classes

ζ

ζ

δ

η

Table 3.3 (continued)

Plasmodium falciparum

Thalassiosira weissflflogii

Thalassiosira weissflflogii

Thalassiosira weissflflogii

Source

83

58

82

170

AZ

EZ

oxalate

malate

citrate

Del Prete et al. (2014b)

Del Prete et al. (2014a)

Simone and Supuran (2012)

Simone and Supuran (2012)

References

68 3 The Diversity, Plasticity and Roles of Carbonic Anhydrase in Inorganic …

3.4 Diversity of Carbonic Anhydrase in Plants

69

As shown in Table 3.3, the inhibitory effects of AZ and EZ on different types or the same type of CA isoenzyme vary widely, and these differences can reach 4 orders of magnitude. The inhibitory effect of different organic acids on the same type of CA isoenzyme does not differ much. In general, the inhibition of carbonic anhydrase by organic acids is much greater than that of inorganic anions, which plays an important role in the physiological environment. That is, in the sites of organic acid metabolism, CA does not play a role because its activity is almost completely inhibited. In most organelles, inorganic anions have difficulty effectively inhibiting CA activity due to low concentrations, which ensures that other substance (such as carbon, nitrogen, phosphorus, sulfur) metabolism and CA action can be carried out at the same time. For example, the assimilation of nitrates in plants does not affect the role of CA, ensuring the coupling of carbon and nitrogen.

3.3.4.4

The Activation Profile of CAs with Amino Acids and Amines

The activation profile of CAs with amino acids and amines shown in Table 3.4 demonstrates how differently these isozymes interact with these activators. As shown in Table 3.4, the activation effects of different amino acids and amines and even the same amino acid or amine on different types or the same type of CA isoenzymes are different, and these differences can reach 4 orders of magnitude. This shows that different amino acids and amines or even the same amino acid and amine have different activation effects on different types or the same type of CA isoenzymes.

3.4 Diversity of Carbonic Anhydrase in Plants Plants have several different genes encoding CA. Cyanobacteria has at least 5 genes encoding CA isoenzymes, and Chlamydomonas reinhardtii has at least 13 genes encoding CA isoenzymes. Oryza sativa has at least 16 genes encoding CA isoenzymes, and in Arabidopsis and sorghum, there are also at least 17 genes encoding CA isoenzymes. Soybean has at least 25 genes encoding CA isoenzymes. Allotetraploid Triticum aestivum has at least 79 genes encoding CA isoenzymes. Table 3.5 shows the number of genes encoding CA in different plant species. In addition, the distribution and expression of different types of CA isoenzymes in tissues and organs also have significant differences. In sorghum, of the 9 genes encoding α-class CA, at least 4 genes were highly expressed, of which 2 genes were extremely highly expressed in leaves and anthers, respectively; five genes encoding β-class CA were highly expressed in different tissues and organs, and 2 genes were highly expressed in leaves; three genes encoding γ-class CA were most

70

3 The Diversity, Plasticity and Roles of Carbonic Anhydrase in Inorganic …

Table 3.4 Activation constant (μM) of amino acids and amines on different classes of CA isozymes for the CO2 hydration reaction at 20°C (Akocak and Supuran 2019) Classes Isozymes Source

L-His L-Phe L-Trp L-Tyr Histamine Serotonin

α

h CAI

Human

0.03

0.07

44

0.02

2.1

α

h CAII

Human

10.9

0.013

27

0.011

125

50

α

h CAIII

Human

35.9

34.7

20.5

34.1

36.9

0.78

α

h CAIV

Human

7.30

36.3

37.1

25.1

25.3

3.14

α

h CAVA

Human

1.34

9.81

1.13

2.45

0.010

6.33

α

h CAVB

Human

0.97

10.45

0.89

0.044

3.52

0.11

α

h CAVII

Human

0.92

10.93

57.5

20.3

37.5

0.93

α

h CAIX

Human

9.71

16.3

37.5

25.3

35.1

33.1

α

h CAXII

Human

37.5

1.38

26.0

25.8

27.9

0.30

α

m CAXIII

Murine

0.13

1.02

16

4.6

0.51

α

h CAXIV

Human

0.90

0.24

16.5

21.8

0.010

6.5

α

m CAXV

Murine

32.1

33.4

13.5

8.9

18.5

7.5

β

Cab

Methanobacterium thermoautotrophicum

69

70

16.9

10.5

76

62

β

scCA

Saccharomyces cerevisiae

82

86

91

85

20.4

15.0

γ

Zn-Cam

Methanosarcina thermophila

68

68

38

24

63

38

γ

Co-Cam

Methanosarcina thermophila

135

70

47

53

9.2

0.97

γ

BpsγCA

Burkholderia pseudomallei

24.7

1.73

0.43

0.20

0.12

0.10

γ

PhaCA

Pseudoalteromonas haloplanktis

12.6

15.8

7.12

1.02

0.48

9.05

δ

TweCA

Thalassiosira weissflflogii

0.75

2.15

0.93

1.52

1.34

0.90

η

PfCA

Plasmodium falciparum

1.06

0.43

5.21

1.02

9.86

7.18

45

highly expressed in anthers (Makita et al. 2014; DiMario et al. 2017a, b). Differential expression of genes encoding different CA isoenzymes provides the basis for the functional diversity of CA. This conclusion can be drawn based on the structural and functional diversity of carbonic anhydrase and the extent of its regulation by organic and inorganic

3.4 Diversity of Carbonic Anhydrase in Plants

71

Table 3.5 Total number of genes encoding CA in different plant species (From DiMario et al. 2017a, b; Jensen et al. 2020; Momayyezi et al. 2020) Plant type

Species

Photosynthetic type

Algae

Cyanobacteria

C3

CA classes and the number of genes encoding CA α

Moss Monocots

Dicots

1

β

γ

δ

ζ

θ

4

Chlamydomonas reinhardtii

C3

3

6

3

Phaeodactylum tricornutum

C3

5

2

2

Thalassiosira pseudonana

C3

1

Selaginella moellendorffii

C3

10

5

4

4

Physcomitella patens

C3

5

6

5

Brachypodium distachyon

C3

6

4

3

Oryza sativa

C3

9

3

4

Hordeum vulgare

C3

14

9

1

Triticum aestivum

C3

49

23

7

Sorghum bicolor

C4

9

5

3

Setaria italica

C4

9

4

3

Zea mays

C4

9

8

1

Ananas comosus

CAM

4

3

3

Arabidopsis thaliana

C3

8

6

5

Brassica oleracea

C3

7

10

2

Solanum lycopersicum

C3

9

5

1

Medicago truncatula

C3

8

7

4

Gossypium raimondii

C3

10

10

2

Citrus clementina

C3

5

5

1

Medicago trunculata

C3

9

10

1

Lactuca sativa

C3

8

7

1

Prunis persica

C3

11

6

1

Populus trichocarpa

C3

8

7

5

1 3

1

substances. Because carbonic anhydrase has extensive structural and functional diversity, it leads to the diversity of carbon metabolic pathways such as carbon migration and conversion. There are several orders of magnitude differences in the activation or inhibition effects of organic and inorganic substances on different carbonic anhydrase isoenzymes, which resulted in each well-organized carbon metabolism pathway. In other words, carbonic anhydrase is the regulating switch of carbon metabolism and its related metabolism. Carbonic anhydrase is an important switch of plant substance metabolism.

72

3 The Diversity, Plasticity and Roles of Carbonic Anhydrase in Inorganic …

3.5 Plasticity of Carbonic Anhydrase Carbonic anhydrase is an inducible enzyme, and its activity is greatly affected by environmental factors, mainly water, pH, light, CO2 and HCO3 − concentration as well as nutrient concentrations. In particular, the decrease in CO2 concentration in the environment has led to increasing carbonic anhydrase activity (Merrett et al. 1996; Moroney and Somanchi 1999; Burkhardt et al. 2001; Badger et al. 2002; Chen and Gao 2003; Yoshioka et al. 2004; Price 2011; Hofmann et al. 2013; Sun et al. 2016; Aslam et al. 2018; Hang and Wu 2019; Jones et al. 2021).

3.5.1 Response of Carbonic Anhydrase to Water Water is the raw material of photosynthesis, and photosynthesis cannot be carried out without water. As an important regulating enzyme of photosynthesis, CA also shows different activities due to different water conditions to adapt to different water environments and meet the requirements of photosynthesis. Under drought stress, the change in CA activity is consistent with the change in photosynthetic rate, and CA may affect the whole photosynthesis process (Downton and Slatyer 1972). When the flag leaf of rice underwent soil drought treatment at the reversible decline stage, if the drought was not very serious, photosynthetic parameters such as stomatal conductance decreased correspondingly, but the CA activity increased significantly due to the induction of adversity, which promoted the increase in stomatal conductance of mesophyll cells and the increase in the effective supply of CO2 and compensated for the decrease in the photosynthetic rate due to stomatal closure to a certain extent. The reciprocal fifth leaves were in the irreversible decline stage, and their photosynthetic organs had declined. Drought treatment did not induce an increase in CA activity but caused a large decline. This indicates that CA activity has a regulatory effect on the photosynthetic rate under water stress conditions, and the fact that some varieties of plants can still maintain a high photosynthetic rate under water stress conditions may be the reason for their adaptability, so the response of CA to water stress may be one of the indicators of the adaptability of varieties (Dai et al. 2000). Dehydration and polyethylene glycol-induced osmotic stress have significant effects on carbonic anhydrase gene expression in Arabidopsis (Wu et al. 2012). Under water stress, the expression of carbonic anhydrase genes in olive trees was downregulated and upregulated after rehydration (Perez-Martin et al. 2014). Under severe drought, the expression of carbonic anhydrase genes in Orychophragmus violaceus was significantly higher than that under well-watered conditions. However, the expression of carbonic anhydrase genes in tomato leaves was significantly lower than that in well-watered leaves (Sun et al. 2016).

3.5 Plasticity of Carbonic Anhydrase

73

3.5.2 Response of Carbonic Anhydrase to pH The pH can regulate the enzymatic reaction of plants. Each enzyme in the cells has an optimum pH value and an optimum pH microenvironment. Even a subtle change in pH will lead to a change in the rate of enzymatic reaction. Under acidic conditions, inorganic carbon mainly exists in the form of CO2 ; under alkaline conditions, inorganic carbon mainly exists in the form of HCO3 − , which must be rapidly converted into CO2 by CA catalysis to meet the needs of plants. The CA activity of Chlorella ellipsoidea and Chlamydomonas reinhardtii grown under acidic conditions is significantly lower than that grown under alkaline conditions (Rotatore and Colman 1991; Janette and John 1994). Alkaline conditions may also directly affect the expression of CA in plants to affect the growth of plants (Williams and Colman 1996). The activity of extracellular CA in Chlamydomonas reinhardtii induced at pH 7.2 was the highest, while that induced at pH 5.5 was significantly lower than that induced at pH 7.2 and pH 9.0 (Chen and Dai 2000). For Dinoflagellates, pH 7.5 has the best induction effect on extracellular CA (Dai et al. 2011). Soil pH is the primary driver of CO2 –H2 O isotopic exchange catalyzed by CA (Sauze et al. 2018; Jones et al. 2021). However, the optimal pH for the photosynthetic O2 evolution function of the lumenal CA (CAH3) from Chlamydomonas reinhardtii was 6.2–6.5 (Terentyev et al. 2019).

3.5.3 Response of Carbonic Anhydrase to Light Intensity CA is an important photosynthetic enzyme involved in CO2 conduction into carboxylation sites, and its activity has a great impact on photosynthesis. Light is an important environmental factor affecting photosynthesis. Plants adapt to changes in the light environment by regulating CA activity. In C3 plants, CA can regulate photosynthesis by promoting the liquid-phase diffusion of CO2 (Burnell et al. 1990). CA (including CA-like photosystem II) catalyzes the conversion of CO2 to bicarbonate and bicarbonate to CO2 required by the photosynthetic carboxylation site. The change in CA activity is regulated by light intensity, which is similar to the change in the Rubisco carboxylation rate. The increase in Rubisco activity and carboxylation rate induced by light is accompanied by a corresponding increase in CA activity (Haglund et al. 1992). Because the enhancement of carboxylation reduces the CO2 partial pressure at the photosynthetic carboxylation site, it is conducive to promoting the catalytic reaction of CA (Rawat and Moroney 1995). The increase in CA activity in rice can increase the affinity of Rubisco for CO2 and improve the photosynthetic CO2 fixation capacity. The DIC transmission rate of Gracllaria fanuishtipzata grown under strong light increased with increasing CA activity. The increase in CA activity is also regarded as one of the means by which rice alleviates photoinhibition under strong light (Ji et al. 1997; Mercado et al. 2000).

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3.5.4 Response of Carbonic Anhydrase to Anions Most anions, such as F− , Cl− , HCO3 − , Br− , I− , SO4 2− , and NO3 − , reduce the activity of the enzyme due to their reduced affinity or competition with the substrate of the enzyme (Maren et al. 1976; Wieth 1979; Innocenti et al. 2004a, b; Nishimori et al. 2007; Nishimori et al. 2009). CA can also bind with carboxylate, phenol, alcohol, imidazole, carboxylic acid phthalamide, thiophthalamide and SCN− , which will inhibit the catalytic activity of CA (Jönsson et al. 1993). Glycine, glucose, acetic acid and other organic carbon inhibit the induction of CA (Umino and Shiraiwa 1991; Merrett et al. 1996; Simone and Supuran 2012). Some amino acids and amines can activate CA (Akocak and Supuran 2019). However, the effect of some anions on carbonic anhydrase depends on the concentration of anions. The gene expression of the chloroplast carbonic anhydrase isoform of Orychophragmus violaceus and the cytosolic carbonic anhydrase isoform of Brassica juncea was promoted at low concentrations and inhibited at high concentrations (Hang and Wu 2019). Fluoride ions below 2.0 mM can obviously promote the extracellular carbonic anhydrase of Chlamydomonas reinhardtii (Wu et al. 2007a, b). A low concentration of bicarbonate ions less than 0.5 mM can promote the extracellular carbonic anhydrase of Chlamydomonas reinhardtii. Beyond this concentration, the inhibition increases with increasing concentration (Wu et al. 2015). Insufficient nitrogen will reduce the activity of CA, and at the same time, it can slowly return to its original activity by increasing nitrate (Burnell et al 1990).

3.5.5 Response of Carbonic Anhydrase to Cations Many heavy metal elements (Co2+ , Cu2+ , Zn2+ , Ag+ , Cd2+ , Pb, and Hg) reduce the activity of carbonic anhydrase because they can either replace zinc in the action center of carbonic anhydrase, directly change the biological activity, or indirectly change the metabolism of carbonic anhydrase (lionetto et al. 1998; Gilbert et al. 2001; Sas et al. 2006; Soyut et al. 2012, Topchiy et al. 2019). Divalent metal ions such as Co2+ , Mn2+ , Ni2+ , Cu2+ , Fe2+ , and Cd2+ can replace the zinc of carbonic anhydrase and show different activities (Lindskog 1963). However, some kinds of heavy metals promote carbonic anhydrase activity at low concentrations and inhibit carbonic anhydrase activity at high concentrations, showing obvious “hormone” effects. Although cadmium can inhibit chloroplast carbonic anhydrase in Spinacia oleracea (Topchiy et al. 2019), when it was added to cultured marine phytoplankton, carbonic anhydrase activity was enhanced (Cullen et al. 1999). Cadmium increases the extracellular carbonic anhydrase activity of Chlamydomonas reinhardtii at low concentrations and inhibits its activity at high concentrations (Wang et al. 2005). Two micromoles of manganese, iron, cobalt, nickel and cadmium can increase the extracellular carbonic anhydrase activity of Chlamydomonas reinhardtii, and when the concentration is 100 μM, it has an inhibitory

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effect (Wang et al. 2006). Adding low concentrations of zinc and iron to culture duck erythrocytes can increase the activity of carbonic anhydrase, while adding high concentrations of zinc and iron to culture duck erythrocytes inhibits the activity of carbonic anhydrase (Wu et al. 2007a, b). The heavy metal hyperaccumulated plant Thraspi caerulescens increased its carbonic anhydrase activity when exposed to cadmium (Liu et al. 2008). Zinc, iron and cadmium (1 mM) increase the extracellular carbonic anhydrase activity of Pseudomonas fragi (Sharma et al. 2009). The application of appropriate Zn enhanced the CA activities of both Zn-efficient and Zn-inefficient genotypes of wheat compared to no Zn (Singh et al. 2019). At 40 mg L−1 , zinc can induce the expression of the β-CA family in millet (Cao et al. 2020). In the range of 0–50 μM, the carbonic anhydrase activity of Picris divaricata, a Zn/ Cd hyperaccumulated plant, was significantly positively correlated with the content of cadmium in the shoots (Ying et al. 2010).

3.5.6 Response of Carbonic Anhydrase to Plant Hormones There are few reports on the effect of plant hormones on carbonic anhydrase activity. In mustard plants, 10–6 M indole butyric acid (IAA), gibberellin (GA3 ), kinetin (Kin) and 10–8 M epibrassinolide (HBR) can increase the activity of carbonic anhydrase. The order of the stimulatory effect of the above hormones on carbonic anhydrase is epibrassinolide > gibberellin > indole butyric acid > kinetin. Abscisic acid (ABA) can reduce the activity of carbonic anhydrase (Hayat et al. 2001). Indole butyric acid and chloroauxin (4-Cl-IAA) at 10–10 to 10–6 M can increase the activity of carbonic anhydrase in leaves. Chloroauxin (10–8 M) can maximally promote the activity of carbonic anhydrase (Ali et al. 2008). Low concentrations of 6-benzylaminoadenine and gibberellin can improve the activity of carbonic anhydrase, while high concentrations can inhibit the activity of carbonic anhydrase in Brassica juncea. The optimal concentration of 6-benzylaminoadenine and gibberellin for the activity of carbonic anhydrase is 5 μM. Similarly, naphthalene acetic acid increases the activity of carbonic anhydrase at low concentrations and inhibits its activity at high concentrations. The optimal concentration of naphthalene acetic acid is 0.5 μM (Li 2007). Therefore, plant hormones have dual effects on carbonic anhydrase.

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3.6 Role of Carbonic Anhydrase on Inorganic Carbon Assimilation in Plants 3.6.1 Unique Thylakoid Carbonic Anhydrase Versus Photosynthetic Oxygen Evolution 3.6.1.1

Intrinsic and Extrinsic CA in PS II-Membranes

There are two types of carbonic anhydrase in chloroplasts: soluble CA in the stroma and thylakoid CA, which is tightly bound to thylakoid membranes (tCA) (Pronina et al. 2002; Rudenko et al. 2006, 2007). Various studies on the character of tCA have demonstrated that photosystem (PS) II membranes of plants, such as Chlamydomonas reinhardtii, Arthrospira maxima, Thermosynechococcus elongatus, spinach, pea, wheat and maize, have two CAs, intrinsic CA, which is tightly associated with the core PS II complex, and extrinsic CA, which can be removed by washing PS II membrane fragments with 1 M CaCl2 (Dai et al. 2001; Villarejo et al. 2002; Moskvin et al. 2004; Ignatova et al. 2006; Rudenko et al. 2006, 2007; Shitov et al. 2009). The intrinsic CA and the extrinsic CA of PS II exhibit significant differences in characteristics, as shown in Table 3.6, in accordance with the available data. From Table 3.6, we find that the intrinsic CA, the extrinsic CA in PS II, and the soluble CA all have very different characteristics, and they all perform very distinct functions. To date, similar to the PS II core complex, the intrinsic CA of PS II has not yet been separated. It seems that PS II is a multienzyme complex with the functions of photosynthetic oxygen evolution and carbonic anhydrase.

3.6.1.2

PS II Did Have CA Activity

There is much evidence that PS II has CA activity according to previous studies on tCA from Arabidopsis, pea, wheat, and maize chloroplasts (Stemler 1986; Moskvin et al. 1995; Khristin et al. 2004; Ignatova et al. 2011). In fact, for the last 20 years, it has been widely accepted that PS II functions as a carbonic anhydrase. Photosynthetic electron transfer in PS II was inhibited by acetazolamide, and the inhibition caused by acetazolamide was completely reversed by the addition of bicarbonate (Shitov et al. 2011). Meanwhile, the photoinduced yield of chlorophyll fluorescence was suppressed by acetazolamide and imidazole (Pronina et al. 2002). PS II inhibitors such as hydroxylamine and 3-(3,4-dichlorophenyl)-1,1’-dimethylurea (DCMU) both inhibited tCA activity (Rudenko et al. 2015). Strong light inhibited both tCA activity and electron transport in PS II, causing photoinhibition, which was prevented by the PS II modifiers atrazine and hydroxylamine (Stemler 1986; Kyle et al. 1984). Formate, an anionic inhibitor, inhibited tCA activity similarly, and other anionic inhibitors, such as I− and bicarbonate, inhibited both PSII and tCA activity (Stemler

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Table 3.6 Comparison of the intrinsic and extrinsic CA characteristics in PS II-membranes Characteristics

Materials

Extrinsic CA

Intrinsic CA

Reference

Position in PS II

Pea

Near PS II in the vicinity of the OEC, at the lumenal side of thylakoid membrane

PS II core-complex

Khristin et al. 2004; Lu and Stemler 2002; Shitov et al. 2009;

Apparent molecular mass

Pea

33 kD, 50 kD, 24 kD, 18 kD

?

Lu and Stemler 2002; Rudenko et al. 2006; Shitov et al. 2009

Direction of reaction

Maize

dehydration

hydration

Lu and Stemler 2007

Effect of divalent cations

Pea

Zn2+ inhibited, and Mn2+ , stimulated the activity

ND

Shitov et al. 2009

Effect of pH

Maize

The highest dehydration activity at pH below 6, and immeasurable above 6.5

Hydration activity insensitive to pH

Lu and Stemler 2007

Effect of Cl−

Maize, Pea

Maximum of activity at 5 ~ 20 mM, afterwards declined, no activity at 80 mM

Continue to increase the activity up to at least 400 mM

Lu and Stemler 2007

SA inhibitors action

Pea, A. thaliana

AZ (at 10–8 -10–5 M) stimulated the activity, high sensitivity to EZ with I50 = 10–9 M

High sensitivity Ignatova et al. to EZ with I50 2006; Shitov et al. = 10–9 M 2009, 2011

Effect of Triton X-100

Pea

No detected effect

Maximum of Pronina et al. activity at triton/ 2002; Khristin Chl ratio of 1.0 et al. 2004

Site of bicarbonate-bound

Spinach

None

?

Tikhonov et al. 2018

Abbreviations: AZ, acetazolamide; A. thaliana, Arabidopsis thaliana; EZ, ethoxyzolamide; OEC, oxygen-evolving center; PS = 2 \* ROMAN II, Photosystem = 2 \* ROMAN II; SA, Sulfonamide; ND, not detected

1980, 1986). Zn2+ inhibits both PS II photosynthetic and tCA activities (Tripathy and Mohanty 1980; Rashid et al. 1991; Stemler 1997). Cl− , Mn2+ and Ca2+ had similar effects on tCA as they did on PS II activities (Stemler 1986, 1997; Lu and Stemler 2007). The tCA activity was sensitive to the surrounding redox potential and was similar to that of PSII, making it unique among known CA types (Bearden and Malkin 1973; Moubarak-Milad and Stemler 1994). Far-red light also stimulates the Hill reaction of PS II and tCA activity (Govindjee et al. 1960; Stemler 1997). Furthermore, it was discovered that an antibody produced against Chlamydomonas reinhardtii thylakoid

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lumen CA(Cah3) reacts with a protein in enriched PS II membranes (Lu and Stemler 2002). The similarities between the above tCA and PS II seem to also imply that PS II is a multienzyme complex with oxygen evolution and CA functions.

3.6.1.3

Mn is Necessary for Oxygen Evolution in the PS II Multienzyme Complex

The characteristics and functions of PS II’s intrinsic CA are obviously different from those of zinc (cadmium)-containing CA. Some inhibitors have a strong inhibitory effect on CA (including intrinsic and extrinsic CA) in PS II but have little effect on photosynthetic activity (Karacan et al. 2014). Although some inhibitors can inhibit both CA activity and photosynthetic activity in PS II, the degree of inhibition varies. For instance, one of the Cu(II)-phenyl sulfonylhydrazone complexes can completely inhibit CA activity in PS II, but it decreases photosynthetic activity by only 33.8% (Rodionova et al. 2017). CA Mn-substituted active-site zinc (CA[Mn]) can act as a peroxidase and produce oxygen in the presence of bicarbonate and hydrogen peroxide (Okrasa and Kazlauskas 2006). In PS II, the activity of oxygen evolution was suppressed after removing Mn clusters, but the intrinsic CA activity remained unchanged, demonstrating that photosynthetic oxygen evolution is not directly correlated with CA activity but is dependent on the presence of Mn clusters (Dai et al. 2001). Mn clusters (Mn4 CaO5 ), in which O atoms act as oxo bridges linking Mn atoms, catalyze dioxygen formation to yield oxygen and are involved in PS II photosynthetic O2 release (Umena et al. 2011).

3.6.1.4

Hydration and Dehydration of Thylakoid CA Depends on pH

A previous study demonstrated that the extrinsic CA near the oxygen-evolving center in PS II had dehydration activity that was sensitive to pH, with the highest dehydration activity at pH below 6, but the intrinsic CA in PS II had hydration activity that was insensitive to pH. Furthermore, the hydration of intrinsic CA is 5 times greater than the dehydration of extrinsic CA in PS II (Lu and Stemler 2007). This suggests that under normal physiological conditions, the hydration of intrinsic CA provides bicarbonate, and the dehydration of extrinsic CA provides protons for photosynthetic oxygen evolution. The combination of extrinsic CA and the PS II core complex provides two basic substrates, bicarbonate and protons, for bicarbonate photolysis (Wu 2021a b, 2023).

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3.6.1.5

79

Photosynthetic Oxygen Evolution

The most direct effect of CA on inorganic carbon assimilation is manifested in the role of thylakoid CA in chloroplasts in photosynthetic oxygen evolution. Under the action of thylakoid CA and photosystem II core-complex (CA-like), the reaction of bicarbonate photolysis occurs in plants: H2 O + CO2 → H+ + HCO− 3 → 1/2 O2 + 2e− + 2H+ + CO2 , which drives photosynthetic oxygen evolution from water (water photolysis), provides CO2 for ribulose-1,5-bisphosphate carboxylase/ oxygenase (Rubisco), elevates and concentrates the CO2 concentration at Rubisco, and accelerates the absorption and reduction of inorganic carbon. Meanwhile, bicarbonate photolysis and water photolysis equally contribute to the total photosynthetic oxygen evolution, which accurately controls photosynthetic oxygen evolution and CO2 assimilation according to the stoichiometric relationship of 1:1 (mol/ mol) (Wu 2023). In addition, carbonic anhydrase accelerates photosynthetic electron transport and transfer (Shitov et al. 2011), depletes the light-generated proton accumulated in the thylakoid under light, thus facilitating the production of CO2 from HCO3 − (Moskvin et al. 2000), and protects photosystem II from photoinhibition under conditions of high illumination (Villarejo et al. 2002).

3.6.2 Direct Effect of Carbonic Anhydrase on Photosynthetic Inorganic Carbon Assimilation 3.6.2.1

Facilitating the Efficient Supply of Carbon Sources and Carbon Fixation Efficiency

Carbonic anhydrase influences photosynthetic inorganic carbon assimilation of plants by facilitating the efficient supply of carbon sources and carbon fixation efficiency. Leaf stomata are the main passages of inorganic carbon in plants. The main pathway for plants to absorb CO2 from the atmosphere is free diffusion. CO2 enters the leaf cell space through stomata and reaches the carboxylation site through the mesophyll cell wall, and the transport process is mainly subject to resistance from the stomata and mesophyll cells (Mooney 1972). Carbonic anhydrase can regulate the opening of stomata to influence the supply of carbon dioxide (Hu et al. 2010, 2015). The effectiveness of CO2 at the carboxylation site is greatly increased under the synergistic effect of CA with ribulose-1,5-diphosphate carboxylase/oxygenase (Rubisco) (Graham and Reed 1971; Sharwood et al. 2016). In C3 plants, carbonic anhydrase accounts for up to 2% of total leaf protein, while 95% of total CA activity is in the chloroplast matrix (Okabe et al. 1984; Tsuzuki et al. 1985); therefore, on the one hand, CA catalyzes the CO2 formed by rapid dehydration of HCO3 − stored in plants and more conveniently diffuses through the cytoplasmic membrane and chloroplast membrane, providing a potential substrate for the carboxylation of Rubisco. On the other hand, CA converts CO2 into HCO3 − ,

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which increases the efficiency of the active transport of inorganic carbon (Moroney et al. 2001; Tiwari et al. 2005). Carbonic anhydrase can be used as a “pump” to regulate the supply of inorganic carbon to Rubiso, which affects photorespiration and the assimilation of photosynthetic inorganic carbon (Igamberdiev and Roussel 2012). CA may also be involved in the transport of carbon dioxide from mitochondria to chloroplasts during photorespiration, facilitating photosynthetic reuse of carbon dioxide produced by respiration or photorespiration (Braun and Zabaleta 2007; Zabaleta et al. 2012; FloryszakWieczorek and Arasimowicz-Jelonek 2017), thereby increasing photosynthetic efficiency, reducing photorespiration, and enhancing the antioxidant capacity of plants (Soto et al. 2015). In C4 plants, both the mesophyll and vascular sheath chloroplasts are involved in CO2 assimilation. The vast majority of CA activity is localized in the cytoplasm of mesophyll cells, which catalyzes the CO2 hydration reaction to form HCO3 − , which provides a substrate for phosphoenolpyruvate carboxylase (PEPC) of C4 and CAM plants (Hatch and Burnell 1990).

3.6.2.2

Facilitating Bicarbonate Use by Plants

Plants may not only use CO2 from the atmosphere for photosynthesis but also use the stored HCO3 for photosynthesis through the action of CA (Wu and Xing 2012; Wu et al. 2018). Carbon dioxide in the atmosphere can freely enter mesophyll cells and be fixed into organic carbon in chloroplasts. Negatively charged bicarbonate ions, which need corresponding carrier proteins or are catalytically converted into CO2 by CA, enter mesophyll cells and then generate organic carbon through a series of biochemical reactions. Under adversity, CA will catalyze the stored bicarbonate ions to generate CO2 and H2 O to compensate for the shortage of photosynthetic substrates when the supply of carbon dioxide in cells is insufficient. It has been reported that βCA1 in chloroplasts may not participate in the process of CO2 transport to Rubisco, and the decrease in chloroplast CA activity has little effect on the photosynthetic CO2 assimilation capacity of plants (Price et al. 1994; Fabre et al. 2007). CA of the cell membrane may form free CO2 by catalyzing bicarbonate ions originating from the soil solution and then transports it to chloroplasts for photosynthesis through CA in the cytoplasm or is catalyzed into bicarbonate ions again for other reactions or storage (Price et al. 1998). The higher the CA activity of plants is, the stronger their ability to utilize HCO3 − (Xing and Wu 2012). Because CA catalyzes the reversible reaction between bicarbonate ions and CO2 very quickly, bicarbonate ions or CO2 in tissues/cells accumulate in a short time, which affects the types and proportions of different inorganic carbons used by plants in this process. Due to the functional diversity and regional distribution of carbonic anhydrase, the fast reversible reaction of bicarbonate ions and CO2 catalyzed by CA not only affects the share of exogenous inorganic carbon source utilization by plants but also affects the total photosynthetic carbon assimilation capacity of plants (Raven 1990; Xing and Wu 2012).

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Karst-adaptable plants generally have a strong ability to use HCO3 − and can alternately use HCO3 − and CO2 with changes in the environment (Wu et al. 2011). For karst-adaptable plants, the CA activity in leaves increases, which on the one hand leads to the reduction or closure of stomatal conductance, reduces transpiration to prevent further dehydration of plants, and on the other hand converts HCO3 − in cells into water and CO2 to cope with the shortage of water and CO2 during photosynthetic carbon assimilation caused by the reduction or closure of stomatal conductance under karst habitats such as karst drought (Wu et al. 2005; Hu et al. 2010). The concentration of bicarbonate in karst soils is too high, which will lead to a high pH value in soil and hinder the absorption of some nutrients, such as iron and zinc, by plants, resulting in the lack of these nutrients by plants, which seriously affects the growth and development of plants (Wallihan 1961; Yang et al. 1994; Misra et al. 2016). The utilization of HCO3 − by plants can increase the utilization of inorganic carbon, reduce the concentration of HCO3 − in the rhizosphere environment, and reduce the pH value of soil, which in turn can further promote the assimilation of plant inorganic carbon.

3.6.3 Indirect Effect of Carbonic Anhydrase on Photosynthetic Inorganic Carbon Assimilation 3.6.3.1

Improving Protective Capacity

Carbonic anhydrase improves chloroplast photoprotection by promoting the utilization of bicarbonate ions and indirectly increases the photosynthesis of plants. Sodium bicarbonate at 0.5 ~ 3 mM can significantly increase the number of leaves, increase dry and fresh weights, and reduce cell death. CAs can act as a switch between Rubisco and phosphoenolpyruvate carboxylase (PEPC), regulate the inorganic carbon pool, stimulate chloroplast photoprotection mechanisms, such as nonphotochemical quenching and an increase in protective cytochrome, and ultimately affect photosynthetic efficiency and crop productivity (D˛abrowska-Bronk et al. 2016).

3.6.3.2

Facilitating the Infiltration of Small Molecules into Macromolecules

The infiltration of inorganic carbon or organic small molecules into organic macromolecules also strongly affects carbon metabolism in plants. Carbonic anhydrase of plants has a significant effect on the infiltration of inorganic carbon. In C4 plants, CA and PEPC have similar distributions in plants, so they use HCO3 − instead of CO2 to promote carboxylation reactions in neutral or slightly alkaline environments (Tiwari et al. 2005). The synergistic effect of CA and PEPC can continuously provide HCO3 − to the carboxylation site (Rathnam and Das 1975), increase the efficiency of

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carboxylation of phosphoenolpyruvate to form oxaloacetic acid catalyzed by PEPC (Del Prete et al. 2016a), accelerate the penetration of inorganic carbon into succinic acid (Park et al. 2017), and promote the assimilation of inorganic carbon. In plants of crassulacean acid metabolism (CAM), carbonic anhydrase also promotes the penetration of inorganic carbon into malic acid (Holtum et al. 1984) and even accelerates the penetration of carbon dioxide into formic acid (Wang et al. 2015). In the synthesis of fat, carbonic anhydrase has also been found to play a promoting role in the penetration of acetic acid into the fat chain (Hoang and Chapman 2002). Carbonic anhydrase promotes the infiltration of inorganic carbon or organic small molecules into organic macromolecules, drives the relevant carbon cycle, and strongly affects the relevant carbon metabolism process.

3.6.3.3

Facilitating the Formation of Photosynthetic Carbon Sinks and Karst Carbon Sinks

The transformation of inorganic carbon into the precipitation of calcium carbonate also strongly affects the carbon metabolism of plants. Carbonic anhydrase significantly promotes the conversion of carbon dioxide to calcium carbonate in organisms (Kim et al. 2012) and accelerates the capture of carbon dioxide by potassium carbonate solution (Zhang and Lu 2015; Hu et al. 2017). The formation of calcium carbonate from many biogenic sources is controlled by carbonic anhydrase in biogenic organisms (Uchikawa and Zeebe 2012; Müller et al. 2014). The deposition of calcium carbonate changes the balance of inorganic carbon in solution and ultimately leads to changes in the photosynthetic carbon sink and karst carbon sink (Xie and Wu 2014, 2017).

3.6.3.4

Regulating the Water Relations of Plants

Water is an important component of plants, participating in various physiological activities of plants, and an important ecological factor affecting plant morphology, structure, growth and development (Dumais and Forterre 2011). Transpiration is a main power of water absorption and transportation of plants, and transpiration is mainly affected by stomatal opening (Müller et al. 2014). When plants suffer from adversity stress, CA is stimulated, and its activity increases, leading to changes in stomatal opening, which eventually inhibits transpiration and reduces water loss (Hu et al. 2010; Xing and Wu 2012). In response to adversity stress, plants increase the efficiency of inorganic carbon acquisition by improving water use efficiency (WUE), that is, to improve the acquisition ability of inorganic carbon under the same unit water loss (Yang et al. 2016) (See chapter 4). At the same time, CA catalyzes the conversion of HCO3 − stored in plants into H2 O and CO2 , providing potential water sources for itself. When the water in plants tends to be balanced and the situation improves, stomata slowly open, transpiration gradually increases, and water consumption rises (Xing and Wu 2012; Perez-Martin et al. 2014).

3.6 Role of Carbonic Anhydrase on Inorganic Carbon Assimilation in Plants

3.6.3.5

83

Regulating Nutrition of Plants

Nutrients necessary for the normal growth and development of plants are divided into macroelements and microelements. Various nutrients have different physiological functions in the metabolism of plants, which are equally important and irreplaceable (Taiz and Zeiger 2006). Nutrient ions in the soil solution reach the rhizosphere through mass flow (water potential gradient) or diffusion (nutrient concentration gradient) and are absorbed by root cells (Chen and Tang 2007). Changes in any environmental factors around the root system (pH, water status, nutrient status, etc.) can affect or even inhibit the absorption of nutrients by plants. The reaction catalyzed by carbonic anhydrase involves two key ions, namely, bicarbonate ions and hydrogen ions. Cations such as K+ , Na+ , Ca2+ , and Mg2+ required for plant growth can easily enter plants through exchange with hydrogen ions. Anions such as PO4 2− and NO3 − can also be rapidly utilized by plants through ion exchange with bicarbonate. Carbonic anhydrase inhibitors can reduce the production of protons and bicarbonate ions in Hydrodictyon reticulatum, thereby affecting ion absorption and reducing photosynthesis (Rybova and Slavikova 1974). In addition, carbonic anhydrase releases hydroxide ions during the dehydration of bicarbonate, which is conducive to inhibiting environmental acidification. In the hydration process of CO2 , H+ is released to inhibit the pH of the environment from being too high. A stable pH environment guarantees nutrients absorption and effective utilization by plants. The utilization of HCO3 − by plants under the action of CA can effectively regulate the rhizosphere environment of plants and promote the absorption of nutrients by plants (Wu et al. 2011). CA can promote the activity of mitochondria by quickly removing inorganic carbon. On the one hand, it can regulate respiration, and on the other hand, it can provide energy for the active transport of ions in plants and promote the active transport of nutrient ions by plants (Henry 1996). CA can also create a favorable environment for the protonation of NH3 and promote the diffusion and transport of nitrogen in plants (Kalloniati et al. 2009). In addition, the synergistic action of CA and PEPC can affect the synthesis of amino acids such as aspartic acid, glycine and serine (Shi et al. 2015; Dimario et al. 2016), and amino acids can also couple the carbon/nitrogen cycle by activating carbonic anhydrase to improve the nitrogen utilization rate (Ghiasi et al. 2017).

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3.7 Possible Significance of CA on the Origin and Evolution of Life The earliest carbonic anhydrase on Earth appeared 3000 to 4000 million years ago (Ma) (Hewett-Emmett and Tashian 1996; Tripp et al. 2004), which is basically the same age as the origin of life (Knoll and Nowak 2017). This ancient carbonic anhydrase belongs to the γ-CA class, in which the metal ion of the active center is Fe (II) (Hewett-Emmett and Tashian 1996; Tripp et al. 2004). It is generally accepted from biological and geological records that oxygenic photosynthetic organisms appeared approximately 2500 Ma (Des Marais 2000; Dismukes et al. 2001; Fischer 2008; Knoll and Nowak 2017). However, the oldest phototrophy (photosynthetic microbial mats) is found in the earlier 3416 Ma (Tice and Lowe 2004, 2006), and the oldest photoferrotrophy is found in 3700 to 3800 Ma (Czaja et al. 2013). It is obvious that oxygen in the biosphere appeared approximately 1000 Ma earlier than the original oxygenic photosynthetic cyanobacteria (Des Marais 2000; Fischer et al. 2016). The γ-CA Mn-substituted active-site iron has the functions of hydration, dehydration and oxygen evolution, which can be used to explain the above fact. High levels of Mn2+ have been found in both shallow water and deep water in Archean basins (Fischer et al. 2015). The iron in the active center of γ-CA is replaced by manganese, forming Mn-substituted CA (CA[Mn]). On the one hand, because the pH of the early Archean is between ~ 6.5 and 7.0 (Halevy and Bachan 2017), the hydration of carbon dioxide in seawater produces very little bicarbonate; however, CA[Mn] accelerates this hydration process and increases the concentration of bicarbonate. On the other hand, CA[Mn] can also catalyze the photolysis of bicarbonate and water, produce protons and electrons, concentrate carbon dioxide, and release oxygen, which not only facilitates the reduction of carbon dioxide but also provides oxygen for the biosphere, accelerating the evolution of life. Approximately 1000 Ma before the occurrence of the first oxygenic photosynthetic organism, cyanobacteria, organic macromolecules that released oxygen may be similar to γ-CA. The hydration and oxygen evolution catalyzed by CA[Mn] are more likely to occur in carbonate rocks. Therefore, we can see the presence of oxygen in the stromatolitic carbonates, but no microfossils such as cyanobacteria are found before 2500 Ma. It seems that CA [Mn] is probably the precursor of the ancestral PSII, which requires more synthesis of chemical, geological and biological evidence.

3.8 Conclusion Carbonic anhydrase can rapidly catalyze the reversible conversion between CO2 and HCO3 − . To date, at least nine CA genetic families have been identified, designated α-, β-, γ-, δ-, ζ-, ε-, η-, θ-, and ι-CA. Carbonic anhydrase has structural and functional diversity. Different organisms, different tissues and organs, different organelles, and

References

85

Fig. 3.1 Inorganic carbon utilization regulated by plasticity and diversity of carbonic anhydrase in plants. Note IC, inorganic carbon; CA, carbonic anhydrase

even carbonic anhydrases of the same organelle have different structures due to their different amino acid sequences and different metal cofactors. The diversity of carbon metabolic pathways, such as carbon migration and conversion, results from the extensive diversity of CAs. Wide differences in the activation or inhibition effects of organic and inorganic substances on different carbonic anhydrase isoenzymes resulted in each well-organized carbon metabolism pathway under normal physiological conditions. Carbonic anhydrase not only provides great convenience for inorganic carbon assimilation in plants but is also an essential key enzyme for inorganic carbon assimilation. The high plasticity of carbonic anhydrase makes the utilization of inorganic carbon in plants highly adaptable to the environment. In the process of adapting to the environment, plants can carry out efficient inorganic carbon assimilation by regulating the diversity and plasticity of carbonic anhydrase (Fig. 3.1). Mn-substituted CA is probably the precursor of the ancestral PSII.

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Chapter 4

Bidirectional Isotope Tracing Culture Technology and Bicarbonate Use by Plants

Abstract Isotope technology is an important means to trace the source of substances. Isotope technology plays an important role in the study of the process and mechanism of photosynthetic oxygen evolution and photosynthetic assimilation. This chapter introduces the process by which scientists discovered several photosynthetic pathways using 14 C labels. The experiment of 18 O tracing photosynthetic oxygen evolution is summarized, the photosynthetic oxygen evolution only from water was debated and questioned, and it was concluded that the oxygen released by photosynthesis comes not only from water photolysis but also from bicarbonate photolysis. A reasonable explanation of the Dole effect is given according to the photosynthetic oxygen evolution equally from bicarbonate and water. Meanwhile, this chapter also explained why the δ18 O of cellulose in tree rings can trace the paleoclimatic environment, although it has little relationship with the source water. Due to the strong isotopic exchange and fractionation, it is difficult to quantitatively distinguish the utilization of root-derived bicarbonate and carbon dioxide from the atmosphere by traditional isotope techniques. This chapter focuses on the principle and technical essentials of bidirectional isotope tracing culture technology. The inorganic carbon sources and utilization pathways of microalgae were described, and the determination methods of direct carbon sinks and indirect carbon sinks of plants were proposed. This chapter describes in detail the measurement of the rate of bicarbonate uptake by plants, the average stable carbon isotopic composition of atmospheric carbon dioxide, the ability of root-derived bicarbonate use by plants and the total photosynthetic capacity by means of bidirectional isotope tracing culture technology. Finally, this chapter also introduced the method of quantitative determination of bicarbonate utilization capacity of wild plants and discussed the metabolic diversity characteristics of karst-adaptable plants. Quantitative determination of the root-derived bicarbonate use capacity of plants provides a scientific basis for the scientific assessment of productivity in karst areas, accurate measurement of karst carbon sinks, and screening of karst-adaptable plants. Keywords Isotope technology · Bidirectional isotope tracing culture · Inorganic carbon assimilation · Inorganic carbon utilization pathway · Dole effect · Indirect carbon sink · Root-derived bicarbonate

© The Author(s), under exclusive license to Springer Nature Singapore Pte Ltd. 2023 Y. Wu and S. Rao, Root-Derived Bicarbonate Assimilation in Plants, https://doi.org/10.1007/978-981-99-4125-4_4

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4.1 Introduction The basic characteristic of plants is photosynthesis. The process by which green plants use the light energy of the sun to assimilate inorganic carbon and water to produce organic substances and release oxygen is called photosynthesis. It mainly includes two stages of light reaction and dark reaction, involving light absorption, electron transfer, photophosphorylation, carbon assimilation and other important reaction steps. It is of great significance to realize the energy conversion of nature and maintain the carbon dioxide and oxygen balance of the atmosphere. Isotope technology can better trace the source of material. Isotopic techniques have made outstanding contributions to the study of photosynthesis. It plays an important role in research on the CO2 fixation pathway, inorganic carbon absorption, photosynthetic oxygen evolution and photosynthetic metabolic pathway. Carbon and oxygen isotopes are commonly used in photosynthesis research. There are 15 known carbon isotopes, including 8 C to 22 C. Carbon isotopes (8 C ~ 19 C) with mass numbers of 8–19 have been found, of which 12 C and 13 C are natural stable isotopes, 14 C are natural radioisotopes, and the rest are synthetic radioisotopes. 12 C accounts for 98.93% of natural carbon, and 13 C accounts for 1.07% of natural carbon. 14 C has the longest lifespan, with a half-life of 5,730 years. The carbon isotope with the shortest lifespan is 8 C, with a half-life of 1.98739 × 10−21 s. Seventeen kinds of oxygen isotopes are known, including 12 O to 28 O, of which 16 O, 17 O and 18 O are stable. The relative abundances in nature are 99.756%, 0.039% and 0.205%, respectively. Other known isotopes are radioactive, and their lifespans are all less than three minutes. Radiocarbon is the most widely used isotope in photosynthesis research. Today, we can clearly understand that the path of carbon dioxide fixation in plants and the photosynthetic types of different plants are involved in the use of 14 C tracing. Although the conclusion that photosynthetically evolved oxygen originated from water is deeply questioned, 18 O also plays an important role in the study of photosynthetic oxygen evolution (Wu 2023). In addition, measuring the stable carbon isotope composition of plants can also enable us to distinguish the photosynthetic pathways between CAM, C3 , and C4 plants. However, both the description of the pathways of carbon dioxide fixation and the process of photosynthetic oxygen evolution, as well as the differentiation of photosynthetic pathways, are qualitative. However, plants can use not only CO2 but also bicarbonate. The capacity to utilize atmospheric CO2 can be measured with a photosynthesis measurement instrument, while the capacity to utilize HCO− 3 is not directly measured with an instrument at present. As inorganic carbon such as CO2 and HCO− 3 is constantly transformed mutually and differential isotope fractionation occurs when plants use CO2 and HCO− 3, these conversions and fractionations are obviously different under different environmental conditions, so it is difficult to obtain the proportion of HCO− 3 used by plants quantitatively with a single isotope tracer (even 14 C). In this chapter, we introduce in detail the technical system for the quantitative utilization of inorganic carbon in plants, which is based on bidirectional isotope tracing culture technology.

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4.2 Radioactive Isotopic Tracing Technology in Photosynthesis Research 4.2.1

11 C as a Tracer to Study CO

2

Fixation in Photosynthesis

Samuel Ruben (1913–1943), Martin Kamen (1913–2002), and their colleagues were the first to apply isotope technology to the study of photosynthesis. First, they used radioactive carbon, 11 C, as a tracer to study the pathway of photosynthetic carbon assimilation. Much work was done between 1939 and 1940 (Ruben et al. 1939a, b, 1940a; Ruben and Kamen 1940a, b). For example, they used 11 C as a tracer to study the CO2 assimilation of barley and confirmed that the fixation of CO2 does not depend on the presence of light. However, after the leaves were placed in the dark for two and one-half to three hours prior to the administration of 11 CO2 , radioactive carbohydrates could not be detected in the absence of light (Ruben et al. 1939a). In addition, they measured the variation in 11 CO2 uptake with time and confirmed that the reduction in radioactivity was mainly due to the assimilation of CO2 rather than the exchange of isotopes. Photosynthetic inhibitors such as hydrogen cyanide, phenylurethan and ultraviolet light can partially or completely block photosynthesis but have no effect on respiration. The dark assimilation of CO2 independent of chlorophyll concentration is reversible (Ruben et al. 1940a). They used 11 C as an indicator to study the CO2 assimilation of Chlorella pyrenoidosa and found that the molecular weight of the radioactive molecules formed by Chlorella pyrenoidosa in the light and dark is approximately four times that of sucrose (Ruben et al. 1940b). The experiments by them used 11 C as a tracer and reached at least two conclusions. One is the conversion of carbon dioxide to carboxyl on a large molecule, not the formation of an additional compound between carbon dioxide and chlorophyll during CO2 fixation by plants. The other is that the subsequent photochemical reduction does not produce formaldehyde (Ruben and Kamen 1940b). Ultimately, their 11 C labeling experiments confirmed that photosynthesis consists of a series of complex chemical reactions. Photosynthesis can be divided into at least four processes: (1) photochemical reactions involving chlorophyll, (2) carbon dioxide fixation into organic carboxyl compounds, (3) organic carboxyl compounds reduction and (4) molecular oxygen evolution (Ruben 1943). Obviously, human beings are not satisfied with the above superficial and preliminary understanding from the short half-life 11 C (20 min) experiments of photosynthesis. People would like to know how the organic matter is converted during the fixation of CO2 , what is the receptor for carbon dioxide, what are the intermediates and so on. The fixation products would have been difficult to identify within 2–5 h owing to short-lived radioactive 11 C. It is impossible to solve these problems without considerable chemical transformations and separations. The 14 C discovered by Samuel Ruben and Martin Kamen shows the dawn for solving the above problems (Ruben and Kamen 1940a, b, 1941).

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4.2.2

14 C

4.2.2.1

14

Tracing of the Pathways of CO2 Fixation in Photosynthesis C Tracing of the Pathways of CO2 Fixation in the Calvin–Benson Cycle

Samuel Ruben and Martin Kamen would like to use the long-lived 14 C to conduct in-depth and detailed research on the structure and properties of carboxyl compounds during the fixation of 14 CO2 . Unfortunately, Samuel Ruben died young in an accident in 1943, and Martin Kamen also encountered political troubles. Therefore, they could not use the 14 C they “invented” to further study CO2 fixation in photosynthesis. However, luckily in misfortune, Melvin Calvin (1911–1997) and Andrew Benson took over the work. From 1947 to 1954, Melvin Calvin, Andrew Benson and James Bassham and coworkers carried out a series of experiments to study the dark fixation of carbon dioxide in Chlorella pyrenoidosa, Scenedesmus D-3 and barley by the exposure of the plants to 14 CO2 following per-illumination and published 22 papers titled “The path of carbon in photosynthesis” (Benson and Calvin 1947, 1950a, b; Calvin and Benson 1948, 1949; Benson et al. 1949, 1950; Benson 1951; Bassham et al. 1954; Wilson and Calvin 1955; Bassham and Calvin 1960; Calvin 1962, Bassham and Calvin 1962). Using extraction, precipitation, and ion exchange column procedures to separate 14 CO2 fixation products, they found that phosphoglyceric acid was the major product in their experiments with the shortest duration (5 s) and is believed to be the first stable product of CO2 reduction. Combining two-dimensional paper chromatography and radioautography, they separated and identified a large number of compounds of carbon assimilation, including phosphoglyoeric acid, carboxylic acids, triose phosphates, amino acids, hexose, ribulose diphosphate (now called ribulose bisphosphate) and sedoheptulose monophosphate, during short periods of photosynthesis with 14 CO2 . According to the chronological order of the occurrence of these compounds, they concluded that ribulose bisphosphate was the acceptor of CO2 and that there was a cycle to regenerate the acceptor. Ultimately, they arranged approximately twelve intermediate compounds in some sequence to form the photosynthetic carbon reduction cycle involving many enzymatic reactions, which was called the Calvin cycle. However, in recognition of the pioneering contributions of Andrew Benson and James Bassham, the photosynthetic carbon reduction cycle was also called the Calvin–Benson cycle or Calvin–Benson–Bassham cycle.

4.2.2.2

14

C Tracing of the C4 -dicarboxylic Acid Pathway of Photosynthesis in C4 Plant Species

However, not all plant species have the Calvin cycle in photosynthetic carbon reduction. After exposure of sugarcane leaves and soybean leaves to 14 CO2 for 15 s,

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Kortschak et al. detected that the radioactivity in 3-phosphoglyceric acid (PGA) accounted for 80% of the total radioactivity in soybean and less than 34% of the total radioactivity in sugarcane. They found that the first stable compounds formed in photosynthesis in sugarcane are malic and aspartic acids (Kortschak et al. 1965). Kortschak et al.’s work is just the beginning. Much work on new pathways of photosynthetic carbon dioxide fixation in sugar cane was completed by Hatch and Slack. After brief exposure to 14 CO2 in sugar-cane and several species of Gramineae, Hatch and Slack found that most fixed radioactivity was located in oxaloacetate, malate and aspartate. The labeling pattern in hexoses was consistent with their formation from 3-phosphoglycerate (Hatch and Slack 1966, 1968, Hatch et al. 1967). Meanwhile, they found that phosphopyruvate carboxylase was apparently the major photosynthetic carbon dioxide-fixing enzyme, and its activity was approximately 60 times greater in sugar cane, sorghum and maize than in oat, wheat and silver beet. Phosphopyruvate carboxylase, but ribulose diphosphate carboxylase activity (now called Rubsico) was only less than one-tenth in sugar-cane, sorghum and maize than in oat, wheat and silver-beet (Slack and Hatch 1967). They also found the enzyme in leaves of sugar cane, sorghum and maize that catalyzes the reversible conversion of pyruvate into phosphopyruvate, not in leaves of extracts from oat, wheat and silver beet (Hatch and Slack 1968). Species in which the C4 -dicarboxylic acid pathway of photosynthesis occurs, such as sugar canes, sorghum and maize, possess a special structure in leaf anatomy and two morphologically distinct types of chloroplasts around the vascular bundle, the inner layer cells near the vascular bundle are called parenchyma sheath cells, and the outer layer cells around the sheath cells are mesophyll cells. The mesophyll cells contain chloroplasts with a developed grana structure, while the chloroplasts of the parenchyma sheath have few grana but contain numerous starch grains. They investigated the distribution of photosynthetic enzymes, 14 CO2 incorporation and intramolecular labeling of intermediates in mesophyll and parenchyma-sheath chloroplasts. From these investigations, they concluded that phosphoenolpyruvate in mesophyll cells combines with CO2 to form malic acid or aspartic acid through the action of phosphopyruvate carboxylase. These C4 -dicarboxylic acids are transferred to parenchyma sheath cells and release CO2 through the action of decarboxylase. The latter enters the Calvin cycle through the action of ribulose diphosphate (RuBP) carboxylase in parenchyma sheath cells (Slack et al. 1969; Johnson and Hatch 1969; Hatch 1971). The above metabolic pathway, which forms C4 -dicarboxylic acids from PEP and then decarboxylates to release CO2 , is called the C4 -pathway or Hatch-Slack pathway (Hatch 1971, 2002). Similar photosynthetic pathways were also found in plants that perform crassulacean acid metabolism (CAM), which is called the CAM pathway (Ranson and Thomas 1960; Black and Osmond 2003).

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14 C

Tracing of the Route of Root-Derived Bicarbonate

Similarly, at first, 11 C with short-lived radioactive was used to trace the carbon of root-derived bicarbonate. For example, Overstreet et al. used radioactive 11 C to study the absorption of bicarbonate ions by barley plants and found that more than 95% of all radioactive carbon was in the form of reduced organic carbon in plants, and only 3–4% of radioactive carbon was in the form of free HCO− 3 . The detailed composition, transportation and allocation of these carbons are difficult to determine because of the short life of 11 C (Overstreet et al. 1940). Using long-lived 14 C to trace the absorption and fixation of bicarbonate by plants can help people understand more detailed footprints of root-derived bicarbonate and the fate of the fixation products. In the 14 C-uptake experiments, Stolwijk and Thimann found that the products of root-derived inorganic carbon fixation can be converted to malic and citric acids and sugars, shifting them into shoots approximately 3 times as great in the light as in the dark (Stolwijk and Thimann 1957). In 14 C-labeled sodium bicarbonate feeding experiments, the 4 C-label was transferred through the roots to the shoots and leaves both in light and in darkness. After the 6 h feeding period, the 14 C-label had been transferred to the shoots and leaves, and more label was in the form of acid-labile products, whereas most of the label was in acid-stable products during the 48 h feeding period. Incorporation of 14 C in light was twice that in darkness at the end of the 24-h feeding period. Both in darkness and in light, the amount of 14 C increased in all parts of plants with time. The uptake of root-derived inorganic carbon might affect carbon budgeting in willow plants (Vapaavuori and Pelkonen 1985; Vourinen et al. 1989). In 14 C-labeled KHCO3 culture experiments, approximately 39% of the total radioactivity was found in shoots and leaves and 61% in roots. KHCO3 enhanced the level of malic acid by approximately 854%, 150%, and 134% in roots, shoots, and leaves, respectively, in relation to the control. The absorption of root-derived bicarbonate might play an important role as an alternative inorganic carbon source in addition to CO2 from atmospheric air (Bialczyk and Lechowski 1992). In the 14 C-labeled inorganic carbon fixation experiments in the roots, the rates of dark incorporation of inorganic carbon attributable to the activity of phosphoenolpyruvate carboxylase in nitrate-fed maize were one-fifth that in ammonium-fed plants after a 30-min pulse of 14 C. The proportion of 14 C located in the shoots was significantly lower in nitrate-fed plants than in ammonium-fed nitrate-fed plants. Nitrate-fed plants allocated relatively more 14 C into organic acids, while ammoniumfed plants favored the incorporation of 14 C into amino acids. Inorganic carbon fixation in the roots of maize plants provides carbon skeletons for ammonium assimilation (Cramer et al. 1993). Similarly, the shift of root-derived organic acids to the shoot and conversion to carbohydrates in tomato (Lycopersicon esculentum) were found in nitrate-fed plants more than in ammonium-fed plants, 14 C partitioning to amino acid synthesis in ammonium-fed plants more than in nitrate-fed plants. Root-derived

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organic acids may contribute to photosynthetic carbon acquisition through decarboxylation of the organic acids in the shoot and the refixation of released CO2 in leaves (Viktor and Cramer 2005) (also see Fig. 2.6 in Chapter 2).

4.3 Stable Isotopic Tracing Technology in Photosynthesis Research 4.3.1 Isotopic Exchange and Carbon Isotope Discrimination in Photosynthesis 4.3.1.1

Limitation of Radioactive 14 C Tracing

Radioactive 14 C is an isotope contained in the human body that exists in the human body itself. Because the abundance of 14 C in the human body is small, it is generally less harmful to the human body. However, the high dose of radioactive 14 C has a great impact on the human body. It can not only produce external radiation, which has a strong radiation damage effect on various parts of the human body, skin and other tissues. It can also penetrate into the human body through the mouth and skin or be resuspended and enter the body through the respiratory tract, causing internal radiation hazards. The harmful effects are divided into somatic effects (cataracts, radiation sickness, cancer, fetal teratogenesis caused by mother’s exposure to radiation, etc.) and genetic effects (chromosome aberration or gene mutation can be transmitted to offspring due to radiation damage to the gonadal germ cells). Therefore, research on radioisotope 14 C as a tracer in photosynthesis is greatly limited. With the rapid development of mass spectrometer technology, stable isotopes have gradually come to the foreground of studying photosynthesis. In the study of inorganic carbon assimilation, the most commonly used stable isotopes are 13 C and 18 O.

4.3.1.2

Isotopic Fractionation

Isotopic fractionation refers to the phenomenon in which the isotopes of an element are distributed among different substances in different proportions in the process of physical, chemical and biological reactions. It includes isotopic thermodynamic equilibrium fractionation and isotopic kinetic nonequilibrium fractionation. Isotopic thermodynamic equilibrium fractionation refers to the fractionation of light and heavy isotopes in different molecules due to changes in the state, phase state, valence state and chemical bond properties between reactants and products in chemical reactions, which is called equilibrium fractionation, also called the isotope exchange reaction. Kinetic nonequilibrium fractionation refers to the fractionation that deviates from isotope equilibrium and is related to time; that is, the distribution of isotopes between phases varies with time and reaction process. Many isotopic fractionations in nature

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have chemical kinetic properties, such as one-way chemical reaction, evaporation, diffusion and biological processes (photosynthesis, respiration, bacterial reduction).

4.3.1.3

Carbon and Oxygen Isotope Exchange

In organisms, on the one hand, carbon isotope exchange often takes place between carbon dioxide and bicarbonate; on the other hand, oxygen isotope exchange between carbon dioxide, bicarbonate, carbonate and water also occurs. The exchange of carbon isotopes is predominated by the following reaction (4.1), which is independent of the hydration of carbon dioxide. CO2 + OH− ↔ HCO− 3

(4.1)

The above reactions can only occur in alkaline environments but rarely in organisms. When the pH value is less than 8, the reaction rate is very slow even if the above reaction occurs (Mills and Urey 1940; Reid and Urey 1943). Oxygen exchange between bicarbonate ions, carbon dioxide, carbonate and water occurs only through the reversible hydration of carbon dioxide when the pH is less than 8. The reversible hydration of carbon dioxide in natural water is very slow. It is not affected by salt, hydrochloric acid and acetic acid but by temperature and other factors. In addition, buffer solution and catalyst can accelerate the reaction (Mills and Urey 1940; Reid and Urey 1943). However, in alkaline aqueous solutions, the exchange of oxygen between carbon dioxide and carbonate ions is considerably fast (Tu and Silverman 1975). The increase in temperature can accelerate the exchange of oxygen isotopes between bicarbonate ions and water. The rate of exchange at 35°C was almost five times that at 25°C (Halas and Wolacewicz 1982). Carbonic anhydrase is ubiquitous in various organelles of plants (please see Chapter 3). It can catalyze the exchange of oxygen between bicarbonate and water due to its catalysis of bicarbonate dehydration and carbon dioxide hydration (Silverman 1973; Silverman and Tu 1975, 1976). Carbonic anhydrase can catalyze the exchange of oxygen between bicarbonate and water not only at pH less than 8 but also at pH between 8.0 and 9.4 (Silverman 1973; Silverman and Tu 1975). The higher the activity of carbonic anhydrase, the higher the efficiency of oxygen exchange between bicarbonate and water. All carbonic anhydrase isoenzymes have the ability to catalyze the exchange of oxygen between bicarbonate and water (Sultemeyer et al. 1990).

4.3.1.4

Carbon and Oxygen Isotope Fractionation of Plants

Carbon and oxygen isotope fractionation of plants mainly comes from photosynthesis and subsequent metabolism of photosynthetic products. The largest fractionation occurs in the process of photosynthesis, while the fractionation of other metabolic processes is generally small (Park and Epstein 1960). In the process of photosynthesis, water and carbon dioxide are incorporated into plant biomass. The abundance

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of 18 O relative to 16 O released by photosynthesis is higher than that of seawater and lower than that of carbon dioxide in the atmosphere (Metzner 1975; Metzner et al. 1979). The abundance of 13 C relative to 12 C in organic matter in plants is usually less than that in atmospheric carbon dioxide (Park and Epstein 1960; Farquhar et al. 1989). Photosynthesis is an extremely complex process that is affected by many physical, chemical and biological factors (O’leary et al. 1992). The fractionation of stable oxygen and carbon isotopes in photosynthesis is therefore inevitably affected by many factors. The fractionation of photosynthetic oxygen isotopes mainly comes from the exchange of oxygen isotopes at each interface in the H2 O-CO2 system. Photosynthetic carbon isotope fractionation is affected by the kinetic processes of inorganic carbon conversion, migration and diffusion, carboxylation, and organic carbon conversion. These kinetics processes are inevitably affected by environmental and physiological factors. The stable carbon isotope ratio of plant organic material varies among different kinds of species and among plants growing in different environments (Cernusak et al. 2013).

4.3.2

18 O

4.3.2.1

18

Tracing Photosynthetic Oxygen Evolution

O Tracing Photosynthetic Evolution and Bicarbonate Photolysis

In photosynthesis research, the most critical experiments to prove whether photosynthetic oxygen evolution comes from water or inorganic carbon are 18 O tracing experiments (Ruben et al. 1941; Stemler and Radmer 1975; Radmer and Ollinger 1980; Hillier et al. 2006). In these photosynthetic oxygen evolution experiments, Ruben et al. made groundbreaking work in 1941. They added 18 O-labeled water and bicarbonate to the photosynthetic oxygen evolution reaction of Chlorella and found that the isotopic composition of oxygen released by Chlorella was consistent with that of water but not with that of inorganic carbon (Ruben et al. 1941). In 1958, Warburg and Krippahl found that bicarbonate significantly stimulated photosynthetic oxygen evolution (Warburg and Krippahl 1958). Thereafter, 18 O labeling experiments on photosynthetic oxygen evolution used 18 O-labeled bicarbonate to stimulate oxygen evolution and were performed with HCO− 3 -depleted broken chloroplasts lacking carbonic anhydrase. Stemler and Radmer carried out experiments with HCO− 3 -depleted maize chloroplast fragments and found that almost all the oxygen stimulated by bicarbonate was unlabeled (Stemler and Radmer 1975). In addition, there are some classic 18 O-labeled photosynthetic oxygen evolution experiments under short saturating light flashes, such as the experiment from Radmer and Ollinger (1980) and that from Hillier et al. (2006) (Radmer and Ollinger 1980; Hillier et al. 2006). At that time, investigators all concluded that the oxygen evolved by HCO− 3 -depleted chloroplasts (free-carbonic anhydrase) is independent of exogenous HCO− 3 used for activation under short saturating light flashes.

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If there was no isotope exchange, it can be believed that the photosynthetic oxygen evolution truly came from the photolysis of water. However, as mentioned earlier in this chapter, the exchange of oxygen isotopes generally occurs between water, carbon dioxide and bicarbonate. Under the action of carbonic anhydrase and photosystem II core complex-like CA, this exchange is extremely fast. Therefore, the above four classical 18 O-labeled photosynthetic oxygen evolution experiments can be interpreted as oxygen evolved not only from bicarbonate but also from water during photosynthesis. Figure 4.1 shows how bicarbonate photolysis can well explain two classical experiments of photosynthetic oxygen evolution using 18 O labeling under continuous illumination, namely, the work of Ruben and his colleagues in 1941 and the work of Stemler and Radmer in 1975 (from Wu 2023). Figure 4.2 demonstrates how bicarbonate photolysis can well explain two other classical experiments of photosynthetic oxygen evolution using 18 O labeling under flashes, namely, the work of Radmer and Ollinge in 1980 and the work of Hillier and his colleagues in 2006 (from Wu 2023). Generally, the above classical experiments of photosynthetic oxygen evolution using 18 O labeling concluded that not only the oxygen released by photosynthesis comes from water photolysis but also may come from bicarbonate photolysis.

4.3.2.2

The Stoichiometric Relationship Between Bicarbonate and Water Photolysis

Some new knowledge and interdisciplinary evidence can lead us to conclude that photosynthetic oxygen evolution is a joint effort of bicarbonate photolysis and water photolysis working together in a 1:1 stoichiometric ratio (Wu 2023). The synthetic formula of photosynthetic oxygen evolution, H2 O + H+ + HCO− 3 → O2 + 4e− + 4H+ + CO2 , can explain how plants accurately control the stoichiometric relationship between photosynthetic oxygen evolution and CO2 assimilation as 1:1 in the presence of inorganic carbon and why the bicarbonate effect occurred during photosynthetic oxygen evolution. Meanwhile, the synthetic formula of photosynthetic oxygen evolution is helpful to understand sulphid photolysis in anaerobic photosynthetic bacteria during photosynthesis and the convenience of bicarbonate photolysis with low free energy compared with water photolysis with high free energy (Wu 2023).

4.3.2.3

Dole Effect and the Origins of Water and Bicarbonate

Importantly, the concentration of atmospheric oxygen remains basically unchanged on the geological scale (Broecker 1970; Bender et al. 1994; Beerling 1999). Previously, it was generally believed that the sole source of oxygen came from water photolysis by plants, and seawater accounted for the vast majority of water on Earth.

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Fig. 4.1 Two classic experiments of 18 O labeling under continuous illumination were explained using bicarbonate photolysis (from Wu 2023). a CA of Chlorella incredibly quickly catalyzes bicarbonate dehydration and CO2 hydration from the experiment by Ruben et al. The 18 O in HC18 O− 3 exchanges rapidly with common oxygen in water and in bicarbonate. Since the concentration of water is much higher than that of inorganic carbon, the oxygen in bicarbonate exchanges nearly all of the oxygen in water (Ruben et al. 1941). b The effect of photosystem II (like CA) led to the fast exchange of oxygen in COOH, OH, and C = O in Stemler and Radmer’s experiment. Since the amount of solvent water is far greater than the amount of H2 18 O and C18 O2 converted from HC18 O− 3 , the vast majority of bicarbonate used for photolysis is unlabeled (Stemler and Radmer 1975)

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Fig. 4.2 Two classic experiments of 18 O labeling under flash were explained using bicarbonate photolysis (from Wu 2023). a In Radmer and Ollinger’s experiment, the oxygen released by bicarbonate-depleted chloroplasts during a brief saturated light flash is derived from bound bicarbonate. It is also not dependent on exogenous bicarbonate used for activation (Radmer and Ollinger 1980). b In the experiment of Hillier et al., the oxygen in most bicarbonate of photosystem II originates from the solvent water and is not labeled. Furthermore, the vast majority of bicarbonates used in photolysis are unlabeled (Hillier et al. 2006)

Therefore, it was deduced that the content of 18 O in the atmosphere should be consistent with that in seawater. However, contrary to the mentioned idea, the content of 18 O in atmospheric oxygen is significantly higher than that in seawater and lower than that in CO2 of seawater. Atmospheric O2 is nearly 24‰ more enriched in 18 O than seawater, and this enrichment is known as the Dole effect (Dole 1935; Dole and Jenks 1944; Metzner 1975; Luz and Barkan 2011).

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Many scientists have tried to analyze the cause of the Dole effect by studying the process of oxygen isotope fractionation in nature but have not found the maximum contributor of the Dole effect. The transport of oxygen released by photosynthesis has almost no isotopic fractionation (Guy et al. 1993). The photosynthesis-respiration cycle is basically responsible for the Dole effect (Kiddon et al.1993). The global 18 O enrichment of leaf water with respect to ocean water is only 4.4‰ (Farquhar et al. 1993). Even if global canopy transpiration is taken into account, the reduction of terrestrial vegetation on the Dole effect would be very small (0.3–0.4‰) (Bender et al. 1994); the ratio of terrestrial to marine gross primary production from 1.8 to 1.0 did not affect the Dole effect in the mid-Holocene (Beerling 1999). Fractionation of oxygen isotopes by respiration and diffusion in soil was also small, and δ18 O of O2 in soils ranged from −1.6 to 0.06‰. Even if global soil respiration is taken into account, it would reduce the Dole effect by only 1–1.5‰ (Angert and Luz 2001; Angert et al. 2001, 2003). The photochemical isotope exchange between CO2 and O2 in the stratosphere resulted in a decrease in δ18 O of atmospheric O2 by only 0.4‰ (Bender et al. 1994). Overall, even if scientists take all factors (including physical, chemical and biological factors) into account, the global 18 O enrichment in atmospheric O2 with respect to ocean water of the Dole effect would still be 21–24‰ (Bender et al. 1994; Hoffmann et al. 2004; Mader et al. 2017). It can be seen from the above that thus far, scientists have not found the real cause of the Dole effect. We believe that the Dole effect is caused by only considering the photosynthetically oxygen released from water but not considering the photosynthetically oxygen evolved from both bicarbonate and water in a 1:1 (mol/ mol) stoichiometric relationship (Wu 2023). The content of 18 O in the atmosphere is a comprehensive reflection of water photolysis and bicarbonate photolysis. Table 4.1 shows the δ18 O and 18 O contents of different constituents of natural H2 O-CO2 systems (data from Metzner 1975). Thus, we can calculate 18 O enrichment in atmospheric O2 according to the fact that bicarbonate photolysis and water photolysis account for half of the photosynthetic oxygen evolution, respectively, as shown in Table 4.2. The 18 O enrichment in atmospheric O2 , which was calculated according to the photosynthetically derived oxygen evolved from water and bicarbonate, is very close to the observed value. Therefore, it can be seen that the oxygen source in the atmosphere is undoubtedly from water and bicarbonate in a 1:1 (mol/mol) stoichiometric relationship, and the Dole effect does not exist when 18 O enrichment in atmospheric O2 with respect to water and bicarbonate is considered. On the geological time scale, water, carbon and oxygen neutralization occur in nature, which keeps the concentrations of CO2 and oxygen in the atmosphere constant (Broker 1970; Bender et al. 1994; Beerling 1999; Wu and Wu 2022). In the biosphere, water, carbon dioxide and oxygen tend to be in dynamic equilibrium, as shown in Fig. 4.3. It can be seen from Fig. 4.3 shows that many physical, chemical and biological processes involving water and carbon dioxide in the biosphere are “reciprocal processes”. For example, oxygen diffusion from leaves to the atmosphere and oxygen

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Table 4.1 δ18 O and 18 O contents of different constituents of natural H2 O-CO2 systems (data from Metzner 1975) (the data of the maximum oxygen exchange between carbon dioxide and bicarbonate ions were from Reid and Urey (1943), and the maximum exchange coefficient is 1.012)

18 O

content

δ18 O

Oxygen 0.2039

22.06‰

0.2042

23.56‰

0.2041

23.06‰ (calculated)

Ocean water

0.1995

0‰

Fresh water

0.1981

0.70‰

Ocean water

0.2078

41.60‰

Fresh water (ave.)

0.2067

36.09‰

Ocean water

0.2103

54.14‰

Fresh water (ave.)

0.2092

48.62‰

Ocean water

0.2090

47.62‰

Fresh water (ave.)

0.2079

42.11‰

Atmosphere Atmosphere (ave.) Water

HCO− 3 (0-exchange)

HCO− 3

(Maximum exchange)

HCO− 3 (Moderate exchange)

Table 4.2 18 O enrichment in atmospheric O2 calculated using the data from ocean water and fresh water according to that photosynthetically oxygen evolved from water and bicarbonate in a 1:1 (mol/ mol) stoichiometric relationship

HCO− 3 HCO− 3 HCO− 3

Ocean water

Fresh water

(0-exchange)

0.2036

0.2031

(Maximum exchange)

0.2049

0.2044

(Moderate exchange)

0.2043

0.2037

absorption by plants, oxygen production by photosynthesis and oxygen consumption by respiration, carbon dioxide entering plants and release from plants, carbonate dissolution and carbonate sedimentation, hydrolysis reactions and dehydration reactions are all “reciprocal processes”. These “reciprocal processes” held the oxygen isotope composition and the concentration of atmospheric O2 unchanged (Broecker 1970), and oxygen isotope fractionation will not be produced in the biosphere. The Dole effect is caused by an insufficient understanding of oxygen sources.

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Fig. 4.3 Carbon dioxide and oxygen of biosphere in dynamic equilibrium. Note ➀ Carbonate rock dissolution; ➁ carbonate sedimentation; ➂ photosynthetic oxygen evolution; ➃ carbon dioxide assimilation; ➄ dissimilation

4.3.3

18 O

of Cellulose in Tree Rings Tracing Past Temperatures

The δ18 O of cellulose in tree rings can also be used as a “thermometer” to measure past temperatures (Gray and Thompson 1976). Although there is still a significant correlation between the δ18 O of whole wood and the average annual temperature, the correlation is poor. There is no significant correlation between the δ18 O of lignin and the average annual temperature. However, the correlation between the δ18 O of cellulose and the average annual temperature is very high. Therefore, only the oxygen isotope composition of cellulose can trace the paleoenvironmental temperature well (Gray and Thompson 1977). The oxygen isotope composition of aquatic and terrestrial plant cellulose shows a systematic difference. Only the oxygen isotope composition of terrestrial coniferous plant cellulose can be applied to measure past temperatures (Epstein et al. 1977). The stable oxygen isotope composition of precipitation resulted from climatic environmental conditions. The δ18 O of cellulose in tree rings closely reflects the stable oxygen isotope composition of atmospheric precipitation and therefore also reflects the climatic information of plant growth, such as temperature and humidity (Burk and Stuiver 1981). Humidity and air temperature influence the stable oxygen isotope composition of cellulose in tropical ecosystems equally by affecting the leafto-air vapor pressure difference (Kahmen et al. 2011). Therefore, it is necessary to treat different measurements with caution when using the stable carbon isotope composition of coniferous plant cellulose in tree rings to determine the paleoclimatic environment. It can be seen from the above facts that the δ18 O of cellulose in tree rings of coniferous plants has little relationship with source water but is closely related to atmospheric precipitation, which indicates that the oxygen of carbohydrates produced

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Fig. 4.4 Photosynthate affected by atmospheric precipitation. Note ➀ photosynthetic oxygen w generation of metabolic water evolution; ➁ carbon dioxide assimilation; ➂ dissimilation; ◯

by photosynthesis does not come from water but from atmospheric CO2 . The interaction among atmospheric temperature, humidity and atmospheric CO2 is reflected in atmospheric precipitation. The δ18 O of cellulose in tree rings reflects the oxygen isotope exchange between H2 O and CO2 before cellulose was synthesized. Some scientists speculated that the oxygen isotope exchange took place before photosynthesis (Gray and Thompson 1976; Epstein et al. 1977). One author of this book, Yanyou Wu, questioned this view and proposed that the isotope exchange was in the process of photosynthetic oxygen evolution during photosynthesis, as shown in Fig. 4.4. The leaves of coniferous plants have thick cuticles, and the stomata are sparse and sagging, which greatly reduces the transpiration of plants and makes them cold resistant. The net photosynthesis and growth of conifers largely depend on the production of stored carbohydrates in winter. Winter temperature may directly affect the isotopic composition of cellulose produced in the next year. The atmospheric precipitation and atmospheric CO2 transformed the bicarbonate. The bicarbonate and water were split into oxygen, and the released CO2 underwent complete oxygen isotope exchange. The released CO2 is assimilated into cellulose precursors; therefore, cellulose inherits the oxygen isotope information of atmospheric CO2 and precipitation affected by evaporation transpiration (Epstein et al. 1977; Kahmen et al. 2011). This can well explain why cellulose δ18 O has the function of tracing atmospheric temperature and humidity. Cernusak et al. found that the oxygen isotope composition of the lamina leaf water from phloem sap was enriched by 14–22‰ compared with source water (Cernusak et al. 2003). This fact demonstrated that the lamina leaf water from phloem sap is metabolic water modified by photosynthesis (Fig. 4.4). It can be seen from Fig. 4.4 that the oxygen of the photosynthate of cellulose precursors comes from the CO2 released from bicarbonate photolysis, while twothirds of the oxygen of bicarbonate comes from dissolved carbon dioxide and onethird from water. This is very consistent with one of the models proposed by Espstein et al., which is obviously better than the other model in explaining the relationship between hydrogen and oxygen in cellulose from aquatic plants (Epstein et al.

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1977). This exactly proved that the stable oxygen isotope information of cellulose in tree rings is mechanistically determined by the photolysis of bicarbonate and water through photosynthesis.

4.3.4

13 C

4.3.4.1

13

as a Tracer and δ 13 C in Photosynthesis Research

C as a Tracer in Photosynthesis Research

Photosynthesis is the process of converting inorganic carbon into organic carbon. Therefore, 13 C is also an effective tool to trace photosynthesis and carbon metabolism pathways. 13 CO2 is widely used as an important tool to characterize plant photosynthetic types, transfer and allocation of photosynthetic products, and source–sink relationships and to trace plants’ carbon footprint and fate in various carbon metabolism processes. The unique biochemical processes of different photosynthetic types can be better understood from 13 CO2 labeling experiments. Osmond et al. studied the regulation of malic acid metabolism in Crassulacean acid metabolism plants in the dark and light using 13 CO2 labeling and found that both ribulose-1,5-biphosphate carboxylase and phosphoenolpyruvate carboxylase are active after short-term exposure to 13 CO2 in the light (Osmond et al. 1988). Arrivault et al. analyzed carbon flow in C4 photosynthesis to reveal how CO2 is incorporated into four-carbon metabolites in the mesophyll according to 13 CO2 labeling kinetics and found the presence of multiple CO2 -concentrating shuttles (Arrivault et al. 2017). The 13 CO2 labeling experiment can also conveniently and effectively study carbon allocation and carbon transfer. Many 13 C-enriched plant materials can be generated using a 13 CO2 pulse-labeling method (Bromand et al. 2001); therefore, it is very convenient to use this method for 13 CO2 labeling experiments. Simard et al. compared carbon allocation and transfer patterns between Betula papyrifera and Pseudotsuga menziesii using the 13 C pulse-labeling method and found that B. papyrifera fixed more total carbon and allocated a greater proportion to its root system than P. menziesii (Simard et al. 1997). Mannerheim et al. investigated carbon allocation to the root system of the tropical tree Ceiba pentandra using the 13 CO2 pulse-labeling method. They found 13 C in root phloem as early as 2 h after labeling. After 5 days of pulse labeling, 27% of the tracers were absorbed by trees in leaves, 21% were absorbed in other tissues of trees, and 52% were lost (Mannerheim et al. 2020). Aranjuelo et al. studied carbon partitioning and dark respiration in cereals subjected to water stress using 13 C labeling, found that photoassimilates stored can reallocate carbon from shoots to ears prior to anthesis during grain filling, and concluded that plants had more carbon available for grain filling (Aranjuelo et al. 2009). Han et al. studied the partitioning and assimilation processes of CO2 absorbed by roots influenced by rootzone CO2 using 13 C labeling and found that the distributional proportion of 13 C in leaves under 0.2% 13 CO2 was significantly lower than that under 0.5% and 1% CO2 concentrations, but carbon assimilation was reduced under long-term high root-zone

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CO2 (Han et al. 2022). Mildner et al. studied temporal and spatial carbon allocation patterns in mature Picea abies using long-term 13 C labeling and demonstrated that the new carbon fixed first arrives at the fast turnover carbon pools in the canopy, then moves away from the crown downward and mixes with older carbon sources (Mildner et al. 2014). 13 C can be used to trace plant source–sink relationships among different individuals and organs. Volpe et al. used 13 CO2 to trace the carbon autonomy of peach shoots and found that both the branches and the fruiting branches of peach trees are relatively autonomous, but they can also import carbon from adjacent branches due to the source–sink relationship among branches (Volpe et al. 2008). Dethloff et al. compared sucrose utilization in Arabidopsis sink predominantly and source predominantly leaves using in situ labeled 13 C-sucrose and found a great difference in 13 C sucrose utilization between them (Dethloff et al. 2017). Finally, 13 C can be used to trace plants’ carbon footprint and fate in various carbon metabolism processes. Dirks et al. studied 13 C incorporation into sugars, starch, proteins, and protein precursors during photorespiration in soybean leaves using 13 C labeling and demonstrated that all glycine incorporated into proteins in photosynthetically active zones was from the photorespiratory pathway under high-light and low CO2 conditions (Dirks et al. 2012). Ma et al. studied fluxes throughout the photosynthetic metabolism of leaves in Arabidopsis thaliana using 13 CO2 labeling and found that the photorespiration flux increased only from 17 to 28% of net CO2 assimilation, while the carboxylation rate doubled with high light acclimation (Ma et al. 2014). Bloemen et al. studied the fate of xylem-transported 13 C-labeled CO2 in leaves of poplar and found that the uptake rate of 13 C was greater under a high vapor pressure deficit than under a low vapor pressure deficit, the enrichment of 13 C in the petiole and veins was higher than that in other tissues, and 13 C may also be fixed in the mesophyll of nonlabeled leaves (Bloemen et al. 2015). Xu et al. performed a flux analysis to study the metabolic origins of nonphotorespiratory CO2 release during photosynthesis using 13 CO2 labeling and found that oxidation of glucose-6phosphate to pentose phosphate via 6-phosphogluconate can account for the majority of CO2 released by respiration in the light (Xu et al. 2021). 13 C-HCO− 3 is also often used to trace the absorption, utilization and conversion of soluble inorganic carbon by plants. Collos et al. compared the discrepancies between net particulate carbon production and bicarbonate uptake by Alexandrium catenella (Dinophyceae) using 13 C-labeled bicarbonate (Collos et al. 2013). Rao et al. investigated the incorporation of newly fixed bicarbonate into plant organs using NaH13 CO3 as a tracer to feed the roots (Rao et al. 2019). Shevela et al. studied bicarbonatemediated CO2 formation on both sides of the photosystem II using NaH13 CO3 as a tracer (Shevela et al. 2020).

4.3.4.2

δ13 C in Photosynthesis Research

Under physiological conditions, carbon isotope exchange fractionation is small. However, in the process of inorganic carbon assimilation, larger carbon isotopic

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fractionation will occur. In the process of carbon dioxide assimilation, there are two reaction steps, which are rate-limiting steps that lead to major carbon isotope fractionation. The first step is from atmospheric carbon dioxide absorption to photosystem II, and the second step is the carboxylation of carbon dioxide at the carboxylation site. The subsequent conversion of tricarboxylic acid into the final photosynthate was also accompanied by carbon isotope fractionation, but the fractionation effect was small. From this, it can be seen that the concentration of CO2 is obviously the most important factor affecting the fractionation of photosynthetic inorganic carbon, while light intensity is not very important (Park and Epstein 1960). The photosynthetic inorganic carbon fractionation of blue–green algae reaches a minimum when the algae are cultured in an environment with a high concentration of inorganic carbon (at a CO2 concentration of 0.2% in air) (Calder and Parker 1973). The enrichment of 12 C in photosynthetic products is not an inevitable result of photosynthesis of carbon dioxide assimilation (Calder and Parker 1973), which is in conflict with the view that plants preferentially use 12 C from the atmosphere in the process of photosynthetic carbon fixation (Park and Epstein 1960), indicating that the isotope fractionation of plants in the metabolic process is related to the rate of physical and chemical reactions, i.e., Plants with low metabolic rates and slow growth have large carbon isotope fractionation (Pardue et al. 1976). For terrestrial plants, diffusion and carboxylation of carbon dioxide are two ratelimiting steps, and the largest fractionation occurs in these two steps. The carbon isotopic discrimination of dark respiration and photorespiration was small. Farquhar and his collaborators developed a set of theories to explain the stable carbon isotope fractionation of plants, including C3 , C4 and CAM species (Farquhar et al. 1982, 1989; Farquhar 1983). They successfully built a simple relationship between discrimination and the ratio of the intercellular and atmospheric partial pressures of CO2 , which can explain the effect on stable carbon isotope fractionation of various environmental factors via their effects on intercellular partial pressures of CO2 (Farquhar et al. 1982; Farquhar 1983). Therefore, the stable carbon isotope composition of plants can be used to study the photosynthetic functions and photosynthetic types of plants (Farquhar et al. 1989; O’leary et al. 1992; Cernusak et al. 2013). Plants have evolved different photosynthetic types, such as C3 , C4 and CAM plant species, to adapt to different environments. Stable carbon isotope compositions in plants vary significantly with different photosynthetic types (Sternberg and DeNiro 1983). The δ13 C of C3 plant species ranges from −20 to −37‰, with an average of −27‰ (Kohn 2010). C4 plant species, whose δ13 C ranges from −9.2 to −19.3‰ (Hattersley 1982), with an average of −12.5‰ (Cerling et al. 1997), have more 13 C enrichment than C3 plant species. Therefore, the photosynthetic type of plants can be identified by measuring the stable carbon isotope composition of different plants under different environments. The photosynthetic function of terrestrial plants controlled by the diffusion of gaseous CO2 and fixation of carbon dioxide is affected by various environmental factors (Scheidegger et al. 2000). Therefore, measuring the stable carbon isotope composition of plants growing under different environmental gradients can reflect the physiological and ecological response of plants to different environments (Cernusak

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et al. 2013). On the one hand, the humidity and concentration of carbon dioxide in the atmosphere affect the atmospheric CO2 entering the stomata and subsequent carboxylation (Madhavan et al. 1991; Feng and Epstein 1995; Yakir and Sternberg 2000); on the other hand, it is also affected by the climate environment and geographical environment (Tieszen 1991). Therefore, the δ13 C of plants can be used as proxy indicators of crop water use efficiency, atmospheric CO2 concentration and precipitation (Körner et al. 1991; Stewart et al. 1995; Condon et al. 2004; Seibt et al. 2008) and can help to analyze the relationship between plants and geographical and climate environments (Diefendorf et al. 2010).

4.4 Quantitative Determination of the Inorganic Carbon Source and Inorganic Carbon Use Pathways of Microalgae 4.4.1 Quantitative Determination of the Share of Different Carbon Sources Used by Microalgae The stable carbon isotope composition of microalgae can reflect their utilization of different inorganic carbon sources in the environment. Generally, there are two sources of inorganic carbon utilized by microalgae, namely, CO2 from the atmosphere and dissolved inorganic carbon in water. In addition, for each inorganic carbon source, there are two inorganic carbon utilization pathways of microalgae, namely, CO2 use and bicarbonate use (Chen et al. 2009). However, due to the continuous mutual transformation of CO2 , HCO− 3 and other dissolved inorganic carbon in the water and the isotope fractionation when microalgae uses CO2 and HCO− 3 , these conversion amounts and isotope fractionation values are obviously different under different environmental conditions, so it is difficult to obtain the proportion of HCO− 3 used by microalgae quantitatively with single isotope labeling (even 14 C). Therefore, we have developed a bidirectional isotope tracer culture technology to solve the problem of signal interference in the process of inorganic carbon transformation and isotope exchange to analyze and quantify the utilization information of different inorganic carbon sources. Bidirectional isotope tracing culture technology is used to simultaneously culture algae with the same biomass in two kinds of culture medium supplemented with sodium bicarbonate with significantly different stable carbon isotope compositions under the same culture conditions. After a certain period of culture, the stable carbon isotope compositions of the newborn algae cultured in the two kinds of culture medium were obtained. According to the difference in the stable carbon isotope composition of the newly formed algae cultured in two kinds of culture media, information on the utilization of bicarbonate by microalgae was calculated (as shown in Fig. 4.5).

4.4 Quantitative Determination of the Inorganic Carbon Source …

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Fig. 4.5 Schematic diagram of bidirectional isotope tracing culture technology for microalgae

Regardless of how CO2 and HCO− 3 are converted, there are carbon dioxide use and bicarbonate use pathways in the process of inorganic carbon utilization by microalgae, and there is approximately 9‰ carbon isotope fractionation between the two inorganic carbon use pathways (Emrich et al. 1970; Mook et al. 1974; Wu et al. 2012). The stable carbon isotope composition of microalgae can be expressed by the isotope mixture model of two-end members, such as Eqs. (4.2, 4.3). δTA = (1 − fbai )δa + fbai (δa + 90/00)(i = 1, 2)

(4.2)

δTB = (1 − fbbi )δai + fbbi (δai + 90/00)(i = 1, 2)

(4.3)

δTA refers to microalgae’s δ13 C value when microalgae use inorganic carbon from the atmosphere, δTB refers to microalgae’s δ13 C value when microalgae use inorganic carbon from added sodium bicarbonate, and microalgae here possess carbon dioxide use and bicarbonate use pathways. fbai refers to the proportion of bicarbonate use by microalgae via the bicarbonate use pathway, in which bicarbonate originated from the conversion of atmospheric CO2 dissolved in water; (1 − fbai ) refers to the share of CO2 use by microalgae sourced from the atmosphere via the CO2 use pathway; fbbi refers to the proportion of bicarbonate use by microalgae via the bicarbonate use pathway, in which bicarbonate was added to culture media; (1 − fbbi ) refers to the share of CO2 use by microalgae via the CO2 use pathway, in which CO2 is sourced from the conversion of bicarbonate added to culture media. We are based on the following assumption: the share of each inorganic carbon use pathway is the same under the same treatment for the same microalgae when microalgae use inorganic carbon from the atmosphere or the added inorganic carbon. Therefore, fbai = fbbi = fbi . δa refers to microalgae’s δ13 C value when microalgae

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fully use carbon dioxide from the atmosphere and completely perform the CO2 use pathway; (δa + 9‰) refers to microalgae’s δ13 C value when microalgae fully use carbon dioxide from the atmosphere and completely perform the bicarbonate use pathway; δai refers to microalgae’s δ13 C value when microalgae use certain inorganic carbon source added and completely perform the CO2 use pathway; (δai + 9‰) refers to microalgae’s δ13 C value when microalgae use certain inorganic carbon source added and completely perform the bicarbonate use pathway. Microalgae in natural water bodies can all use inorganic carbon sources in the atmosphere and inherent inorganic carbon sources in water bodies. For the use of each inorganic carbon source, there are two kinds of inorganic carbon use pathways, including CO2 use and bicarbonate use pathways. For this reason, we have established the isotope mixture model of two end members as shown in Eq. 4.4. δTi = (1 − fBi )δTA + fBi δTB = (1 − fBi )[(1 − fbi )δa + fbi (δa + 90/00)] + fBi [(1 − fbi )δai + fbi (δai + 90/00)](i = 1, 2)

(4.4)

δTi refers to the δ13 C of microalgae cultured in the culture medium with added sodium bicarbonate with a known δ13 C value, fBi is the share of added inorganic carbon in the total carbon inorganic source used by microalgae, and (1 − fBi ) is the share of atmospheric carbon dioxide in the total inorganic carbon source used by microalgae. For the same species of microalgae cultured under the same conditions, Eqs. (4.4) can be expressed as Eqs. (4.5) and (4.6), respectively, depending on the stable carbon isotope composition of sodium bicarbonate added to the culture medium. δT1 = (1 − fB1 )[(1 − fb1 )δa + fb1 (δa + 90/00)] + fB1 [(1 − fb1 )δa1 + fb1 (δa1 + 90/00)]

(4.5)

δT2 = (1 − fB2 )[(1 − fb2 )δa + fb2 (δa + 90/00)] + fB2 [(1 − fb2 )δa2 + fb2 (δa2 + 90/00)]

(4.6)

In Eqs. (4.5) and (4.6), δT1 and δT2 refer to the δ13 C values of microalgae cultured in the culture medium supplemented with the first and second kind of sodium bicarbonate with known δ13 C values, respectively; fB1 and fB2 refer to the shares of the first and second kind of added sodium bicarbonate to the total carbon source for microalgae utilization, respectively; δa1 and δa2 refer to microalgae’s δ13 C values when microalgae use bicarbonate from the culture medium of the first and second kind of sodium bicarbonate added, respectively, and completely perform the CO2 use pathway; (δa1 + 9‰) and (δa2 + 9‰) refer to microalgae’s δ13 C values when microalgae use bicarbonate from the culture medium of the first and second kind of sodium bicarbonate added, respectively, and completely perform the bicarbonate use pathway; fb1 and fb2 refer to the proportion of the first and second kind of added bicarbonate use by microalgae via the bicarbonate use pathway, respectively.

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The same understanding is based on the following two points: (1) Regardless of what kind of labeled sodium bicarbonate is added, the share of the same species of microalgae using the added bicarbonate to the total carbon source is the same under the same culture conditions. Therefore, we can obtain the equation fB1 = fB2 = fB . (2) Regardless of what kind of labeled sodium bicarbonate is added, the share of the same species of microalgae using the bicarbonate pathway is the same under the same culture conditions. That is, fb1 = fb2 = fb . On this basis, by making the difference and simplification between Eq. (4.5) and Eq. (4.6), we can obtain Eq. (4.7). fB =

δT1 − δT2 δa1 − δa2

(4.7)

In Eqs. (4.7), δa1 and δa2 are difficult to obtain, but δa1 should be the sum of the δ13 C (δC1 ) of the first isotope labeled sodium bicarbonate and carbon isotope fractionation value (△1 ) of microalgae using inorganic carbon. At this time, the microalgae used the added bicarbonate and completely performed the CO2 use pathway. Similarly, δa2 should be the sum of the δ13 C (δC2 ) of the second isotope labeled sodium bicarbonate and carbon isotope fractionation value (△2 ) of microalgae using inorganic carbon. At this time, the microalgae used the added bicarbonate and completely performed the CO2 use pathway. As the culture conditions and growth conditions of microalgae are identical, △1 and △2 are exactly equal, that is, △1 = △2 = △; (δa1 − δa2 ) can be converted into the difference between δC1 and δC2 . Therefore, Eq. (4.7) can be further expressed as shown in Eq. (4.8). fB =

δT1 − δT2 δC1 − δC2

(4.8)

In Eq. (4.8), δC1 and δC2 refer to the δ13 C of the first and second isotope-labeled sodium bicarbonate, respectively. In addition, the stable carbon isotope fractionation (△), when microalgae completely perform the CO2 use pathway to assimilate inorganic carbon, can be obtained by measuring the average stable carbon isotope composition (δair ) of atmospheric carbon dioxide in the culture environment, such as Eqs. (4.9) or (4.10). △ = δT1 − δair − fB δC1 + fB δair

(4.9)

△ = δT2 − δair − fB δC2 + fB δair

(4.10)

We successfully quantified the share of different inorganic carbon sources used and the proportion of different inorganic carbon use pathways of the microalgal species Chlorella pyrenoidosa and Chlamydomonas reinhardtii by applying the bidirectional isotope tracing culture technique (Wu et al. 2015). Both C. pyrenoidosa and C. reinhardtii have considerable use of the added bicarbonate, but its use share is small. These microalgae mainly use dissolved inorganic carbon in the water environment

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via the bicarbonate use pathway. The proportion of the bicarbonate use pathway was found to be above 76% in two microalgal species. In addition, we determined the stable carbon isotope fractionation (△) in the process of CO2 assimilation of C. pyrenoidosa, C. reinhardtii and compound algae collected from the surface water of Hongfeng Lake using the bidirectional isotope tracing culture technique. The △ values of C. pyrenoidosa, C. reinhardtii and the compound algae are 14.8‰, 15.3‰, and 21.7‰, respectively. The proportions of the bicarbonate use pathway calculated by △ were 81.1%, 100% and 97.8% in C. pyrenoidosa, C. reinhardtii and compound algae, respectively, which demonstrated that bicarbonate use is the main pathway of inorganic carbon use by microalgae in karst lakes (Zhao et al. 2016).

4.4.2 Quantify the Share of the Pathways of Different Inorganic Carbon Use by Microalgae Equation (4.4) can be simplified as: δTi = δa + fBi (δai − δa ) + 90/00 fbi (i = 1, 2)

(4.11)

where δai − δa can be converted into the difference between the δ13 C (δCi , δC1 or δC2 ) of inorganic carbon in the culture medium with the addition of isotope-labeled bicarbonate and the δ13 C (δC0 ) of inorganic carbon (only from the atmosphere) in the culture medium without the addition of bicarbonate. On this basis, we establish Eq. (4.12) as follows. δai −δa = δCi − δC0 = Di (i = 1, 2)

(4.12)

Substituting Eq. (4.12) into Eq. (4.11), Eq. (4.11) can be simplified as: δTi = δa + fBi Di + 90/00 fbi (i = 1, 2)

(4.13)

According to a previous study, the pathway of bicarbonate use by microalgae was completely inhibited under the condition of adding a high concentration of sodium bicarbonate (16.0 mM NaHCO3 ) and an inhibitor of extracellular carbonic anhydrase (10.0 mM acetazolamid) (Wu et al. 2012). That is, fbi = 0. The algae’s δ13 C (δa ) was obtained as shown in Eq. (4.14). At this time, the microalgae fully used carbon dioxide from the atmosphere and performed the CO2 use pathway. δa = δTi − fBi Di (i = 1, 2)

(4.14)

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Therefore, we can obtain δa . Next, δa is substituted into Eq. (4.13), and finally, the proportion (fbi ) of the bicarbonate use pathway of microalgae can be calculated as shown in Eq. (4.15): fbi = 1000 (δTi − δa − fBi Di )/9(i = 1, 2)

(4.15)

4.4.3 Quantify the Direct Carbon Sink and Indirect Carbon Sink The process of algae assimilating inorganic carbon from the atmosphere and from the water, regardless of whether the pathway of CO2 use or bicarbonate use by algae, are called the direct carbon sink (CSD ) and indirect carbon sink (CSID ), respectively. The direct carbon sink of algae directly removes carbon dioxide from the atmosphere, while the indirect carbon sink of algae reduces the content of soluble inorganic carbon by removing the inherent inorganic carbon in the water body. According to the dynamic balance between carbon dioxide and bicarbonate in the open system, the indirect carbon sink of algae will result in atmospheric carbon dioxide entering the water body and finally indirectly remove carbon dioxide from the atmosphere. The total carbon sink capacity (CST ) of microalgae is the sum of the direct carbon sink and indirect carbon sink. Therefore, we have established the following models to calculate the carbon sink capacity of microalgae. CSA−ai = (1 − fBi ) × (1 − fbi ) × (P − 1)

(4.16)

CSA−bi = (1 − fBi ) × fbi × (P − 1)

(4.17)

CSB−ai = fBi × (1 − fbi ) × (P − 1)

(4.18)

CSB−bi = fBi × fbi × (P − 1)

(4.19)

CSD = CSA−ai + CSA−bi

(4.20)

CSID = CSB−ai + CSB−bi

(4.21)

CSA−ai : Carbon sink capacity when microalgae use carbon dioxide in the atmosphere and conduct the carbon dioxide use pathway. CSA−bi : Carbon sink capacity when microalgae use carbon dioxide in the atmosphere and conduct the bicarbonate use pathway.

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CSB−ai : Carbon sink capacity when microalgae use added inorganic carbon sources and conduct the carbon dioxide use pathway. CSB−bi : Carbon sink capacity when microalgae use added inorganic carbon sources and conduct the bicarbonate use pathway. fBi : The share of inorganic carbon added use in total inorganic carbon use by microalgae. fbi : The share of the pathway of bicarbonate use by microalgae. P: The proliferation of microalgal biomass before and after treatment. It can be seen from the above models that different inorganic carbon source uses and the share of pathway of inorganic carbon use by microalgae can be successfully obtained by measuring the changes in carbon isotope composition of microalgae cultured in sodium bicarbonate medium with different stable carbon isotope compositions using bidirectional isotope tracing culture technology. Compared with the methods of pH drift, inorganic carbon balance dynamics, isotope fractionation effect and spectral analysis (Rost et al. 2007), bidirectional isotope tracing culture techniques can quantitatively obtain various inorganic carbon sources, inorganic carbon use pathways and carbon sinks of microalgae. We studied the effect of algal extracellular carbonic anhydrase on carbon sinks (including direct and indirect carbon sinks) by applying the bidirectional isotope tracing culture technique and found that extracellular carbonic anhydrase mainly increased the utilization of inorganic carbon from the atmosphere and slightly increased the utilization of bicarbonate in aquatic medium. Microalgae mainly utilize CO2 from the atmosphere, accounting for 92% of the total carbon sequestration. Microalgal depletion of bicarbonate in aquatic media may be involved in “missing carbon sinks” (Wu et al. 2015).

4.5 Quantification of Bicarbonate Utilization by Terrestrial Plants 4.5.1 Bidirectional Isotope Tracing Culture In most cases, because the pH value of the soil solution is slightly acidic, the bicarbonate content in the soil is small, and root-derived bicarbonate absorption and utilization by plants may be ignored. However, in the karst environment, due to long-term karstification, the pH and bicarbonate content in karst soils are high; plants grown in karst soils have evolved a set of characteristics that adapt to growth in karst areas. The most important is the efficient use of bicarbonate characteristics. Therefore, plants growing in karst areas can not only use atmospheric CO2 but also use root-derived bicarbonate for photosynthesis (Wu and Xing 2012; Hang and Wu 2016; Rao and Wu 2017; Wang et al. 2017).

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123

Bicarbonate uptake and use by plants, on the one hand, promotes the dissolution of carbonate, conversely accelerating the formation of bicarbonate and making great contributions to “carbon neutralization” (Wu and Wu 2022); on the other hand, it promotes photosynthesis and carbon and nitrogen metabolism, which is conducive to the growth and development of plants, increases the carbon sink capacity of plants, and provides food and energy for the ecosystem. Therefore, determining plants’ capacity for bicarbonate uptake and use can provide new knowledge for plant carbon metabolism and provide basic data for screening karst-adaptable plants and ultimately provide solutions for “carbon peak” and “carbon neutralization”. The absorption and utilization of atmospheric CO2 by plant leaves is a direct carbon sink, while the utilization of root-derived bicarbonate by plants is an indirect carbon sink. Inorganic carbon in soil solution has a variety of forms, and constant transformation and isotope exchange occur among various inorganic carbon forms. Therefore, conventional stable isotope technology has difficulty distinguishing the inorganic carbon from the soil or from the atmospheric inorganic carbon used by plants. Therefore, we also used bidirectional isotope tracing culture technology to identify the use of different carbon sources by plants. Bidirectional isotope tracer culture technology was used to simultaneously label and culture two identical plants in two kinds of culture medium supplemented with sodium bicarbonate with significantly different stable carbon isotope compositions under the same culture conditions. After a certain period of culture, the stable carbon isotope compositions of the culture medium and (or) the newborn leaves cultured in the two kinds of culture medium were obtained. According to the difference in parallel isotope signals, the signal interference in the process of inorganic carbon transformation and isotope exchange was eliminated. Finally, information on different inorganic carbon sources used by plants can be obtained (as shown in Fig. 4.6).

Fig. 4.6 Schematic diagram of bidirectional isotope tracing culture technology for terrestrial plants

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4.5.2 Quantifying the Daily Average Stable Carbon Isotope Composition of Atmospheric CO2 In the past, the main method to determine the stable carbon isotope composition of atmospheric CO2 was to collect air from the testing environment and to measure δ13 C. This method has difficulty obtaining the stable carbon isotope composition of atmospheric CO2 with regional characteristics due to the complexity of the composition in the air from the testing environment and its variability over time. The δ13 C of atmospheric CO2 at some time points can only be obtained, and these values bring certain measurement error due to the complexity. Therefore, it is of great significance to determine the stable carbon isotope composition of CO2 in the atmosphere that can represent regional characteristics for the study of global change. The daily average stable carbon isotope composition of atmospheric CO2 can be obtained by using the characteristics of plants that can utilize bicarbonate by plants based on bidirectional isotope tracing culture experiments. Plants with consistent growth were cultured in the testing environment. Two kinds of sodium bicarbonate with a large difference in δ13 C labeling were added to the nutrient solution used for plant culture. After a period of culture, the stable carbon isotope composition in the culture solution was determined. Due to the continuous absorption and utilization of bicarbonate by plants, the carbon dioxide from the air is continuously replenished to the nutrient solution, resulting in constant changes in the stable carbon isotope composition in the culture solution. The stable carbon isotope change of the culture medium is described by the isotope mixture model of two-end members. The proportion of exogenous bicarbonate in the culture medium to the total inorganic carbon source in the culture medium is obtained through the model, and the δ13 C of carbon dioxide dissolved from the air in the culture medium is calculated. Then, the daily average stable carbon isotope composition of atmospheric carbon dioxide is obtained. The isotope mixture model of two-end members can be expressed as shown in Eq. (4.22). δi = δCa − fBi δCa + fBi δci

(4.22)

where δi refers to the δ13 C of inorganic carbon in the medium after a certain period of plant culture, δCa refers to the δ13 C of inorganic carbon from atmospheric CO2 dissolved in the culture medium, δci refers to the δ13 C of bicarbonate in the initial culture medium, and fBi refers to the share of exogenous bicarbonate to the total inorganic carbon source in the culture medium after a certain period of plant culture. Obviously, δCa can be obtained when δci , δi and fBi are known. Although δci and δi can be measured practically, we can obtain fBi , which cannot be measured, through bidirectional isotope tracing culture technology. For labeled 1 and labeled 2, Eq. (4.22) can be rewritten as shown in Eqs. (4.23) and (4.24), respectively.

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δ1 = δCa − fB1 δCa + fB1 δC1

(4.23)

δ2 = δCa − fB2 δCa + fB2 δC2

(4.24)

where δ1 and δ2 refer to the δ13 C value of inorganic carbon in the culture medium with the first and second labeled sodium bicarbonate added after a certain time of plant culture, respectively; δC1 and δC2 refer to the δ13 C value of the first and second labeled sodium bicarbonate in the initial culture solution, respectively; and fB1 and fB2 refer to the shares of the first and second labeled sodium bicarbonate to the total inorganic carbon source after a certain time of plant culture, respectively. Comparing Eqs. (4.23) and (4.24), fB1 = fB2 = fB , and Eqs. (4.23) and (4.24) are simultaneously solved to obtain Eq. (4.25). fB =

δ 1 − δ2 δC1 − δC2

(4.25)

The fB value calculated according to Eq. (4.25) is the proportion of exogenous bicarbonate added to the total inorganic carbon source in the culture medium after a certain period of plant culture. The fB value is between 0 and 1. The greater the fB is, the less carbon dioxide from the air enters the culture medium. The less carbon dioxide enters the culture medium, the more difficult it is to accurately determine the δ13 C (δCa ) of atmospheric carbon dioxide entering the culture medium during this period of plant culture. The smaller the fB is, the more carbon dioxide from the air enters the culture medium. The more carbon dioxide enters the culture medium, the easier it is to accurately determine the δ13 C (δCa ) of atmospheric carbon dioxide entering the culture medium during this period of plant culture. Therefore, we chose to cultivate plants that can rapidly use bicarbonate to allow more carbon dioxide from the air to enter the culture medium. Through many experiments, the critical value of fB is determined to be 0.6 during plant culture for 24 h. When fB is less than 0.6, the above data can be brought into Eq. (4.26). δCa =

δ1 δC2 − δC1 δ2 δ1 + δC2 − δC1 δ2

(4.26)

δCa can be converted into the average stable carbon isotope composition of atmospheric carbon dioxide (δAa ). The conversion expression is shown in Eq. (4.27): δAa = δCa + △CO2(air)− HCO3(aq)

(4.27)

In Eq. (4.27), △CO2(air)− HCO3(aq) is the carbon isotope fractionation between bicarbonate (aq) and carbon dioxide (air) under a nonequilibrium state. The formation of bicarbonate from carbon dioxide dissolved in solution will undergo carbon isotope fractionation (Mook et al. 1974). The vigorously growing

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plants can quickly use the exogenous bicarbonate added, and the carbon dioxide from the atmosphere can be quickly replenished into the culture medium, which is similar to the results of the rapid removal of carbon dioxide by Mook et al. (1974). Therefore, △CO2(air)− HCO3(aq) is 1.1‰ (Mook et al., 1974), and the above Eq. (4.27) is δAa = δCa + 1.1‰. As an example of measurement, the daily average stable carbon isotope composition of atmospheric CO2 in the same culture chamber was −14.62‰ and −14.61‰ in the environment for the culture of Orychophragmus violaceus and for the culture of Brassica juncea using the bidirectional stable carbon isotope tracing culture technique, respectively (Hang et al. 2015).

4.5.3 Quantify Bicarbonate Uptake by Plants The absorption of bicarbonate by plant roots is a key step for plants to utilize bicarbonate. Because of the mutual transformation of various inorganic carbon and the stable carbon isotope fractionation of the process, it is difficult to obtain the amount of bicarbonate absorbed by plants by measuring the consumption of bicarbonate or the change in the stable carbon isotope of bicarbonate. However, we can measure the bicarbonate consumed by plants through bidirectional isotope tracing culture technology and then obtain the bicarbonate absorption rate of plants. This experiment was set up in two culture systems: a plant culture system and a blank culture system. The plant culture system was used to culture two identical plants in culture medium supplemented with bicarbonate with stable carbon isotope label 1 and stable carbon isotope label 2. The blank culture system was used to place the culture containers without plants containing the culture medium with bicarbonate with stable carbon isotope label 1 and stable carbon isotope label 2 under the same conditions as the plant culture system. The solution volume and δ13 C of the plant culture system and blank culture system at the same treatment time were determined. The proportion of the remaining labeled bicarbonate in the total inorganic carbon of the culture medium in the two culture systems was obtained. Subsequently, the cumulative consumption of labeled bicarbonate and dissolved bicarbonate from the atmosphere in the two systems was calculated. The cumulative consumption of total bicarbonate in the two systems was further obtained. Experiments have demonstrated that the cumulative consumption of total bicarbonate changes with absorption (standing) time in an excellent linear proportional relationship (Fang and Wu 2022). The linear relationship model between the cumulative consumption of total bicarbonate and absorption (standing) time in the two systems was constructed. Subsequently, the rates of total consumption of bicarbonate in the two systems were obtained. Finally, the rate of bicarbonate uptake by the plants was determined. The fresh weight and dry weight of plants and the fresh weight and dry weight of roots/shoots were measured after culture. The rate of bicarbonate uptake by the plants based on the unit weight was calculated.

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127

The proportion of the remaining labeled bicarbonate in the total inorganic carbon of the culture medium in the two culture systems was obtained according to 4.5.2. In the plant culture system, two processes consume the added bicarbonate. On the one hand, the added bicarbonate is absorbed by the plants; on the other hand, it exchanges with the bicarbonate from the transformation of atmospheric CO2 . After these processes take place for a period of time, the proportion (fB ) of the added exogenous bicarbonate to the total inorganic carbon of the culture medium can be expressed in Eq. (4.25). In the blank culture system, the exchange of the added bicarbonate with the bicarbonate from the transformation of atmospheric CO2 consumes the added bicarbonate. After this process takes place for a period of time, the proportion (f0 ) of the added exogenous bicarbonate to the total inorganic carbon of the culture medium can be expressed by Eq. (4.28). f0 =

δ01 − δ02 δC1 − δC2

(4.28)

where i is the sampling (culture) time. δ01 and δ02 represent the stable carbon isotope composition of the culture medium with bicarbonate label 1 and label 2 added to the blank culture system at the same treatment time, respectively. At a certain sampling (culture) time, the proportion (fBi ) of the added exogenous bicarbonate to the total inorganic carbon of the culture medium in the plant culture system can be expressed as Eq. (4.29). Similarly, at a certain sampling (culture) time, the proportion (f0i ) of the added exogenous bicarbonate to the total inorganic carbon of the culture medium in the blank culture system can be expressed as Eq. (4.30). fBi =

δ1i − δ2i δC1 − δC2

(4.29)

f0i =

δ01i − δ02i δC1 − δC2

(4.30)

where i is the sampling (culture) time. δ1i and δ2i represent the stable carbon isotope composition of the culture medium with bicarbonate label 1 and label 2 added to the plant culture system at a certain sampling time, respectively. Similarly, δ01i and δ02i represent the stable carbon isotope composition of the culture medium with bicarbonate label 1 and label 2 added to the blank culture system at a certain sampling time, respectively. The labeled bicarbonate cumulative consumption (pi ) in the plant culture system at different sampling (culture) times can be expressed as Eq. (4.31). Similarly, the labeled bicarbonate cumulative consumption (mi ) in the blank culture system at different sampling (culture) times can be expressed as Eq. (4.32). pi = (cv0 fB0 − cv1i fBi ) − AC1i

(4.31)

mi = (cv0 f00 − cv0i f0i ) − AC0i

(4.32)

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4 Bidirectional Isotope Tracing Culture Technology and Bicarbonate Use …

where i is the sampling (culture) time. c is the original concentration of added bicarbonate concentration, v1i and v0i are the volume of culture solution in the plant culture system and blank culture system at different sampling (culture) times, respectively, fB0 and f00 are the initial proportion of labeled bicarbonate to the total inorganic carbon of the culture medium in the plant culture system and blank culture system, respectively, and AC1i and AC0i are the cumulative sampling consumption in the plant culture system and blank culture system at different sampling (culture) times, respectively. The cumulative sampling consumption (AC1i ) of the labeled bicarbonate in the plant culture system at different sampling (culture) times can be expressed as Eq. (4.33). Similarly, the cumulative sampling consumption (AC0i−1 ) of labeled bicarbonate in the blank culture system under different sampling (culture) times can be expressed as Eq. (4.34). AC1i = AC1i−1 + cvd fBi

(4.33)

AC0i = AC0i−1 + cvd f0i

(4.34)

where vd is the sampling volume during sampling and analysis, i is the sampling time, AC1i and AC0i are both 0, and AC1i−1 and AC0i−1 are the cumulative sampling consumption of the labeled bicarbonate in the last sampling of the plant culture system and blank culture system, respectively. The cumulative consumption (qi ) of dissolved bicarbonate from air in the plant culture system at different sampling (culture) times can be expressed as Eq. (4.35). Similarly, the cumulative consumption of dissolved bicarbonate from the air (ni ) in the blank culture system under different sampling (culture) times can be expressed as Eq. (4.36). qi = pi

(1 − fBi ) fBi

(4.35)

(1 − f0i ) f0i

(4.36)

ni = mi

The cumulative consumption of total bicarbonate (TBC1 ) in the plant culture system at different sampling (culture) times can be expressed as Eq. (4.37). Similarly, the cumulative consumption of total bicarbonate (TBC0 ) in the blank culture system at different sampling (culture) times can be expressed as (4.38). TBC1 = pi + qi =

pi fBi

(4.37)

TBC0 = mi + ni =

mi f0i

(4.38)

4.5 Quantification of Bicarbonate Utilization by Terrestrial Plants

129

Linear relationship models between TBC1 , TBC0 and absorption (standing) time were established. The slopes K1 and K0 of the models are the consumption rates of total bicarbonate in the plant culture system and blank culture system, respectively. The bicarbonate absorption rate (Vb ) of plants can be expressed as Eq. (4.39). Vb = K1 − K0

(4.39)

Finally, the bicarbonate absorption rate (Vb ) of plants is converted into the bicarbonate uptake rate per unit mass. As an example of measurement, we determined the bicarbonate uptake rate per unit mass of Broussonetia papyrifera (Bp) and Morus alba (Ma), and the results are shown in Table 4.3 (Wu et al. 2021). Table 4.3 shows that the ability of B. papyrifera to absorb and utilize bicarbonate is smaller than that of M. alba, and light can promote the utilization of bicarbonate, which is consistent with the fact that B. papyrifera is a karst-adaptable plant. Bicarbonate is bound to cause damage to plants due to its weak alkalinity if it is absorbed too much and concurrently is not used by plants. Therefore, bicarbonate absorbed by B. papyrifera is not only less than that of M. alba but can also be used as a carbon source for photosynthetic organs, further reducing the damage of bicarbonate to plants. At the same time, light can promote the absorption and utilization of bicarbonate by plants, indicating that the bicarbonate absorbed is transported to shoots and leaves through the xylem along with transpiration flow. On the one hand, B. papyrifera absorbed an appropriate amount and was used in a great amount, which greatly reduced the damage; on the other hand, it is also used by plants as an inorganic carbon source, which is an important manifestation of its karst adaptability. However, M. alba is difficult to grow in karst environments, which is related to its high absorption and low utilization. The adaptability of B. papyrifera to higher bicarbonate may also be related to its proper absorption and great utilization. In addition, another reason for the low absorption of bicarbonate by B. papyrifera is that B. papyrifera will exude (produce) much more organic acid than M. alba under high pH and high concentrations of bicarbonate, reducing the pH in the vascular bundle and resulting in reduced absorption of bicarbonate (Zhao and Wu 2017).

4.5.4 Quantification of Root-Derived Bicarbonate Use and Total Photosynthetic Carbon Assimilation of Plants The capacity of atmospheric CO2 use by plant leaves can be directly measured by a photosynthetic apparatus, but the capacity of bicarbonate utilization by plants has not been directly measured by any instrument at present. The capacity of total photosynthetic carbon assimilation in plants includes the ability of CO2 assimilation and root-derived bicarbonate assimilation. Therefore, to determine the total photosynthetic carbon assimilation capacity of plants, it is necessary to determine the total

130

4 Bidirectional Isotope Tracing Culture Technology and Bicarbonate Use …

Table 4.3 Bicarbonate uptake rate per unit mass (VFW , VRFW , VSFW , VDW , VRDW and VSDW ) (μmol/h.g) of Broussonetia papyrifera (Bp) and Morus alba (Ma) under different treatments (VFW , VRFW , VSFW , VDW , VRDW and VSDW are the bicarbonate uptake rates per unit mass based on plant fresh weight, root fresh weight, shoot fresh weight, plant dry weight, root dry weight and shoot dry weight, respectively) Treatment Sample VFW VDW VRFW VSFW VRDW VSDW No. (μmol/h.g (μmol/h.g (μmol/h.g (μmol/h.g (μmol/h.g (μmol/h.g FW) DW) FW) FW) DW) DW) 1 Bp 12-12/L-D 2 (control) 3 Mean Ma 1 12-12/L-D 2 (control) 3 Bp 24 L

Bp 24 D

Ma 24 L

16.79

14.69

6.80

62.23

23.00

13.69

10.68

5.40

42.21

20.25

4.77

17.90

12.46

7.72

52.37

27.20

4.34

16.13

12.61

6.64

52.27

23.48

6.16

24.83

24.32

8.26

125.17

30.97

5.18

19.49

18.37

7.22

88.24

25.01

5.27

21.11

21.56

6.97

117.28

25.74

Mean

5.54

21.81

21.42

7.48

110.23

27.24

1

1.64

5.90

3.05

3.53

13.24

10.64

2

2.45

8.30

5.05

4.77

19.97

14.21

3

2.48

7.70

4.83

5.10

17.88

13.52

Mean

2.19

7.3

4.31

4.47

17.03

12.79

1

0.97

3.42

2.08

1.80

8.23

5.84

2

0.47

1.74

0.89

0.99

3.73

3.25

3

1.30

4.46

2.67

2.53

9.82

8.16

Mean

0.91

3.21

1.88

1.77

7.26

5.75

1

7.89

36.06

29.37

10.79

174.19

45.47

2

7.18

32.77

26.88

9.80

163.77

40.97

3

7.86

35.69

30.51

10.58

183.33

44.31

Mean Ma 24 D

4.65 3.59

7.64

34.84

28.92

10.39

173.76

43.58

1

−0.04

−0.18

−0.12

−0.07

−0.61

−0.26

2

1.19

4.78

3.63

1.77

16.22

6.79

3

2.26

9.52

6.18

3.56

31.35

13.66

Mean

1.14

4.71

3.23

1.75

15.65

6.73

Note 12-12/L-D, 12-h photoperiod (control); 24 L, 24 h of continuous light; 24 D, 24 h of continuous dark

bicarbonate utilization capacity of plants. The total share of bicarbonate utilization by plants is the sum of the share of the added bicarbonate utilization and the share of the atmospheric CO2 -derived dissolved bicarbonate use. The principle and procedure of bidirectional isotope tracing culture technology to determine the share of the added bicarbonate use by terrestrial plants is similar to that by microalgae, such as 4.4.1. Here, we describe the principle and process according to the actual situation of terrestrial plants.

4.5 Quantification of Bicarbonate Utilization by Terrestrial Plants

131

Bidirectional isotope tracer culture is similar to 4.5.1. The stable carbon isotope composition of the new leaves met the isotopic mixing model of two end members, as shown in Eq. (4.40). δT = δA − fB δA + fB δB

(4.40)

For labeled 1 and labeled 2, Eq. (4.40) can be rewritten as shown in Eqs. (4.41) and (4.42), respectively. δT1 = δA − fB1 δA + fB1 δB1

(4.41)

δT2 = δA − fB2 δA + fB2 δB2

(4.42)

In Eqs. (4.41) and (4.42), δT1 and δT2 refer to the foliar δ13 C values of plants cultured in the culture medium supplemented with the first and second kind of sodium bicarbonate, respectively; fB1 and fB2 refer to the shares of the added bicarbonate use to total carbon source use by plants in the labeled 1 and the labeled 2 culture medium, respectively; δA refers to the foliar δ13 C values assuming that plants completely use CO2 as the sole carbon source; δB1 and δB2 refer to the foliar δ13 C values assuming that plants completely use the added bicarbonate as the sole carbon source of the labeled 1 and the labeled 2 culture medium, respectively. In Eqs. (4.41) and (4.42), fB = fB1 = fB2 , and Eqs. (4.41) and (4.42) were simultaneously solved in Eq. (4.43). fB =

δT1 − δT2 δB1 − δB2

(4.43)

It can be seen from 4.4.1 above that in Eq. (4.43), δB1 – δB2 can be converted into the difference between the δ13 C of labeled 1 bicarbonate (δC1 ) and that of labeled 2 bicarbonate (δC2 ). Equation (4.43) can be further expressed as shown in Eq. (4.44). fB =

δT1 − δT2 δC1 − δC2

(4.44)

Therefore, the share (fB ) of exogenous bicarbonate use to total carbon source use by plants can be obtained by the substitution of δC1 , δC2 , δT1 and δT2 into Eq. (4.44). The above calculated fB refers to the share of the exogenous bicarbonate use to total carbon source use by the first expanded (fast-growing) leaf tested of the plant, and the first expanded leaf at this time includes “new carbon” and “old carbon” tissues. In fact, the “old carbon” tissue of this leaf has not used the bicarbonate added exogenously, and only the “new carbon” tissue is affected by the assimilation of the added exogenous bicarbonate. Therefore, the proportion (fBN ) of net increased organic carbon of plants using exogenous bicarbonate can be expressed as shown in Eq. (4.45).

132

4 Bidirectional Isotope Tracing Culture Technology and Bicarbonate Use …

fBN =

fB fLA

(4.45)

where fLA is the increased proportion of organic carbon in the first expanded leaf tested during a certain period of culture. The increased proportion of organic carbon in the first expanded leaf tested is expressed by the increased proportion of leaf area. In addition, CO2 in the atmosphere will also be dissolved into the culture medium to form bicarbonate, which is used by plants. Therefore, to measure the share of total bicarbonate use by plants, it is also necessary to consider the share of utilization of bicarbonate formed by dissolved atmospheric CO2 in the culture medium of plants. Suppose that a certain amount of bicarbonate is initially added to the culture medium. At this time, the added exogenous bicarbonate accounts for 100% of the total inorganic carbon in the culture medium. The generated bicarbonate from the dissolved atmospheric CO2 into the culture medium and the exogenous bicarbonate added is simultaneously absorbed and utilized by plants. According to 4.5.2 and 4.5.3, the proportion of bicarbonate used by plants is proportional to the concentration of bicarbonate in the solution. After a period of plant culture, the proportion of the added exogenous bicarbonate (fBA ) to the total inorganic carbon source in the culture medium can be obtained by referring to 4.5.2 and 4.5.3. The utilization share of the bicarbonate generated from atmospheric CO2 can be calculated according to the fBA after a period of plant culture. The share of the total bicarbonate (including dissolved from the atmosphere and added exogenously) to the total carbon source of plants (fBT ) is the sum of fBN , and the utilization share of the generated bicarbonate from atmospheric CO2 and fBT can be expressed as Eq. (4.46). fBT =

2fBN 1 + fBA

(4.46)

The net photosynthetic rate (PN ) measured by a photosynthetic instrument based on the measurement of atmospheric CO2 flux is the capacity of plants to assimilate atmospheric CO2 . However, plants can also use bicarbonate for photosynthesis. The capacity to assimilate bicarbonate by plants is expressed as BUC, that is, the utilization capacity of bicarbonate, which can be expressed as Eq. (4.47). BUC =

PN fBT 1 − fBT

(4.47)

The total inorganic carbon assimilation capacity (TPN ) of plants can be expressed as Eq. (4.48). TPN =

PN 1 − fBT

(4.48)

4.5 Quantification of Bicarbonate Utilization by Terrestrial Plants

133

As an example, we measured the total photosynthetic capacity of Orychophragmus violaceus and Brassica juncea seedlings when bicarbonate was added to the culture medium at 5, 10 and 15 mM, as shown in Table 4.4. It can be seen from Table 4.4 that, compared with B. juncea, the bicarbonate use ability of O. violaceus increases with the increase in the concentration of bicarbonate added, which shows that O. violaceus is more adaptable to karst environments with high concentrations of bicarbonate than B. juncea (Hang and Wu 2016). As another example, we measured the total photosynthetic capacity of O. violaceus and B. juncea seedlings under different concentrations of polyethylene glycol (6000) (PEG) with 10 mM bicarbonate added to the culture medium, as shown in Table 4.5. Table 4.5 shows that, compared with B. juncea, the share of bicarbonate use by O. violaceus increases with increasing PEG concentration. It can be inferred that O. violaceus is more suitable to grow in the karst arid environment. The total photosynthetic assimilation capacity determined by the above method is limited by the size of the plant. If the plant is too large, the bicarbonate added to the culture medium is quickly absorbed by the plant, the labeled inorganic carbon in the culture medium is quickly diluted by the dissolved inorganic carbon from the atmosphere, the foliar δ13 C of the plant is less affected by the bicarbonate added, and the difference in foliar δ13 C of plants cultured on the two labeled culture media is small, which is covered by the systematic error of sample measurement and makes the determination result unreliable. Therefore, the above methods can only be used to determine the total photosynthetic assimilation capacity of plant seedlings of appropriate size. To determine the total photosynthetic assimilation capacity of relatively large plants, we adopted the following improved methods. The plants were cultured by changing the culture medium once every two days. Taking 24 h as the base point, the added inorganic carbon in the culture medium 24 h prior is a positive increase Table 4.4 The total photosynthetic inorganic carbon assimilation capacity of Orychophragmus violaceus and Brassica juncea under different concentrations of added bicarbonate Species O. violaceus

B. juncea

Parameters

Concentration of bicarbonate added (mM) 5

10

15

PN (μmol CO2 m−2 s−1 )

5.82 ± 0.41

5.57 ± 0.19

4.80 ± 0.27

BUC (μmol CO2 m−2 s−1 )

0.32 ± 0.01

0.85 ± 0.02

1.01 ± 0.04

TPN (μmol CO2 m−2 s−1 )

6.14 ± 0.31

6.42 ± 0.25

5.81 ± 0.15

PN (μmol CO2 m−2 s−1 )

6.56 ± 0.16

7.71 ± 0.25

7.63 ± 0.35

BUC (μmol CO2 m−2 s−1 )

0.22 ± 0.00

0.25 ± 0.01

0.24 ± 0.02

TPN (μmol CO2 m−2 s−1 )

6.78 ± 0.45

7.96 ± 0.56

7.87 ± 0.45

134

4 Bidirectional Isotope Tracing Culture Technology and Bicarbonate Use …

Table 4.5 The total photosynthetic inorganic carbon assimilation capacity of Orychophragmus violaceus and Brassica juncea under different concentrations of PEG Species O. violaceus

C. juncea

Parameters

Concentration of polyethylene glycol (g/L) 0

10

20

40

fBT

6.68 ± 0.10

13.08 ± 0.10

17.53 ± 0.18

47.58 ± 0.20

BUC (μmol CO2 m−2 s−1 )

0.30 ± 0.00

0.35 ± 0.00

0.45 ± 0.03

1.13 ± 0.06

TPN (μmol CO2 m−2 s−1 )

4.50 ± 0.15

2.68 ± 0.15

2.56 ± 0.10

2.37 ± 0.12

fBT

2.89 ± 0.15

7.63 ± 0.05

7.97 ± 0.06

3.18 ± 0.20

BUC (μmol CO2 m−2 s−1 )

0.17 ± 0.00

0.28 ± 0.00

0.18 ± 0.00

0.01 ± 0.01

TPN (μmol CO2 m−2 s−1 )

5.78 ± 0.50

3.69 ± 0.14

2.23 ± 0.23

0.17 ± 0.31

(up relative to the base point), and the added inorganic carbon in the culture medium 24 h later is a negative increase (down relative to the base point). Both the positive increase and the negative increase in the added inorganic carbon are caused by the influence of atmospheric CO2 on the inorganic carbon of the culture medium. The positive increase in the added inorganic carbon was regarded as equal to the negative increase. Therefore, the δ13 C of inorganic carbon in the medium 24 h after changing the culture medium was measured in the middle of the culture time. The share of the total bicarbonate (including dissolved from the atmosphere and added exogenously) to the total carbon source of plants was obtained as Eq. (4.49). fBT =

δT1 − δT2 δCM1 − δCM2

(4.49)

where δCM1 and δCM2 are the δ13 C of inorganic carbon in the medium with labeled 1 and labeled 2 bicarbonate added 24 h after changing the culture medium in the middle of the culture time, respectively. BUC and TPN can also be obtained according to Eqs. (4.47) and (4.48). Using the above improved method, we measured the effect of osmotic stress caused by different concentrations of polyethylene glycol (PEG 6000) (0, 100, 200 g/ L) in a simulated karst environment (pH 8.3, the concentration of NaHCO3 10 mM) on the use of root-derived bicarbonate of Camptotheca acuminata and then obtained the total photosynthetic assimilation rate. The results showed that the root-derived bicarbonate use by C. acuminata accounted for 10.36% of the total inorganic carbon source under the control without PEG and accounted for 20.05% and 16.60% of the total inorganic carbon source under moderate stress (PEG, 100 g/L) and severe stress

4.6 Measurement of Total Inorganic Carbon Assimilation Capacity of Plants …

135

Table 4.6 Inorganic carbon assimilation (PN [μmol CO2 m−2 s−1 ], BUC [μmol CO2 m−2 s−1 ] and TPN [μmol CO2 m−2 s−1 ]) of Camptotheca acuminata under three osmotic stresses PEG-0 (fBT = 10.36%)

PEG-100 (fBT = 20.05%)

PEG-200 (fBT = 16.60%)

PN

BUC

TPN

PN

BUC

TPN

PN

BUC

TPN

1

4.06

0.47

4.53

4.81

1.21

6.02

3.87

0.77

4.64

3

4.37

0.51

4.88

3.80

0.95

4.75

1.87

0.37

2.24

5

3.96

0.46

4.42

3.46

0.87

4.33

1.35

0.27

1.62

7

4.62

0.53

5.15

5.29

1.33

6.62

1.55

0.31

1.86

9

4.24

0.49

4.73

3.82

0.96

4.78

1.60

0.32

1.92

11

4.18

0.48

4.66

2.88

0.72

3.60

1.70

0.34

2.04

13

5.01

0.58

5.59

3.95

0.99

4.94

1.71

0.34

2.05

15

3.86

0.45

4.31

3.03

0.76

3.79

1.20

0.24

1.44

T-Mean

4.29

0.50

4.79

3.88

0.97

4.85

1.86

0.37

2.23

Days

(PEG, 200 g/L), respectively (Rao and Wu 2017). Table 4.6 shows the influence of different concentrations of PEG on the total photosynthetic assimilation rate of C. acuminata. Plants use bicarbonate for photosynthesis, reducing the use of water. The water use efficiency of plants refers to the dry matter produced by the unit weight of water lost by transpiration. The traditional water use efficiency (WUE) of plants is the ratio of the photosynthetic rate and transpiration rate determined by a photosynthetic instrument. Therefore, when we know that plants not only use atmospheric carbon dioxide but also root-derived bicarbonate, the actual water use efficiency (WUEa ) of plants should be the ratio of the total photosynthetic rate and transpiration rate. Table 4.7 shows the effects of different PEG concentrations on the transpiration rate and actual water use efficiency. Table 4.6 and Table 4.7 show that moderate osmotic stress can greatly improve the utilization of bicarbonate. The total photosynthetic rate is not affected by moderate osmotic stress, and the transpiration rate is greatly reduced by osmotic stress. In summary, moderate osmotic stress can greatly improve water use efficiency without reducing overall photosynthesis.

4.6 Measurement of Total Inorganic Carbon Assimilation Capacity of Plants in Field Habitats Under the same conditions, different plant species have not only different abilities to assimilate carbon dioxide but also different abilities to use other forms of inorganic carbon. Under different environments, the same plant species has different abilities to assimilate different inorganic carbon. Although we have developed a set of methods that can measure the total photosynthetic carbon assimilation capacity

136

4 Bidirectional Isotope Tracing Culture Technology and Bicarbonate Use …

Table 4.7 Water metabolism (Tr [transpiration rate, mmolH2 O m−2 s−1 ], WUE [water use efficiency, μmol CO2 mmol−1 H2 O] and WUEa [actual water use efficiency, μmol CO2 mmol−1 H2 O]) of Camptotheca acuminata under three osmotic stresses PEG-0 (fBT = 10.36%)

PEG-100 (fBT = 20.05%)

Days

Tr

WUE

WUEa

Tr

WUE

WUEa

1

1.65

2.5

2.79

1.01

5.15

6.44

PEG-200 (fBT = 16.60%) Tr 0.79

WUE

WUEa

5.45

6.53

3

1.16

3.95

4.41

0.50

7.91

9.89

0.24

7.81

9.36

5

1.34

3.03

3.38

0.51

6.85

8.57

0.18

7.75

9.29

7

1.74

2.74

3.06

0.88

6.16

7.70

0.25

6.07

7.28

9

1.08

4.06

4.53

0.40

9.76

12.21

0.16

10.25

12.29

11

0.69

6.52

7.27

0.32

9.47

11.84

0.18

10.21

12.24

13

1.28

3.93

4.38

0.59

7.11

8.89

0.22

7.65

9.17

15

0.77

5.13

5.72

0.36

8.51

10.64

0.16

7.77

9.32

T-Mean

1.21

3.98

4.44

0.57

7.62

9.53

0.27

7.87

9.44

of plants cultured in the laboratory, these methods have difficulty measuring the total photosynthetic carbon assimilation capacity of plants at different seedling ages and growth stages in different environments in real time. In the karst limestone area with a high concentration of bicarbonate, the photosynthetic apparatus based on the measurement of atmospheric carbon dioxide flux is only used to measure the inorganic carbon assimilation capacity of plants, which seriously underestimates the productivity of plants in the karst area. Therefore, accurate measurement of the total photosynthetic carbon assimilation capacity of plants at different seedling ages and different growth stages under different environments plays an important role in correctly evaluating plant productivity, screening karst-adaptable plant species with high productivity, and using karst-adaptable plants to manage and restore the fragile karst ecological environment. There is a significant difference between the stable carbon isotope fractionation value of plant leaves when fully assimilating carbon dioxide and that of plant leaves when fully assimilating bicarbonate. Fractionation (△ca) of atmospheric carbon dioxide by leaves satisfies Eq. (4.50) during the C3 pathway (Farquhar et al. 1989). △ca = a + (D − a)(Ci/Ca)

(4.50)

In Formula (4.50), Ci is the concentration of intercellular carbon dioxide, Ca is the concentration of carbon dioxide in the atmosphere, “a” is the fractionation of inorganic carbon during stomatal diffusion, with a value of 4.4 ‰, and D is the fractionation of inorganic carbon during Rubsico carboxylation, with a range of Ci , and substitute the 27‰ ~ 29‰ according to different plant species. Let k = Ca above known parameters into Formula (4.50). △ca = 4.40/00 + (D − 4.40/00)k

(4.51)

4.6 Measurement of Total Inorganic Carbon Assimilation Capacity of Plants …

137

Equation (4.51) represents the fractionation of inorganic carbon when C3 plants fully assimilate carbon dioxide from the atmosphere. However, plants not only use carbon dioxide from the atmosphere but also use root-derived bicarbonate. When carbon dioxide is hydrolyzed to bicarbonate ions, the fractionation of inorganic carbon is −9.9‰, and when plants use bicarbonate, there is no diffusion process of 4.4‰ isotope fractionation. Therefore, the fractionation value (△b) of inorganic carbon from root-derived bicarbonate by leaves meets Eq. (4.52). △b = △ca − 14.30/00

(4.52)

Assuming that f is the share of carbon dioxide used by plants from the atmosphere, the actual stable carbon isotope fractionation value (△aa) of inorganic carbon assimilated by plant leaves is △aa, and f and △aa meet the relationship shown in Eq. (4.53). △aa = f △ca + (1 − f)(△ca − 14.30/00)

(4.53)

Equation (4.53) can be rewritten into Eq. (4.54). f=

△aa − △ca + 14.30/00 14.30/00

(4.54)

The intercellular carbon dioxide concentration (Ci) and the carbon dioxide concentration in the atmosphere (Ca) were measured through the photosynthetic apparatus. According to Eqs. (4.51) and (4.52), △ca and △b can be obtained. Then, according to the foliar δ13 C value and Eq. (4.53), f can be obtained. Finally, the total capacity to assimilate inorganic carbon by plants (TPN ) can be calculated as Eq. (4.55). TPN =

Pn f

(4.55)

As an example, we measured the share of root-derived bicarbonate utilization and the total inorganic carbon assimilation of Camptotheca acuminata and Platycarya longipes of karst secondary forest grown on Jiangjun Mountain (26°39' 22'' N, 106°36' 38'' E, 1200 m altitude) in July and August, as shown in Table 4.8 and Table 4.9. Table 4.8 and Table 4.9 show that the inorganic carbon assimilation ability of P. longipes is greater than that of C. acuminata. The photosynthetic capacity and utilization of root-derived bicarbonate of different plants are obviously different. Carbon dioxide assimilation is still the main mechanism for terrestrial plants. As we took the newly developed leaves of each plant for measurement, the growth environment of each leaf was different, and the photosynthetic function and utilization of inorganic carbon were also significantly different, resulting in great differences in foliar δ13 C. In fact, the δ13 C of different parts of the same leaf varies greatly (Nguyen Tu et al. 2013). Therefore, to reduce the workload, we can determine the

138

4 Bidirectional Isotope Tracing Culture Technology and Bicarbonate Use …

Table 4.8 Carbon dioxide assimilation (PN ) and total inorganic carbon assimilation (TPN ) of Camptotheca acuminata (CA) and Platycarya longipes (PL) in karst secondary forest in July Plant No.

PN μmol m−2 s−1

f %

TPN μmol m−2 s−1

Plant No. PN μmol m−2 s−1

f %

TPN μmol m−2 s−1

CA-1

4.51

88.6

5.09

PL-1

8.26

95.8

8.62

CA-2

6.69

88.6

7.55

PL-2

7.15

90.0

7.94

CA-3

5.74

78.3

7.33

PL-3

10.42

90.9

11.46

CA-4

4.98

95.4

5.22

PL-4

8.94

83.5

10.71

CA-5

8.21

87.4

9.39

PL-5

10.87

90.2

12.05

CA-6

5.55

87.9

6.31

PL-6

12.03

81.0

14.85

CA-7

5.46

88.1

6.20

PL-7

9.99

74.1

13.48

CA-8

7.28

88.4

8.24

PL-8

8.27

78.2

10.58

CA-9

6.79

85.6

7.93

PL-9

10.37

89.8

11.55

CA-10

8.18

78.6

10.41

PL-10

12.13

85.7

14.15

CA-11

9.12

85.2

10.70

PL-11

11.36

78.6

14.45

CA-12

7.33

84.5

8.67

PL-12

11.73

78.2

15.00

Mean

6.65

86.4

7.75

Mean

10.13

84.7

12.07

Table 4.9 Carbon dioxide assimilation (PN ) and total inorganic carbon assimilation (TPN ) of Camptotheca acuminata (CA) and Platycarya longipes (PL) in karst secondary forest in August Plant No.

PN μmol m−2 s−1

f %

TPN μmol m−2 s−1

Plant No. PN μmol m−2 s−1

f %

TPN μmol m−2 s−1 12.72

CA-1

5.55

87.0

6.38

PL-1

8.84

69.5

CA-2

7.74

86.6

8.94

PL-2

7.44

94.5

7.87

CA-3

10.90

73.9

14.75

PL-3

4.99

91.1

5.48

CA-4

6.00

91.8

6.54

PL-4

8.97

92.3

9.72

CA-5

5.46

87.6

6.23

PL-5

6.80

88.1

7.72

CA-6

5.64

94.0

6.00

PL-6

4.24

88.5

4.79

CA-7

5.03

73.7

6.82

PL-7

5.87

87.5

6.71

CA-8

3.88

79.9

4.86

PL-8

8.88

83.2

10.67

CA-9

4.73

74.7

6.33

PL-9

9.83

84.9

11.58 12.27

CA-10

4.99

75.0

6.65

PL-10

9.94

81.0

CA-11

2.63

84.3

3.12

PL-11

8.66

97.0

8.93

CA-12

6.69

66.2

10.11

PL-12

6.64

71.3

9.31

Mean

5.77

81.2

6.38

Mean

7.59

85.7

8.98

4.7 Conclusions and Outlook

139

Table 4.10 Water metabolism (Tr [transpiration rate, mmolH2 O m−2 s−1 ], WUE [water use efficiency, μmol CO2 mmol−1 H2 O], WUEa [actual water use efficiency, μmol CO2 mmol−1 H2 O]), and the share of root-derived bicarbonate utilization (fBT , %) of Camptotheca acuminata (CA) and Platycarya longipes (PL) of karst secondary forest in July Plant No.

Tr

fBT

WUE

WUEa

Plant No.

Tr

fBT

WUE

WUEa

CA-1

2.13

11.4

2.13

2.40

PL-1

2.86

13.0

2.89

3.32

CA-2

3.54

11.4

1.89

2.13

PL-2

2.85

13.4

2.54

2.94

CA-3

3.83

21.7

1.50

1.91

PL-3

3.85

26.1

2.74

3.71

CA-4

2.64

4.6

1.88

1.97

PL-4

4.08

8.2

2.21

2.40

CA-5

3.97

12.6

2.07

2.36

PL-5

4.39

12.4

2.47

2.82

CA-6

2.89

12.1

1.92

2.19

PL-6

4.23

6.0

2.85

3.03

CA-7

3.26

11.9

1.68

1.90

PL-7

3.47

26.3

2.88

3.91

CA-8

3.15

11.6

2.31

2.61

PL-8

3.14

20.1

2.64

3.30

CA-9

3.75

14.4

1.83

2.14

PL-9

2.96

25.3

3.50

4.68

CA-10

3.32

21.4

2.48

3.15

PL-10

3.27

25.0

3.71

4.94

CA-11

3.23

14.8

2.86

3.36

PL-11

3.16

15.7

3.59

4.26

CA-12

3.05

15.5

2.41

2.85

PL-12

3.43

33.8

3.42

5.17

Mean

3.23

13.6

2.08

2.41

Mean

3.47

18.8

2.95

3.71

δ13 C of mixed sampling leaves and calculate the share of bicarbonate utilization using the average value of Ci/Ca of plants. In addition, we measured the share of root-derived bicarbonate utilization and the water metabolism of C. acuminata and P. longipes of karst secondary forest grown on Jiangjun Mountain in July and August, as shown in Table 4.10 and Table 4.11. Tables 4.10 and 4.11 show that water metabolism characteristics, similar to photosynthetic assimilation, in different plant species vary, and transpiration and water use efficiency are significantly different among plants in the same species. Individual diversity in the assimilation of inorganic carbon and water metabolism is another specific embodiment of the diversity of metabolic pathways of karst-adaptable plants and a specific embodiment of the biodiversity of karst-adaptable plants (Wu et al. 2018).

4.7 Conclusions and Outlook Plants can use not only carbon dioxide from the atmosphere but also (root-derived) bicarbonate (from water medium). The 18 O and 14 C labels reveal the secrets of photosynthetic oxygen release and carbon dioxide assimilation in plants. However, traditional isotope techniques have difficulty quantitatively revealing the stoichiometric relationship between oxygen release and carbon dioxide assimilation. Bidirectional isotope tracing culture technology uses parallel isotope signals to eliminate signal

140

4 Bidirectional Isotope Tracing Culture Technology and Bicarbonate Use …

Table 4.11 Water metabolism (Tr [transpiration rate, mmolH2 O m−2 s−1 ], WUE [water use efficiency, μmol CO2 mmol−1 H2 O], WUEa [actual water use efficiency, μmol CO2 mmol−1 H2 O]), and the share of root-derived bicarbonate utilization (fBT , %) of Camptotheca acuminata (CA) and Platycarya longipes (PL) of karst secondary forest in August Plant No.

Tr

fBT

WUE

WUEa

Plant No.

Tr

fBT

WUE

WUEa

CA-1

2.57

13.0

2.16

2.48

PL-1

4.35

30.5

2.04

2.94

CA-2

3.42

13.4

2.27

2.62

PL-2

2.35

5.5

3.17

3.35

CA-3

5.16

26.1

2.11

2.86

PL-3

1.62

8.9

3.08

3.39

CA-4

2.43

8.2

2.47

2.69

PL-4

2.64

7.7

3.39

3.68

CA-5

2.70

12.4

2.02

2.31

PL-5

2.34

11.9

2.92

3.32

CA-6

1.85

6.0

3.09

3.29

PL-6

1.93

11.5

2.25

2.54

CA-7

2.40

26.3

2.09

2.84

PL-7

2.32

12.5

2.53

2.89

CA-8

1.70

20.1

2.29

2.86

PL-8

3.49

16.8

2.55

3.07

CA-9

1.90

25.3

2.52

3.37

PL-9

3.73

15.1

2.65

3.12

CA-10

1.62

25.0

3.12

4.16

PL-10

3.04

19.0

3.28

4.05

CA-11

1.00

15.7

2.68

3.18

PL-11

2.44

3.0

3.58

3.69

CA-12

2.97

33.8

2.26

3.41

PL-12

2.72

28.7

2.47

3.47

Mean

2.48

18.8

2.42

3.01

Mean

2.75

14.3

2.83

3.29

interference in the process of inorganic carbon conversion and inorganic carbon isotope exchange and quantifies the information of bicarbonate utilization by plants. The Dole effect occurs because the origin of oxygen in the atmosphere is not only water photolysis but also bicarbonate photolysis. We have successfully quantified the inorganic carbon sources and utilization pathways of microalgae, direct carbon sinks and indirect carbon sinks, root-derived bicarbonate use and the total photosynthetic capacity of plants by using bidirectional isotope tracer culture technology. In the future, we will use bidirectional isotope tracing culture technology of carbon–oxygen dual elements to quantify the stoichiometric relationship between photosynthetic oxygen evolution (water and bicarbonate photolysis) and carbon dioxide assimilation of plants, providing more reliable direct evidence for the carbon neutralization path in nature (Wu and Wu 2022) (Fig. 4.7).

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Fig. 4.7 Carbon neutralization in nature (Wu and Wu 2022). Carbonate rocks (lithosphere) are dissolved under the action of water (hydrosphere) and carbon dioxide (atmosphere) to form Ca2+ and bicarbonate (biosphere). Ca2+ combines with bicarbonate to precipitate calcium carbonate (CaCO3 ) into the lithosphere. Plants split bicarbonate and water to release oxygen and carbon dioxide (biosphere). Plants then assimilate carbon dioxide to form carbohydrates (CH2 O) (biosphere). Finally, organisms utilize oxygen to decompose carbohydrates into carbon dioxide to the atmosphere and water to the hydrosphere

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Silverman DN, Tu CK (1975) Buffer dependence of carbonic anhydrase catalyzed oxygen-18 exchange at equilibrium. J Am Chem Soc 97(8):2263–2269 Silverman DN, Tu CK (1976) Carbonic anhydrase catalyzed hydration studied by 13 C and 18 O labeling of carbon dioxide. J Am Chem Soc 98(4):978–984 Simard SW, Durall DM, Jones MD (1997) Carbon allocation and carbon transfer between Betula papyrifera and Pseudotsuga menziesii seedlings using a 13 C pulse-labeling method. Plant Soil 191(1):41–55 Slack CR, Hatch MD (1967) Comparative studies on the activity of carboxylases and other enzymes in relation to the new pathway of photosynthetic carbon dioxide fixation in tropical grasses. Biochem J 103(3):660–665 Slack CR, Hatch MD, Goodchild DJ (1969) Distribution of enzymes in mesophyll and parenchymasheath chloroplasts of maize leaves in relation to the C4 -dicarboxylic acid pathway of photosynthesis. Biochem J 114(3):489–498 Stemler A, Radmer R (1975) Source of photosynthetic oxygen in bicarbonate-stimulated Hill reaction. Science 190(4213):457–458 Sternberg L, DeNiro MJ (1983) Isotopic composition of cellulose from C3 , C4 , and CAM plants growing near one another. Science 220(4600):947–949 Stewart GR, Turnbull MH, Schmidt S, Erskine PD (1995) 13 C natural abundance in plant communities along a rainfall gradient: a biological integrator of water availability. Funct Plant Biol 22(1):51–55 Stolwijk JAJ, Thimann KV (1957) On the uptake of carbon dioxide and bicarbonate by roots, and its influence on growth. Plant Physiol 32:513–520 Sultemeyer DF, Fock HP, Canvin DT (1990) Mass spectrometric measurement of intracellular carbonic anhydrase activity in high and low Ci cells of Chlamydomonas: studies using 18 O exchange with 13 C/18 O labeled bicarbonate. Plant Physiol 94(3):1250–1257 Tieszen LL (1991) Natural variations in the carbon isotope values of plants: implications for archaeology, ecology, and paleoecology. J Archaeol Sci 18(3):227–248 Tu CK, Silverman DN (1975) Kinetics of the exchange of oxygen between carbon dioxide and carbonate in aqueous solution. J Phys Chem 79(16):1647–1651 Vapaavuori EM, Pelkonen P (1985) HCO− 3 uptake through the roots and its effect on the productivity of willow cuttings. Plant Cell Environ 8:531–544 Viktor A, Cramer MD (2005) The influence of root assimilated inorganic carbon on nitrogen acquisition/assimilation and carbon partitioning. New Phytol 165(1):157–169 Volpe G, Lo Bianco R, Rieger M (2008) Carbon autonomy of peach shoots determined by 13 Cphotoassimilate transport. Tree Physiol 28(12):1805–1812 Vourinen AH, Vapaavuori EM, Lapinjoki S (1989) Time course of uptake of dissolved inorganic carbon through willow roots in light and in darkness. Physiol Plant 77:33–38 Wang R, Wu Y, Xing D, Hang H, Xie X, Yang X, Zhang K, Rao S (2017) Biomass production of three biofuel energy plants’ use of a new carbon resource by carbonic anhydrase in simulated karst soils: Mechanism and capacity. Energies 10:1370 Warburg O, Krippahl G (1958) Hill-reaktionen. Z Naturforsch B 13(8):509–514 Wilson AT, Calvin M (1955) The photosynthetic cycle. CO2 -dependent transients. J Am Chem Soc 77(22): 5948–5957 Wu Y, Wu Y (2022) The Increase in the karstification–photosynthesis coupled carbon sink and its implication for carbon neutrality. Agronomy 12(9):2147 Wu YY, Xing DK (2012) Effect of bicarbonate treatment on photosynthetic assimilation of inorganic carbon in two plant species of Moraceae. Photosynthetica 50:587–594 Wu YY, Xu Y, Li HT, Xing DK (2012) Effect of acetazolamide on stable carbon isotope fractionation in Chlamydomonas reinhardtii and Chlorella vulgaris. Chinese Sci Bul 57(7):786–789 Wu Y, Li H, Xie T (2015) Biogeochemical action of microalgal carbonic anhydrase. Science Press of China, Beijing, China, pp 75–110 Wu Y, Xing D, Hang H, Zhao K (2018) Principles and techniques of determination on plants’ adaptation to karst environment. Science Press of China, Beijing, China, pp 1–88

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Chapter 5

Root-Derived Inorganic Carbon Assimilation by Plants in Karst Environments

Abstract Root-derived inorganic carbon has been demonstrated to contribute to plant carbon gain by many experiments in the laboratory. However, it remains largely unknown whether and to what extent soil dissolved inorganic carbon (DIC) influences leaf photosynthesis in karst environments. In this chapter, we first review the current knowledge regarding the uptake, transport, allocation, and assimilation of DIC in plant organs and revisit several representative reports concerning the bicarbonate assimilation by plants under simulated karst environments. Then we summarize the characteristics of natural karst enviroments and provide theoretical models to quantify the contribution of soil DIC to leaf total photosynthesis in field conditions. Further, isotope evidence for plants’ use of soil DIC in karst habitas and species-specific induced variation in DIC assimilation as well as their spatial–temporal heterogeneity are discussed. Finally, we highlight ecophysiological and biogeochemical significance of DIC assimilation in the karst environments, for instance the possible strategy for plants adapting to karst environments and the impact on the estimation of global carbon budget. Keywords Dissolved inorganic carbon · Root uptake · Transpiration stream · Leaf photosynthesis · Karst · Isotope · Photosynthetic model · Carbon sink

5.1 Introduction Photosynthesis is the most important reaction on earth and is a foundation for the survival of nearly all life. The classical concept of photosynthesis describes a process in which plants utilize the light to catalyze the reaction of H2 O and CO2 , or precisely the coupling of the Hill reaction and dark reaction, and subsequently produce organic matter as well as releasing energy. For a long time, atmospheric CO2 has been regarded as the only substrate for leaf photosynthesis in terrestrial plants. On the contrary, cyanobacteria, microalgae, macroalgae and seagrasses are capable of using HCO− 3 dissolved in water for photosynthesis (Poschenrieder et al. 2018), which is coordinated by the well-known “Carbon Concentration Mechanisms”. Partly inspired

© The Author(s), under exclusive license to Springer Nature Singapore Pte Ltd. 2023 Y. Wu and S. Rao, Root-Derived Bicarbonate Assimilation in Plants, https://doi.org/10.1007/978-981-99-4125-4_5

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by these discoveries, in the past dacedes researchers show much interests in rootderived inorganic carbon involving in photosynthesis of higher plants. Typical experiments are applying 14 C or 13 C labeling in the root zones or detached organs to track the fate of bicarbonate or CO2 in different parts of plants (Enoch and Olesen 1993; Stringer and Kimmerer 1993). Later studies on soil CO2 efflux also demonstrate that a portion of CO2 respired by living roots, mycorrhizal fungal, micro-organisms, and from the decomposition of dead organic matter can be fixed by photosynthetic cells in woody tissues or leaves (Teskey et al. 2008; Bloemen et al. 2013). However, there are some uncertainties associated with above researches. For example, the labeling experiments are usually conducted in hydropinic culture, sometimes exposing to high concentration of bicarbonate and largely hampering leaf photosynthesis, which may not reflect actual situation of plants growing the soils. While in natural conditions, the concentration of soil CO2 , when compared with xylem CO2 , is too low to be absorded by roots in some species (Bloemen et al. 2016). In addition, based on isotope mixing models, most studies predict a very low contribution of root-derived inorganic carbon to leaf photosynthesis, for instance less than 1%. This has led many people to believe that the effect of root-deirved inorganic carbon on plant carbon gain or leaf photosynthesis can be ignored. Nevertheless, a unique scenario has not been explored. In the karst environments, soils are characterized by high pH, high concentrations of bicarbonate and calcium. There is highly possibility for plants to utilize the inorganic carbon from soils. Considering a few limitations of high abundance of 13 C labeling in natural ecosystems, investigation and quantyfication of root-derived inorganic carbon assimilation in karst habitats seem challenging. In this chapter, we will review the current knowledge of the uptake, transport, allocation, and assimilation of dissolved inorganic carbon (DIC) within plant tissues or organs. We will focus on karst environments and introduce a new approach (natural abundance of 13 C tracer) to estimate the contribution of soil DIC to leaf photosynthesis in karst habitats. To test the robustness of this approach, we will revisit the results of recent studies which evaluate the influences of species, spatial and temporal factors on plants’ use of soil DIC.

5.2 How Plants Utilize Soil Dissolved Inorganic Carbon 5.2.1 Bidirectional Carbon Flow at the Soil–Root Interface It has been evident that carbon flow at the soil–root interface is bidirectional (Jones et al. 2009; Brüggemann et al. 2011): carbon lost from roots (e.g., root respiration, exudation of organic compounds, death and lysis of root epidermal cortical cells) and carbon uptake from soils (e.g., absorption of CO2 or dissolved inorganic

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carbon, active root uptake of sugars and organic nitrogen compounds). But for inorganic carbon, especially bicarbonate, the relevant information is limited. In fact, the processes regarding the uptake, transport, and assimilation of HCO− 3 are rather complicated and inconclusive.

5.2.2 Using CO2 and Bicarbonate as Fertilizers: A Brief History CO2 is a nonpolar molecule that can readily pass through the cell membranes of all living organisms. In addition, the membrane transport of CO2 is also shown to be facilitated by aquaporins (water-permeable complexes) in both animals and plants (Uehlein et al. 2003; Kaldenhoff 2012; Wang et al. 2016; Ermakova et al. 2021). Once absorbed by the plants, CO2 diffuses freely within the cells (Miller 1960), whether in leaves, stems or roots (Teskey and McGuire 2007). The availability of CO2 thus forms the foundation of the dark reaction of CO2 in photosynthetic tissues. The concentration of CO2 in ambient air was previously considered one of the major environmental factors influencing leaf photosynthesis and biomass production (Wilson and Calvin 1955; Ehleringer and Björkman 1977). How plants respond to variable CO2 has continued to intrigue many plant physiologists and ecologists for a better understanding of the relationship between plants and the environment (Luo et al. 1999; Nowak et al. 2004). In laboratory (chamber) experiments, leaves were exposed to elevated CO2 concentrations, for example, 1.3–18 times the background value (Kramer 1981; Dietz and Heber 1984). This treatment caused a significant improvement in photosynthesis in the short term, which was mainly attributed to biochemical and anatomical regulation and acclimation at the compound level (Farquhar et al. 1980; Kramer 1981; Dietz and Heber 1984). In the early 1990s, largescale enrichment experiments, free-air CO2 enrichment (FACE) (475–600 parts per million [ppm], several months to ten years), emerged and allowed the study of the responses of plants and ecosystems to rising CO2 concentrations in natural conditions (Ainsworth and Long 2005). The results obtained in numerous FACE studies were consistent with the general trend observed in chamber experiments (Kimball et al. 1995; Ainsworth et al. 2003; Hickler et al. 2008). Model predictions also showed that the CO2 fertilization effect will continue to stimulate plant biomass in the future despite the limitation of soil nutrients (Terrer et al. 2019). In the context of global change, the CO2 concentration of the atmosphere increases from a preindustrial level of approximately 280 ppm to 421 ppm in 2022 (NOAA 2022). This increasing trend causes worldwide concern about the CO2 greenhouse effect on climate variability (Mercer 1978; Rodhe 1990; Schmidt et al. 2010) but also brings opportunities for exploring the sensitivity of vegetation productivity to elevated atmospheric CO2 concentrations at the global scale from long-term observations via remote sensing or eddy covariance techniques (Wang et al. 2020a; Chen et al. 2022).

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Another approach is to irrigate plants with CO2 -enriched solution in the root zones, which also has the potential to increase photosynthesis and biomass production (Arteca et al. 1979; Mauney and Hendrix 1988), as it is analogous to the traditional fertilizers used in agricultural practice. The irrigation method is much easier to implement and has low cost, which dates back to the nineteenth century (Birner and Lucanus 1866; Moll 1877; Pfeffer 1881; Vines 1882). Enoch and Olesen (1993) summarized the results from a large number of previous studies and concluded that the growth of plants irrigated with enriched CO2 water was significantly higher than that of the control, by 2.9% on average. In particular, crops and vegetables usually exhibited a larger enhancement in plant productivity triggered by root-zone CO2 fertilization. For example, Arteca et al. (1979) recorded an approximately 18% increase in dry matter resulting from root uptake of CO2 in potato. Nakayama and Bucks (1980) observed an increase of 10–20% in the productivity of potato, wheat and cantaloupe. A study conducted by Mauney and Hendrix (1988) showed an increase in dry weight by 15–125% in cotton irrigated with CO2 -saturated water. Altogether, the effect of enriched CO2 on plant growth is marked and species-specific. One of the potential mechanisms was initially proposed by Stoklasa (1929), Barbieri (1930), and Miller (1931) in which CO2 was taken up by roots and then transported aboveground, which was subsequently confirmed by 14 C isotopic evidence (Kuzin et al. 1952). Irrigation with CO2 -enriched water is a promising tool to stimulate plant growth; however, this approach is never applied to agricultural activities on a large scale, possibly due to difficulties in the preservation of CO2 in the water and little effect on the production of some economic plants. Uptake of CO2 by roots has an important implication for plants’ use of bicarbonate, allowing for the interconversion between CO2 and HCO− 3 . Although bicarbonate addition was usually observed to hamper leaf photosynthesis and biomass production and induce symptoms of chlorosis (iron deficiency) in many species (Bloom and Inskeep 1986; Alhendawi et al. 1997; Msilini et al. 2009; Covarrubias and Rombolà 2013; Zhao and Wu 2017; Wang et al. 2019b), a few studies have argued that it is likely that high pH (or alkaline conditions) rather than excess bicarbonate exerts a direct constraint on plant growth and ion nutrition (Bertoni et al. 1992; Pearce et al. 1999; Ding et al. 2020). Specifically, Ding et al. (2020) showed that a high concentration of bicarbonate ions diminished the negative effect of high pH (= 8) on nutrient translocation to aboveground organs of Lupinus angustifolius. This might be the truth for many experiments, in which high concentrations of bicarbonate usually co-occur with high pH. Hence, the real impact of bicarbonate on plant productivity might have been masked or at least partly offset by high pH, especially under the circumstance of a high dose of bicarbonate. In this case, the isotope technique, as we will discuss in the following context, serves as an important tool to quantitatively assess the possible fertilization effect of bicarbonate on plants. In addition, ammonium bicarbonate (NH4 HCO3 ) has been used as a major nitrogen fertilizer in some European and Asian countries (Li and Chen 1980; Zhang et al. 2011), despite its obvious shortcomings (e.g., instability, low nitrogen content, and high hygroscopicity) over other fertilizers

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(Roelcke et al. 2002; Zhang et al. 2011; Drapanauskaite et al. 2021). However, the uptake of HCO− 3 and its individual influence on the photosynthesis and biomass production of crops under field conditions are less studied.

5.2.3 Root Uptake of Bicarbonate Ion 5.2.3.1

Generation of Bicarbonate Ion

Apart from bicarbonate addition in hydroponic experiments, bicarbonate ions are widely present in natural waters (Raymond and Cole 2003; Raymond et al. 2008; Dickson 2010; Zhao et al. 2015; Huisman et al. 2018; Ni and Li 2022) and calcareous or saline-alkali soils (Tsypin and Macpherson 2012; Ma et al. 2013; Taalab et al. 2019; Wang et al. 2019c). The generation of bicarbonate ions usually has two sources, one from the dissociation of carbon acid after the hydration of gas phase CO2 in water (England et al. 2011) and another from the coupling of the hydrolysis of CO2 and dissolution of soluble rocks (Liu et al. 2008), namely, chemical weathering (see following Sect. 5.4.1). The whole products of CO2 dissolved in the water are 2− CO2 (aq), H2 CO3 , HCO− 3 , and CO3 (collectively referred to as dissolved inorganic carbon, DIC). The interconversion between these species is according to the firstand second-order dissociation of H2 CO3 resulting from the hydration of CO2 , which is mainly dependent on pH. In natural ecosystems, H2 CO3 is the most abundant 2− acid, while HCO− 3 and CO3 are the major contributors to total alkalinity in waters (England et al. 2011). The concentration of bicarbonate in soil solution is dynamic and related to the CO2 −HCO− 3 equilibrium system, which is primarily affected by the concentration of calcium carbonate (Sarkar et al. 2008) and soil moisture (Bloom and Inskeep 1986; Zuo et al. 2007).

5.2.3.2

Isotope Evidence and Ion Absorption Characteristics

The absorption of bicarbonate ions by plants is initially performed with 14 C feeding and radioautography, by which the radioactivity of 14 C is allocated into different organs and compounds can be detected. For example, Overstreet et al. (1940) showed that 4–5.1% of bicarbonate ions were fixed by barley, including 0.14–0.22% retained as [H2 CO3 ] or HCO− 3 in plant tissues and 3.86–4.93% retained as reduced carbon. This experiment provides new insight into the assimilation of bicarbonate by plants, although the proportion of HCO− 3 fixation is rather small. The subsequent work was extensively carried out with the advent of isotope techniques (including the widely used 13 C labeling technique) and focused on different aspects of DIC fixation (Arteca et al. 1979; Bialczyk and Lechowski 1992; Bialczyk et al. 2004). At least the following conclusions can be drawn: the 13 C- or 14 C-labeled DIC (mainly in the form of CO2 or HCO− 3 ) in the hydroponic solution is transported upward through the transpiration stream (Amiro and Ewing 1992; Stringer and Kimmerer

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1993; Bloemen et al. 2016), influences the carbon isotope composition of aboveground tissues (Stemmet et al. 1962; Vapaavuori and Pelkonen 1985; Vuorinen et al. 1989), and results in the generation of organic acids in roots and stems (Stringer and Kimmerer 1993; Bialczyk et al. 2004; Shahabi et al. 2005; Msilini et al. 2009; Covarrubias and Rombolà 2013; Zhao and Wu 2017), and accumulation of reduced carbon or starch in leaves and/or stems (Stringer and Kimmerer 1993; Vuorinen et al. 1992; Hibberd and Quick 2002). More recently, it was observed that H13 CO− 3 was rapidly assimilated by plants and became a part of nonstructural carbohydrates in different plant organs of Camptotheca acuminata within 24 h (Rao et al. 2019). The fast assimilation of DIC was consistent with an earlier study showing that H14 CO− 3 labeling within roots was translocated to the leaves of willow within 5–24 h (Vuorinen et al. 1989). In addition, an ion absorption experiment conducted by Rao and Wu (2017b) showed that the uptake rate of bicarbonate solution ranged from 32.60 to 103.59 ml/ day for Camptotheca seedlings under different water regimes, equivalent to 0.16– 0.39 μmol h−1 g−1 for the uptake rate of HCO− 3 ions. In the study of Fang and Wu (2022), a slightly low rate of bicarbonate absorption was observed in Broussonetia papyrifera (0.06–0.15 μmol h−1 g−1 ) and Morus alba (0.02–0.14 μmol h−1 g−1 ).

5.2.3.3

Possible Mechanisms for the Uptake of Bicarbonate Ions

Although a substantial amount of isotopic evidence demonstrates the utilization of bicarbonate by higher plants, it remains largely unknown how bicarbonate (ion) enters root cells and loads into the root xylem. Compared with CO2 , HCO− 3 is charged and thus not freely permeable to the plasma membrane. Previous studies have shown that water and ions can cross the cortex (including the rhizodermis and exodermis) via symplasmic and/or apoplasmic pathways (Ebrahimi and Bhatla 2012; White 2012a; Song et al. 2017). It may also be applicable for bicarbonate ions. Generally, the contribution of symplast to total root uptake is greater than that of apoplast (Zhan et al. 2018; Isayenkov and Maathuis 2019), while in some cases, apoplastic transport is likely more efficient for some ions or molecules (Wang et al. 2020b). However, when molecules arrive at the casparian band, the apoplasmic pathway is blocked. Thus, water and ions have to penetrate the plasma membrane, which becomes a rate limiting step for transport (Foster and Miklavcic 2017). In comparison to the apoplasmic pathway, the transport of HCO− 3 through the symplasmic pathway in root cells remains more uncertain. Current knowledge of the membrane transport of HCO− 3 is mainly obtained from mammals and aquatic plants (e.g., cyanobacteria, algae, seagrasses) (Walker et al. 1980; Larkum et al. 2017). In the following, we summarize four major mechanisms for the transport of bicabronate ions across the membrane in various types of organisms, which have important implications for investigation into the uptake of bicarbonate ions in root cells of higher plants. 1. Membrane HCO− 3 transporters. Transporters embedded in the plasma membrane play critical roles in the regulation of intracellular and extracellular pH (Bernardino et al. 2013; Romero et al. 2013). As has been identified in both

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mammals and aquatic plants (Boron 2004; Liu et al. 2012; Romero et al. 2013; Fang et al. 2021; Nawaly et al. 2022), the direct uptake of HCO− 3 is mediated by a variety of membrane-localized specific transporters according to their functions, such as acid extruders (Na+ –H+ exchangers, Na+ −HCO− 3 cotransporters, − /Cl transporter and V-ATPases, etc.) and acid loaders (Na+ -independent HCO− 3 − − − cotransporter). HCO /Cl transporters, also known as HCO− Na+ /HCO− 3 3 3 /Cl exchangers, are encoded by two distinct genes of the solute carrier family: SLC4A and SLC26A. For example, Chávez et al. (2012) showed that SLC26A3 and SLC26A6 participate in mouse sperm capacitation. In addition, an outward proton pump drives coupled bicarbonate and proton movement across the plasmalemma via an H+ -symport in seagrasses (Walker et al. 1980; Hellblom and Axelsson 2003; Uku et al. 2005; Larkum et al. 2017; Rubio et al. 2017). For land plants, no membrane HCO− 3 transporter has been identified in mesophyll cells of leaves, except for attempts to engineer the transporters of cyanobacteria and proteobacteria into the chloroplast inner-envelope membrane and thylakoids of terrestrial C3 plants (Parry et al. 2011; Price et al. 2011; Rottet et al. 2021). In addition, to the best of our knowledge, no studies have identified HCO− 3 transport embedded in the plasmalemma of root cells. 2. Anion channels. Ion channels have a conduction pathway that allows passive diffusion of cations and anions along the electrochemical gradient. Early studies focused on anion transport were mainly conducted on the cystic fibrosis transmembrane regulator (CFTR) protein of mammals. CFTR is responsible for Ca2+ − dependent HCO− 3 secretion via Cl channels (Seidler et al. 1997; Jung et al. 2013), which occurs in apical, colon, and salivary gland cells. Cl− channels are characterized by nonspecific anion selectivity, which means they can permeate many other anions (Choi et al. 2001; Quinton 2001; Qu and Hartzell 2008). Choi et al. (2001) demonstrated the important role of HCO− 3 transport in epithelial secretion and cystic fibrosis. Subsequent work further showed that the outward movement (or secretion) or influx of HCO− 3 was mediated or activated by CFTR protein, cAMP, short-chain fatty acid, forskolin, and so on in many mammalian cells (Hirono et al. 2001; Vidyasagar et al. 2004; Chávez et al. 2012; Kim et al. 2014; Shah et al. 2016). In higher plants, a variety of anion channel types in the plasma membrane have also been identified in both leaf and root cells (Roberts 2006; Kollist et al. 2011; Xue et al. 2011). It has been demonstrated that HCO− 3 functions as a small molecule activator of SLAC1 (slow anion channel-associated 1) anion channels in guard cells (Xue et al. 2011). SLAC1 was shown to be a physiologically relevant CO2 /HCO− 3 sensor in guard cells (Zhang et al. 2018), which responded to increasing concentrations of CO2 (Pantoja 2021). In addition, anion channels also fulfill several root-specific functions (Roberts 2006), such as nutrient uptake (Pantoja 2021), loading to the xylem and excretion in the root zones (Kollist et al. 2011). However, to date, no study has presented direct evidence of anion channels mediating the transport of HCO− 3 in root cells. 3. Extracellular acidification. Walker et al. (1980) proposed a model for Chara in which the local acidification at the outer surface of the cell was caused by active efflux of protons. This process increased the concentration of CO2 adjacent to

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the outer epidemal walls of some other aquatic plants, for example, seagrasses, the majority of macroalgae, and freshwater green algae (Raven and Hurd 2012; Larkum et al. 2017), thus facilitating the acquisition of CO2 from alkaline media (Prins and Helder 1980; Hellblom and Axelsson 2003). Additionally, periplasmic carbonic anhydrase (CA)-catalyzed HCO− 3 dehydration also occurs in the acid zone (Hellblom and Axelsson 2003; Uku et al. 2005), accelerating the fixation rate of CO2 (Larkum et al. 2018). This conclusion is based on the inhibition of HCO− 3 by acetazolamide (AZ, an inhibitor of extracellular carbonic anhydrase activity) (Uku et al. 2005; Raven and Hurd 2012). In comparison to membrane transport (relying on cotransporters or exchangers), the acidification mechanism seems more economical and faster. In higher plants, this function may be more pronounced. For instance, when plants are exposed to excess bicarbonate, roots can exudate organic acids to the rhizosphere. This, together with an outward proton pump, has the potential to enhance the extracellular acidification and uptake of HCO− 3 via its conversion to CO2 . 4. Carbonic anhydrase-catalyzed CO2 diffusion. CAs catalyze the interconversion between CO2 and HCO− 3 with great efficiency and are ubiquitous in nature (DiMario et al. 2017). Three families (α-, β- and γ-CA) are identified in higher plants, algae, and cyanobacteria, whereas only one type of CA (α-CA) is found in animal cells (Moroney et al. 2001). In algae, CAs are found in the mitochondria, chloroplast thylakoid, cytoplasm, and periplasmic space (Moroney et al. 2001). In C3 plants, CAs are generally located in the plasma membrane, chloroplast stroma, cytoplasm and mitochondria (Moroney et al. 2001; Hu et al. 2015; DiMario et al. 2017), while in C4 plants, β-CA is most abundant in the cytosol (Badger and Price 1994). Of many functions, CAs are characterized by their critical roles in improving the efficiency of inorganic carbon (CO2 and/or HCO− 3) transport and utilization in both aquatic (Raven and Hurd 2012; Larkum et al. 2017; Rubio et al. 2017; Jensen et al. 2020) and terrestrial plants (Wu and Xing 2012; Poschenrieder et al. 2018; Momayyezi et al. 2020). For higher plants, few studies have investigated the essential roles of CAs in root cells, especially when plants grow in alkaline or saline soils. It is also unclear whether CA is excreted from the epidermal cells of roots to catalyze the conversion of HCO− 3 to CO2 and thus facilitate the utilization of soil inorganic carbon. What is certain, however, is that CAs also occur in alkaline soils (Li et al. 2005; Sharma et al. 2009; Achal and Pan 2011) and are even more abundant than those in acidic soils (Jones et al. 2021), because the abundance of CAs increases with soil pH (Sauze et al. 2018; Jones et al. 2021). This has important implications for the CA-catalyzed transport of HCO− 3 in root zones, which may be one of the most likely reasons for the fast assimilation of HCO− 3 by aboveground tissues. Further identification of these possible mechanisms is expected to provide solid evidence for the membrane transport of HCO− 3 in root surface cells of terrestrial plants under alkaline soil conditions. When root cells cope with excess HCO− 3 , it is supposed that the expression of genes encoding membrane transporters, anion channels, and CAs will be upregulated to mediate HCO− 3 transport across the membrane and thus

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maintain the stability and equilibrium of ions and pH. Therefore, the key question for future study is whether the mechanisms presented in mammalian species and aquatic plants will be all or partly shown in the roots of higher plants. Additionally, membrane transport of DIC is not an independent action, which, as will be discussed in the following sections, is closely linked to and driven by some metabolic processes. Only by combining whole stages can we obtain a conceptual understanding of HCO− 3 utilization by plants.

5.2.3.4

Fates of Absorbed Inorganic Carbon

The fates of root-derived inorganic carbon can be tracked by isotope techniques. The metabolic pathways of absorbed bicarbonate include but are not limited to (1) anaplerotic fixation, which allows replenishment of the intermediate products involved in the tricarboxylic acid (TCA) cycle (Vuorinen et al. 1992; Cramer et al. 1995; Ford et al. 2007; for details see Sect. 5.2.4). This portion may be very large, as indicated by a dominant 13 C or 14 C sink in roots (Ford et al. 2007); (2) root exudation, in which organic acids (e.g., malate and citrate) generated from the carboxylation of bicarbonate are exuded to neutralize OH− and HCO− 3 and thus improve the rhizospheric conditions and nutrient uptake (Johnson et al. 1996; Kollist et al. 2011; Rose et al. 2011); and (3) corticular photosynthesis, in which inorganic carbon is photosynthetically fixed by stem or branch tissues (Hibberd and Quick 2002; Ford et al. 2007; for details see Sect. 5.2.6); (4) leaf photosynthesis (Vuorinen et al. 1992; Stringer and Kimmerer 1993; for details see Sect. 5.2.7); and (5) escape from leaves or stems in the form of CO2 (Bloemen et al. 2013; Shimono et al. 2019; Tarvainen et al. 2021; Salomón et al. 2022). Under light conditions, this portion is very small for leaves (Stringer and Kimmerer 1993; Bloemen et al. 2013; Stuz and Hanson 2019a) but higher for leaves in the dark (Stringer and Kimmerer 1993) and stems, the latter of which is variable and mainly depends on species differing in their transpiration stream fluxes and the barriers to radical CO2 diffusion caused by stem anatomy (Steppe et al. 2007; Ubierna et al. 2009; Salomón et al. 2022). In addition, a small quantity of root-derived inorganic carbon remains temporally in plant tissues (Stringer and Kimmerer 1993; Ubierna et al. 2009), particularly in night conditions, but may ultimately be involved in other metabolic processes allowing for a long life span of most plants.

5.2.4 Anaplerotic Fixation in Roots In root cells, part of the absorbed DIC reacts with phosphoenolpyruvate (PEP) with the calalysis of phosphoenolpyruvate carboxykinase (PEPc) and produces oxaloacetic acid (OAA), which is subsequently converted to malic acid. The accumulation of organic acids is a typical response of root cells to excess bicarbonate or CO2 -enriched solution (Zhao and Wu 2018; Molina and Covarrubias 2019). In

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addition to transport from roots to shoots (see Sect. 5.2.5) and exudation from roots to soils (Martínez-Cuenca et al. 2013), malate also plays a vital role in the anaplerotic pathway (Sagardoy et al. 2011), a reaction that replenishes some intermediate metabolites for the TCA cycle (Vega-Mas et al. 2019). This is because a portion of OAA and its precursor α-ketoglutaric acid involved in the TCA cycle are consumed by amino acid metabolism (Cramer et al. 1993; Bialczyk et al. 2004; Ford et al. 2007). The anaplerotic process usually occurs in fungal, mycorrhizal and root tissues (Jackson and Coleman 1959; Wingler et al. 1996; Chalot et al. 2002). As has been recorded by past studies (Vuorinen et al. 1992; Cramer et al. 1993; Bialczyk et al. 2004; Viktor and Cramer 2005; Ford et al. 2007), increased NH+ 4 availability stimulated anaplerotic fixation and increased the amount of soil DIC fixed in roots or mycorrhizae. In contrast, the NO− 3 supply exhibited a relatively low increase in DIC fixation (Cramer et al. 1993). In addition, root-zone DIC increased root PEPc activity (Vuorinen and Kaiser 1997; Matarese 2011), which was associated with the anaplerotic fixation rate (Sagardoy et al. 2011). Taken together, the cascade of DIC carboxylation (or dark fixation), anaplerotic fixation, TCA cycling, and amino acid metabolism represents a small but significant carbon budget in roots. The ecophysiological significance of anaplerotic fixation could be that it benefits root formation and N assimilation, particularly when encountering adverse environments.

5.2.5 Transport of Root-Derived Inorganic Carbon in Xylem Sap 5.2.5.1

Knowns and Unknowns for Xylem Transport

Xylem is a complex tissue that possesses both conducting and mechanical functions in vascular plants. Vessels (dead cells) and accompanying sieve tubes (living cells), the major components of xylem, serve as conduits transporting water and mineral elements (e.g., K+ , Ca2+ , Mg2+ , Zn2+ , Cl− , PO2− 4 , etc.) from roots to shoots. Currently, our knowledge of the thransport of root-derived inorganic carbon in xylem sap is limited, which raises many questions. For example, are these root-derived DIC directly transported upward like water and nutrients? Do they exhibit spatial– temporal variation in their concentrations or fluxes and differ among species? Are all xylem-transported carbon fixed photosynthetically? To address these questions, we may rely on classical isotope tracing techniques as well as some chemical analysis instruments with high precision.

5.2.5.2

Measurements of the Chemical Composition of Xylem Sap

2− As mentioned above, inorganic carbon (CO2 , HCO− 3 , and CO3 ), organic acids (e.g., malate, citrate), and amino acids (e.g., Gln, glu, Asn, Asp, etc.) may all be delivered

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upward within the xylem sap. Understanding the chemical compositions and their variations in xylem may shed light on the physiological responses to environmental cues in root zones, such as Fe or Zn deficiency, high pH, excess bicarbonate or salts. Conventional analysis of most chemical compositions of xylem sap usually requires destructive sampling of stems. As shown by Bialczyk et al. (2004), the epidemal layer and phloem were removed by ring girding, after which this section was washed with distilled water and then dried, eliminating the influence of phloem sap. Subsequently, this part of the stem was pruned with a sharp razor. The initial drops of xylem sap were discarded, and then the samples were collected in tubes for 10–15 min. To improve the collecting efficiency of xylem fluid, pressure chambers, such as the Scholander-type pressure chamber (Soilmoisture Equipment Corp., Santa Barbara, CA, USA), have become popular in recent years (Martínez-Cuenca et al. 2013). After collection, the sample tubes were immediately frozen with liquid nitrogen and subsequently stored at −20 °C until analysis. Organic and inorganic matter in xylem sap were analyzed with different methods and instruments. For example, the contents of carbohydrates, organic acids, and amino acids were determined with a high-performance liquid chromatography (HPLC) system (Bialczyk et al. 2004; Martínez-Cuenca et al. 2013). The column type, mobile phase, temperature, and injection volume of samples differed between these compounds. In addition, the measurements of amino acids should be previously treated with the method of derivatization (Bialczyk et al. 2004). Available data showed that the concentration of malate in xylem sap increased by 100%, with a mean value of 3.09 mM, when tomato seedlings were cultivated on media with 5 mM − HCO− 3 and NO3 (Bialczyk et al. 2004). The total concentration of amino acids was usually less than 2 mM and increased 17.5–48.5% when plants were exposed to NH+ 4 − − and NO− 3 in conjunction with HCO3 compared with non-HCO3 treatment (Bialczyk et al. 2004). As identified in tomato seedlings, the major components of amino acids were Gln, Glu, Asn, and Asp (Bialczyk et al. 2004). The concentration of CO2 in xylem sap can be measured in situ either with a CO2 microelectrode (for example, MI-720, Microeletrodes, Inc., London-derry, NH, USA; or advanced instruments) or an infrared gas analyzer (IRGA, LI 6400, Li-Cor Inc., Lincoln, NE, or an advantage version). The CO2 microelectrode is immersed in the xylem sap through a sealed vial; therefore, the concentration of CO2 is measured after a short stabilization period (Stringer and Kimmerer 1993). For the IRGA method, a purpose-built chamber is usually designed to install and seal onto the tree surface after drilling a hole in the stem (Levy et al. 1999). This chamber contains two outlets: one is used to remove samples, while another is attached to a balloon that allows the atmospheric pressure inside the chamber to be maintained. According to a mass balance model in a mixing system, the partial pressure of CO2 of the sample (pCO2(sample) ) is calculated as:  Vsample + Vsystem · pCO2(final) − Vsystem · pCO2(initial) = Vsample 

pCO2(sample)

(5.1)

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where V sample and V system are the volumes of the sample and IGRA system, respectively, pCO2(final) is the mixture pCO2 of the IGRA system and sample, and pCO2(initial) is the initial pCO2 of the IGRA system. Note that pCO2(sample) only represents the content of gas phase CO2 in xylem sap. Tarvainen et al. (2021) observed a range of 2700–9900 μmol mol−1 (or 0.27–0.99%) of root-derived CO2 in xylem sap of Pinus species, similar to the level (0.5%) observed in Abies grandis (Ubierna et al. 2009). Moreover, Teskey et al. (2008) reported a broad range of CO2 concentrations, varying from 0 to 26.3%. Assuming the gaseous CO2 is in equilibrium with water, the whole products of CO2 after dissolving the water ([CO2 *]) can be converted to the molar volume according to Henry’s Law (Stumm and Morgan 1996):     K 1 (T )K 2 (T ) K 1 (T ) + CO2 ∗ = pCO2 K H (T ) 1 + [H+ ] [H+ ]2

(5.2)

2− where [CO2 *] = CO2 (aq) + H2 CO3 + HCO− 3 + CO3 , K H (T ) is Henry’s constant for CO2 , K1 (T ) and K 2 (T ) are dissociation constants for bicarbonate and carbonate ions, and [H+ ] is the concentration of hydrogen ions. Marshall et al. (1994) estimated that the [CO2 *] of xylem sap ranged from 3 to 16 mM (mean value: 10 mM) in Juniperus osteosperma. The pH of xylem sap is determined by a pH meter. Levy et al. (1999) showed that the sap pH varied from 5.2 to 6.8 in three species (Betula pendula, Musanga cecropioides, Distemonanthus benthamianus), which was similar to that of other studies (Stringer and Kimmerer 1993; McGuire and Teskey 2002; Teskey and McGuire 2007). The weak acid solution in xylem sap indicated that nearly half of the total dissolved inorganic carbon was CO2 (Stringer and Kimmerer 1993). The flux of [CO2 *] in xylem sap can be calculated by two means: (1) multiplying the concentration of [CO2 *] by the transpiration rate at the leaf level (Levy et al. 1999) and (2) estimating from the measurements of the concentration of [CO2 *], the velocity of sap flow, and the atmotic weight of carbon (Teskey and McGuire 2007; Aubrey and Teskey 2009). The second method is based on the heat-pulse technique, and recently, a novel approach termed the “double–ratio method” (three-probe) allows the measurement of very high, very low and even negative velocities of xylem sap (Deng et al. 2020). Levy et al. (1999) and Aubrey and Teskey (2009) reported a large variation in xylem [CO2 *] flux among vegetation types (temperate forest, tropical rainforest, and tropical arid shrub), ranging from 7.5 to 12 μmol m−2 s−1 in the daytime. Ubierna et al. (2009) reported values of 1.8–2.0 μmol m−2 s−1 in two conifer species. Aubrey and Teskey (2021) showed that CO2 flux in the xylem sap of Populus deltoides was approximately in the range of 2–5 μmol m−2 s−1 . The internal transport rate of [CO2 *] seems larger in broadleaf species than in conifer species. It has also been shown that [CO2 *] in xylem sap varies diurnally and seasonally and relates to temperature and water availability (Steppe et al. 2007; Aubrey and Teskey 2009; Etzold et al. 2013; Salomón et al. 2016; Aubrey and Teskey 2021). This variation is linked to the negative relationship between the sap flow rate and the concentration of [CO2 *] (Brüggemann et al. 2011). The sap flow rate is closely related to the transpiration rate, while [CO2 *] is generally influenced by the strength of respired CO2

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from stems, roots and microbes and sap pH. Typically, factors affecting sap flux and tissue respiration will exert a strong influence on the concentration and flux of xylem sap [CO2 *].

5.2.5.3

In What Form Is Root-Derived Inorganic Carbon Transported?

It has been confirmed that 14 C or 13 C labels can be transported from roots to aboveground tissues through the transpiration stream; nonetheless, in what form the label is delivered remains unclear. Two hypotheses have been proposed concerning this long-distance transport. The first and most popular view is the direct transport of DIC within xylem sap (Enoch and Olesen 1993; Stringer and Kimmerer 1993; Ford et al. 2007; Teskey et al. 2008; Grossiord et al. 2012; Bloemen et al. 2013; Bloemen et al. 2016; Stuz and Hanson 2019a; Tarvainen et al. 2021; Salomón et al. 2022). Given the high solubility of DIC in water, it is reasonable that the transpiration stream could provide a conduit for the internal delivery of DIC to shoots (Stringer and Kimmerer 1993). Although the diffusion coefficients of HCO− 3 and dissolved CO2 are low in the water (1.17 × 10−9 m−2 s−1 and 2.02 × 10−9 m−2 s−1 at 25 °C, respectively; Zeebe 2011), the transportation of inorganic carbon is driven by transpiration and root pressure (Stringer and Kimmerer 1993; White 2012b; Bloemen et al. 2016), similar to the function of an elevator. Under natural conditions, the internal transport of DIC can be corroborated with direct evidence from the measurements of fluxes and/ or concentrations of inorganic carbon in xylem sap (Stringer and Kimmerer 1993; Bloemen et al. 2016). As shown by Teskey and McGuire (2007), the diurnal variation in CO2 flux in the xylem sap of Platanus occidentalis ranged from 0 μmol m−2 s−1 at night to approximately 50 μmol m−2 s−1 in the daytime. In Populus deltoides, the maximal value of CO2 flux reached 12 μmol m−2 s−1 at peak sap flux during the day (Aubrey and Teskey 2009). Additionally, recent years have seen advances in commercial CO2 sensors, allowing high-frequency and high-definition measurements of CO2 concentration and flux in the xylem sap of a large number of species, whether for short- or long-term observations (Paudel et al. 2018; Aubrey and Teskey 2021; Salomón et al, 2021). The second viewpoint is that part of root-derived DIC is initially fixed anaplerotically and then transported upward in the form of organic acids (e.g., malate and citrate) and/or amino acids (Vuorinen et al. 1992; Cramer et al. 1993; Hibberd and Quick 2002; Ford et al. 2007; Alhendawi 2011; White 2012b; Fang and Wu, 2022), which constitute a significant quantitative element of the xylem sap (Bialczyk et al. 2004). This conclusion is based on the fact that there is a significant increase in organic acids (mainly malate) in roots, stems, and leaves after exposure to excess bicarbonate or CO2 -enriched solution (Bialczyk and Lechowski 1995; Bialczyk et al. 2004). Furthermore, organic acids are commonly observed in the xylem stream even without a supply of DIC in root zones (Raven and Smith 1976; López-Millán et al. 2009; Larbi et al. 2010). Organic acids are generally low-weight molecules, which have diffusion coefficients of 0.5 × 10−9 m−2 s−1 and 0.71 × 10−9 m−2 s−1 for mailc acid and citric acid, respectively (Delhaize et al. 1993; Liu et al. 2004), slightly lower

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than that of DIC. Similarly, xylem loading allows the translocation of organic acids to the shoots rapidly (Kollist et al. 2011). Another explanation for the presence of organic acids in xylem sap is providing carboxylates to maintain ionic balance (Cramer et al. 1993). Obviously, both hypotheses are supported by considerable experimental data. A further question is which means dominate the long-distance transport of root-derived DIC under different conditions. A labeling experiment conducted by Vuorinen et al. (1992) revealed that in a short time, such as 1 h, most of the 14 C was incorporated into organic acids and amino acids in all tissues of Populus deltoide under both light and dark conditions. Bialczyk et al. (2004) showed that less than 8% of DIC taken up by roots of tomato seedlings was recovered in inorganic matter, suggesting that most DIC was transported to the shoot in the form of organic acids. In addition to the isotope evidence, the content of organic acids (chiefly malate) in the xylem sap ranged from 1.20 to 3.09 mM under different nitrogen regimes in combination with the addition of bicarbonate (Bialczyk et al. 2004). In sugar beet plants, the concentration of malic acid in xylem sap was in the range of 0.46–1.96 mM under different zinc treatments (Sagardoy et al. 2011). In Citrus sinensis (L.) Osbeck. × Poncirus trifoliata (L.) Raf, the level of malate was approximately 0.5 mM, while the concentration of citrate varied from approximately 0.5 to nearly 1 mM under no supply of bicarbonate (Martínez-Cuenca et al. 2013). Salomón et al. (2022) reported 2.4–9.7 mM in Cedar, Maple, and Oak. By comparison, the concentration of DIC in xylem sap varied considerably from less than 1 mM to more than 30 mM among species (Teskey et al. 2008; Aubrey and Teskey 2009, 2021), averaging approximately 10 mM. This may imply that even in normal conditions, for example, no supply of bicarbionate or CO2 -enriched solution in root zones, the DIC concentration represents a large flux of inorganic carbon originating from belowground respirotary CO2 and transported within xylem sap. Howerer, this does not mean that DIC is the major component all the time, as a situation with a high concentration of organic acids and a low level of DIC may exist in some species. Furthermore, organic acids may be the primary form in the early stage of dark fixation in roots and in the dark (Stringer and Kimmerer 1993) but possibly limited to a low level in xylem sap due to low transpiration. There is also another possibility that when the concentration of root-zone DIC is low, DIC is transported preferentially in the form of organic acids. When the DIC concentration is very high (10 mM, for instance), the amount of organic acids transported will increase disproportionately due to the limited increase in PEP content and PEPc activity. In such cases, the remaining DIC will be directly transported to aboveground tissues.

5.2.5.4

Sources and Fates of Chemical Compounds in Xylem Sap

[CO2 *] in the xylem sap has two major sources: (i) root- and/or stem-respired CO2 and (ii) external inorganic carbon absorbed by roots. These components differ in concentration or flux and isotopic signal. The concentration of respiratory CO2 from stem tissues, published ranging from 0.1 to 13.5% (Cernusak and Marshall, 2000; Ubierna et al. 2009), is much higher than that of xylem-transported CO2 . In species

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with ring and diffuse pores, the fates of xylem [CO2 *] are that most [CO2 *] is continually transported upward and a small proportion diffuses outward, acting as a part of stem CO2 efflux to the atmosphere (McGuire and Teskey 2002; Teskey and McGuire 2007; Saveyn et al. 2008; Bloemen et al. 2013; Salomón et al. 2019). Ubierna et al. (2009) estimated that the CO2 flux transported upward represents only 1–3% of the stem CO2 efflux to the air. Salomón et al. (2016) showed that the contribution of xylem-transported CO2 to stem CO2 efflux was in the range of 13–38%. In the study of Tarvainen et al. (2021), most of the vertical-transport CO2 in the xylem sap of mature Pinus sylvestris trees was lost by radical diffusion, constituting an important part of stem CO2 efflux. This flux may be considerable in some conifer species and is significantly related to tree age, pit vessels, sap flow rate, and stem height (Ubierna et al. 2009). Nevertheless, the underlying mechanism probably relates to the relative velocity between stem CO2 efflux and sap flux, which has been observed in both conifer and broadleaf species (Kunert and Edinger 2015). As shown by Stringer and Kimmerer (1993), the concentration of [CO2 *] decreased by more than 60% during the day, indicating that most of the xylem-transported [CO2 *] was translocated to petioles and leaves, either diffusing out or fixed photosynthetically. The sources of organic acids and amino acids are relatively clear, coming from the dark fixation of inorganic carbon (anaplerotic reaction) and nitrogen assimilation (amino acid metabolism coupled with the TCA pathway), respectively. Amino acids are terminally used for synthesizing proteins, facilitating the root uptake of macroelements, improving photosynthesis and plant growth, regulating the osmotic potential, resisting the attack of phytopathogens, and so on. For organic acids, the primary fates involve the regulation of stomatal gurad cells, balancing ion and base movement (Hibberd and Quick 2002), adjusting intracellular or extracellular pH (Molina and Covarrubias 2019), or decarboxylation to CO2, which will be refixed later through photosynthesis (Hibberd and Quick 2002; Rombolà et al. 2005).

5.2.6 Assimilation of Xylem-Transported Carbon Sources Through Corticular Photosynthesis The discovery of corticular photosynthesis dates back to a century ago. It describes that plant parts other than leaves can recycle CO2 via photosynthesis. In the early stages of plant growth or growing seasons, most stems and branches of woody plants form greenish tissues beneath the periderme and rhytidome. These tissues contain chlorophyll and allow recapturing respiratory CO2 and using light penetrating the rhytidome to conduct photosynthesis (Pfanz et al. 2002). In addition, petioles, veins, or calyxes of many species also contain chloroplasts that enable corticular photosynthesis (Foote and Schaedle 1976; Stringer and Kimmerer 1993; Pfanz et al. 2002; Bloemen et al. 2013). Collectively, this pathway is slightly different from conventional foliar photosynthesis, as the former utilizes the respired CO2 by plant tissues, termed “refixation” or “recycling”, while the latter directly absorbs

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atmospheric CO2 (Cernusak and Hutley 2011). The substrate for corticular photosynthesis principally comes from xylem-transported CO2 and/or local respired CO2 in stem tissues (Cernusak and Marshall 2000). Species with thin bark and green tissues or at a young age may refix a larger amount of CO2 compared with thick bark (Bloemen et al. 2016). It is unlikely that external (atmospheric) CO2 can be fixed by the stem bark due to the existence of a concentration gradient of CO2 from stems to ambient air, also known as stem CO2 efflux. In addition, malate produced from dark fixation is an indirect substrate used by corticular photosynthesis. For example, Hibberd and Quick (2002) reported characteristics of C4 photosynthesis in cells of stems and petioles of tobacco (a C3 plant). They demonstrated corticular photosynthesis via both 14 C-labeled bicarbonate and malate supplied to the xylem stream. The incorporation of 14 C into insoluble material, most notably starch, was found in cells around the vascular bundles. This phenomenon was consistent with the evidence of chlorophyll fluorescence and high activities of decarboxylation enzymes and phosphoenolpyruvate orthophosphate dikinase at the same site. Attempts to quantify the rate of corticular photosynthesis have been made since the middle of the twentieth century. Given the difficulties in the direct measurement of corticular photosynthesis of intact organs, most available positive net photosynthesis data were obtained from isolated chlorenchymes or peeled tissues (Pfanz and Aschan 2001). However, this method may not be able to disentangle the influence of stem respiration on the calculation of corticular photosynthesis in intact plants. Relevant models have been proposed to assess the proportional contribution of corticular photosynthesis to the formation of wood tissues (Cernusak and Hutley 2011). Ávila et al. (2014) showed that the corticular CO2 fixation rate can reach up to 60% of the leaf photosynthetic rate, and a higher level up to 75% was observed in some species (Pfanz et al. 2002). The contribution of xylem-transported CO2 to corticular photosysnthesis may be small; as noted previously, the contribution of xylem-transported CO2 to stem CO2 flux is also very low in some species (Ubierna et al. 2009). This is attributed to the strong diffusion barriers created by the cell walls of xylem and cambium tissues (Molina and Covarrubias 2019). Precise determination of the CO2 assimilation rate of the in vivo cortex may still rely on the isotope labeling technique that has been applied to estimate the contribution of root-derived inorganic carbon to leaf total photosynthesis (see Sect. 5.2.7). Furthermore, investigation into corticular photosynthesis helps to realize its important eco-physiological significance. First, corticular photosynthesis has been identified as an important mechanism for species recycling respiratory CO2 before diffusing out of the stem (Levy et al. 1999; Pfanz et al. 2002) and contributing to the growth of stems or branches (Cernusak and Hutley 2011). As indicated by De Roo et al. (2019), photosynthetically fixed CO2 produced by stem respiration can lead to a reduction in stem CO2 efflux of up to 22%. In extreme conditions, corticular photosynthesis can compensate for 60–90% of stem respiratory CO2 (Pfanz et al. 2002). Second, the refixed CO2 respired by local wood tissues is not considered net CO2 uptake (Wittmann et al. 2006; Brüggemann et al. 2011), whereas xylem-transported CO2 and malate originating from root uptake of DIC definitely increase the plant total carbon gain. Third, corticular photosynthesis is shown to be impacted by growing

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stage, light intensity, temperature, pollution, or pathogens. Their response may be different between species, thus inevitably affecting the carbon budget of plants.

5.2.7 Assimilation of Xylem-Transported Carbon Sources via Leaf Photosynthesis 5.2.7.1

Leaf Photosynthesis: A Hotspot for the Assimilation of Various Inorganic Carbon Sources

After traveling through the stems, branches, and petioles, the xylem-transported carbon sources (e.g., DIC, organic acids) finally arrive at leaves. However, not all of the root-derived carbon sources can be conveyed to this location because, as stated before, most of them have been fixed in specific tissues or released to the ambient environment during long-distance xylem transport (Ford et al. 2007). Inside the leaves, xylem-transported carbon sources travel from small veins to mesophyll cells (Hanson et al. 2016; Stuzz and Hanson 2019a), either apoplastically or symplastically (Buckley 2015). The pathway is slightly different from the absorption of atmospheric CO2 , which must cross the stomatal cavity (gas phase diffusion) before entering the mesophyll cells (liquid phase diffusion). Accumulations of DIC and malate in the cytosol will be subject to photosynthetic fixation proceeding in the chloroplast (Brüggemann et al. 2011; Poschenrieder et al. 2018), as could be shown in NaH14 CO3 labeling studies (Stringer and Kimmerer 1993). The magnitude of this fixation pathway has been quantified in many studies; however, larger discrepancies exist among those applying different methods. Below, we list several kinds of representative approaches to calculate the amount of xylemtransported carbon sources fixed by foliar photosynthesis or otherwise estimate its contribution to leaf total photosynthesis or plant total carbon gain.

5.2.7.2

High Abundance of 13 C Labeling and Tracking

This is the most widely used method, which clearly reveals the distributions of 13 C tracers in different organs or compounds (Shimono et al. 2019). For instance, Ford et al. (2007) showed that the average distributions of labeled 13 C were 19.3%, 35.7% and 45% in leaves, stems and roots (including mycorrhizal and nonmycorrhizal) of Pinus taeda, respectively. The total carbon gain could be viewed as the sum of soil DIC fixation, apparent photosynthesis and respiration measured by commercial infrared gas analyzers (Ford et al. 2007; Han et al. 2022). Note that the calculation of soil DIC fixation comprises the leaf photosynthesis contributed by the assimilation of xylem-transported carbon sources. Moreover, the high abundance of 13 C labeling may not be strong enough to be detected in large trees in comparison to seedlings (Ubierna et al. 2009; Rao et al. 2019), which is mainly due to the plant height, the

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volume of stems and the concentration of DIC feeding the roots or stems. Tarvainen et al. (2021) also found that leaf total photosynthesis was almost not influenced by xylem CO2 transport along the root-stem-leaf continuum in mature Pinus sylvestris trees. Therefore, it is often observed that root- or stem-derived DIC contributes only or less than 1% to plant carbon gain, accounting for large biomass of plant (Enoch and Olesen 1993; Ford et al. 2007) or leaf total photosynthesis (also see 14 C labeling; Vapaavuori and Pelkonen 1985; Vuorinen et al. 1989), and thus concludes that the effect of soil DIC on plant growth can be ignored.

5.2.7.3

Xylem CO2 Flux

This method is based on the assumption that root-derived DIC is directly transported upward and fixed by leaf photosynthesis. In early studies, the effect of xylem transport CO2 on leaf gas exchange is commonly quantified by expressing the ratio of stem CO2 flux to leaf photosynthesis, assuming that all aqueous transport of CO2 is assimilated by leaves (Stringer and Kimmerer 1993). For example, Levy et al. (1999) showed a range of 0.5–7.1% of the net photosynthetic rate originating from stem CO2 , which is similar to the 2–9% of photosynthesis in Scots pine observed by Hari et al. (1991). The magnitude of this flux is comparable to the leaf photosynthetic rate (Bloemen et al. 2016); thus, it usually represents an important fraction contributing to the total photosynthetic rate. Nevertheless, this approach may overestimate the contribution of xylem CO2 . Studies have shown that a proportion of xylem CO2 may diffuse outward through stems and/or leaf stomata or is fixed via corticular photosynthesis.

5.2.7.4

Parasitism Model

This is a special example once used for estimating the contribution of nutrition of the host tree to parasitic mistletoe (Marshall et al. 1994). In addition to atmospheric CO2 , the mistletoe acquires either dissolved inorganic carbon from xylem sap (xylem-trapping pattern) or organic carbon from the phloem (phloem-trapping pattern). For example, uptake of DIC from the host, which imparts a more negative or positive isotopic signal to the mistletoes, can cause a significant difference between the carbon isotopic ratios of xylem-trapping mistletoes (observed δ13 C) and those estimated from the photosynthetic carbon discrimination model (predicted δ13 C). Such discrepancy can be offset by accounting for the contribution of carbon assimilation from xylem sap of the host through a simple isotopic mixing model. Marshall et al. (1994) reported a mean value of 15% over 11 species for this contributor, whereas other studies reported a much higher proportion up to 60% (Marshall and Ehleringer 1990; Schulze et al. 1991).

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5.2.7.5

167

Bidirectional Isotope Labeling (or “Two-Source 13 C Labeling”)

It was developed by the research group of Yanyou Wu (Wu and Xing 2012; Rao and Wu 2017b), who used a pair of isotope mixing models to eliminate some uncertainties or unknown parameters when considering the photosynthetic assimilation of both atmospheric CO2 and root-derived DIC in chloroplasts of mesophyll cells (for more details, see Chap. 4). The relevant studies usually show relatively high contributions (10–28%) of root-derived DIC to leaf total photosynthesis under the condition of high concentrations of bicarbonate (e.g., 10 mM).

5.2.7.6

On-Line Measurement of 13 C Flux

The laser-based 13 C technique enables real-time measurements of 13 CO2 and 12 CO2 entering and leaving the leaf cuvette and thereafter the photoassimilated flux of xylem-transported CO2 on the basis of mass balance (Stutz and Hanson 2019a, b). The results of Stutz and Hanson (2019a) revealed that approximately 2.5% of the total photosynthesis was attributed to xylem-transported CO2 in two C3 species under saturated irradiance and a supply of 11.9 mmol/L NaH13 CO3 . However, under low irradiance or low intercellular CO2 , the contribution of xylem CO2 increased up to 10%.

5.3 Root-Derived Bicarbonate Assimilation by Plants Under Simulated Karst Environments 5.3.1 Revisiting the Simulation Experiments In the previous sections, we have already discussed how plants assimilate xylemtransported inorganic carbon. Most studies have been conducted with detached twigs and leaves exposed to low levels of DIC; however, few of them have focused on the influence of DIC on intrinsic photosynthesis. In this section, we revisit some studies that investigated how intact plants utilized root-derived bicarbonate through foliar photosynthesis under simulated karst environments. In contrast to normal soil conditions, the karst soil environments,—as we will describe elaborately in Sect. 5.4—are usually characterized by high concentrations of bicarbonate, high pH and a high frequency of soil water limitation. The cases presented below adopt the bidirectional labeling or high abundance of 13 C labeling approaches to quantify the proportional contribution of rhizospheric DIC (mainly in the form of bicarbonate ion) to leaf total photosynthesis or total carbon reservoir in different types of plants.

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5.3.2 Bicarbonate Stress on Photosynthetic Inorganic Carbon Assimilation Case 1 In the study of Wu and Xing (2012), two Moraceae seedlings, Broussonetia papyrifera and Morus alba, were subjected to 10 mM bicarbonate in hydroponic culture for 20 days. Two treatment groups, which contained modified Hoagland nutrient solution with pH set to 8.1, had two different carbon isotope siganals (δ13 C) of NaHCO3 labels. One was −17.4‰ (L group), and the other was −6.7‰ (H group). The calculations of the bicarbonate utilization proportion and total photosynthesis were performed according to a bivariate isotope mixing model as described in Chap. 4. As shown in Table 5.1, the inhibition of bicarbonate on the net photosynthetic rate originating from atmospheric CO2 (A) and root-zone bicarbonate (ADIC ) increased with time in B. papyrifera, while in M. alba, the inhibition decreased, and even an increase in A on Day 20 was observed compared with that on Day 10. The ADIC of M. alba was extremely low during 20 days of treatment. This led to a low contribution of bicarbonate to leaf total photosynthesis ( f DIC ) in M. alba compared with approximately 30% in B. papyrifera. Furthermore, the activity of carbonic anhydrase (CA) seemed to explain the difference in f DIC between the two species because CA catalyzed the interconversion between HCO− 3 and CO2 and therefore facilitated the assimilation of root-derived bicarbonate in leaves. The authors finally concluded that the more bicarbonate was assimilated, the more harmful it was to leaf photosynthesis. Although the double effect of bicarbonate has also been observed elsewhere, the conclusion based on the data in Table 5.1 may be debatable. As mentioned previously, the reduction in foliar photosynthesis after exposure to bicarbonate is more likely due to the associated effect of high pH (Pearce et al. 1999; Ding et al. 2020). Based on this premise, it can be concluded that bicarbonate increases the photosynthetic capacity; however, this effect differs largely in the two species. Case 2 In the study of Hang and Wu (2016), two Brassicaceae seedlings, Orychophragmus violaceus and Brassica juncea, were exposed to 5, 10, and 15 mM Table 5.1 Carbonic anhydrase (CA) activity, net photosynthetic rate originating from atmospheric CO2 (A) and root-zone bicarbonate (ADIC ), leaf total photosynthesis (Atotal ), and the proportional contribution of root-derived bicarbonate to leaf total photosynthesis ( f DIC ) of two Moraceae plants under the supply of bicarbonate in hydroponic culture B. papyrifera

Parameters CA (WAU g−1 FW) m−2 s−1 )

M. alba

Day 10

Day 20

Day 10

Day 20

3818 ± 76

4674 ± 66

758 ± 9

171 ± 8

5.91 ± 0.34

6.13 ± 0.02

4.08 ± 0.39

4.89 ± 0.21

A (μmol m−2 s−1 )

2.49 ± 0.14

1.91 ± 0.10

1.82 ± 0.32

2.55 ± 0.18

ADIC (μmol m−2 s−1 )

0.97

0.74

0.15

0

m−2 s−1 )

3.46

2.65

1.97

2.55

28.03

27.92

7.61

0

Acontrol (μmol

Atotal (μmol f DIC (%)

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Table 5.2 Net photosynthetic rate originating from atmospheric CO2 (A) and the proportional contribution of root-derived bicarbonate to leaf total photosynthesis ( f DIC ) of two Brassicaceae plants under the treatments of different bicarbonate levels in hydroponic culture Species

Parameters

Bicarbonate level (mM) 5

Orychophragmus violaceus

A (μmol m−2 s−1 )

Brassica juncea

f DIC (%)

10

15

5.8

5.5

4.8

f DIC (%)

5.28

13.28

17.31

A (μmol m−2 s−1 )

6.5

7.6

7.5

3.28

3.10

3.09

bicarbonate in hydroponic culture for 7 days. The δ13 C values of bicarbonate in the two labeling groups were −2.45‰ (H group) and −24.41‰ (L group). The pH of the hydroponic solution was set to 8.3. In O. violaceus, A decreased with an increasing concentration of bicarbonate, while an opposite trend was observed in B. juncea (Table 5.2). Obviously, the response of A to excessive bicarbonate in herbaceous plants is totally different from that in tree species reported by Wu and Xing (2012). The f DIC of O. violaceus increased from 5.28 to 17.31% when the concentration increased from 5 to 15 mM, whereas the f DIC of B. juncea remained constant (~3.2%). Their subsequent work (Hang and Wu 2019) also exhibited higher CA activities in O. violaceus than in B. juncea, which partially accounted for the higher f DIC in O. violaceus. In addition, Hang and Wu (2019) considered the contamination of DIC solution by CO2 in ambient air, which altered the δ13 C of the former. Therefore, they calculated the relative contributions of initial DIC and atmospheric CO2 to the mixed solution and finally to total photosynthesis. This is one of the advantages that allows bidictional labeling with a natural abundance of 13 C conducted in an open environment.

5.3.3 Interactive Effect of Bicarbonate Excess and Water Stress on Photosynthetic Inorganic Carbon Assimilation Case 3 In the study of Wang et al. (2017), three biofuel energy plants, O. violaceus, B. juncea and Euphorbia lathyris, were exposed to 10 mM/kg bicarbonate combined with three water regimes (soil water content (SWC): 17%, 14%, and 11%, respectively) in pots (the substrate contained a 1:1 quartz sand to vermiculite mixture, m/m) for 7, 14, and 21 days, respectively. The δ13 C values of bicarbonate in the two labeling groups were −18.81‰ (H group) and −27.28‰ (L group). The pH of Hoagland nutrient solution irrigated to the substrate was set to 8.1. In comparison to the hydroponic culture, plants grown in this substrate were closer to those in karst soils due to aeration conditions. This was favorable for the uptake of water and nutrients by roots. As time passed, A (or shown as Pn in Fig. 5.1) changed slightly under

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5 Root-Derived Inorganic Carbon Assimilation by Plants in Karst …

17% SWC but decreased drastically under 14% and 11% SWC among all species. Under three water regimes, f DIC was generally more than 12.5% in all species. In addition, the f DIC of the three species responded differentially to water stress: O. violaceus initially increased but subsequently decreased when the strength of water stress was enhanced; B. juncea exhibited a decreasing pattern when SWC decreased from 17 to 11%; and E. lathyris continually increased with a decrease in SWC. After 21 days of treatment, the highest values of f DIC were 26.72% for O. violaceus under 14% SWC and 22.10% for B. juncea and 26.95% for E. lathyris under 11% SWC. Such high proportions of f DIC and photosynthetic acclimation to root-zone DIC in the three biofuel enegy plants is most likely responsible for their adaptation to karst soils, where a high content of DIC and frequent water limitation usually threaten plant survival. Case 4 In the study of Rao and Wu (2017b), a karst-adaptive species Camptotheca acuminata was used to assess its bicarbonate assimilation under the conditions of 10 mM bicarbonate combined with variable water deficit (well-watered, moderate and severe stress). The water stress treatments were implemented with the addition of polyethylene glycol (PEG 6000) to control the water potential in the solution. The δ13 C values of bicarbonate in the two labeling groups were −9.76‰ (H group) and −26.78‰ (L group). The pH of the hydroponic solution was set to 8.3. Under well-watered treatment, A increased daily within 14 days. Moderate stress induced a decrease in A by 25–50% during the second 7 days, whereas severe stress strongly hampered A during 14 days of treatment. This led to f DIC increasing from 10.34%

Fig. 5.1 Effect of different levels (Con: 25% moisture, D1 : 17% moisture, D2 : 14% moisture, and D3: 11% moisture) of drought stress on Pn (net photosynthetic rate, (μmol·m2 ·s−1 ); (a–c) and proportion of HCO− 3 (d–f) use in the leaves of three plant species. Ov: Orychophragmus violaceus; Bj: Brassica juncea; and El: Euphorbia lathyris. The different letters above the bars represent significant differences among treatments on the same day (p < 0.05). Reprinted with permission from Wang et al. (2017). Copyright 2017 MDPI Publisher

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Fig. 5.2 Leaf total photosynthesis and the contribution of root-derived bicarbonate under the three water deficits in both L and H. Pre-trt: pretreatment; WW, well-watered; MS: moderate stress; SS: severe stress. Reprinted with permission from Rao and Wu (2017b). Copyright 2017 Springer Press

under well-watered conditions to 20.05% under moderate stress but decreasing to 16.60% under severe stress (Fig. 5.2). The magnitude of f DIC under moderate and severe stress was lower than that of the tree species B. papyrifera observed in Wu and Xing (2012) but close to those of three herbaceous species shown in Wang et al. (2017), indicating a common capability of assimilating bicarbonate under the adverse environment of karst. Additionally, the work of Rao and Wu (2017b) highlights how bicarbonate assimilation contributed to leaf photosynthesis from the perspective of isotopes. Traditionally, the major challenge underlying the natural abundance of 13 C labeling is that the carbon isotope signal of bicarbonate may not be detectable in leaves after undergoing long-distance transport in xylem sap and incorporation into photosynthates. However, Rao and Wu (2017b) showed that the δ13 C of leaves under the three water deficits in the low abundance group (−26.78‰) was approximately 1–1.5‰ less than that in the high abundance group (−9.76‰) (p < 0.01). The systematic discrepancy provided direct evidence for the assimilation of bicarbonate in C. acuminata. Furthermore, Rao and Wu presented the relationship between C i /C a and observed photosynthetic carbon discrimination (△13 Cair-leaves ). Most of the △13 Cair-leaves significantly deviated from the dotted line (predicted photosynthetic carbon discrimination based on C i / C a ), as shown in Fig. 5.3, supporting the high values of f DIC under moderate and severe stress.

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5 Root-Derived Inorganic Carbon Assimilation by Plants in Karst …

Fig. 5.3 Relationship between △13 Cair-leaves and Ci/Ca in L (open dots and dashed line) and H (black dots and solid line). Dot line, theoretical regression equation: y = 4.4 + 22.6x. N = 27. Reprinted with permission from Rao and Wu (2017b). Copyright 2017 Springer Press

5.3.4 Stimulation of Bicarbonate on the Plants’ Total Carbon Gain and the Dynamics of Nonstructural Carbohydrates Case 5 Although the role of root-derived DIC in leaf photosynthesis has been extensively studied, the effect of assimilated DIC on nonstructural carbohydrate (NSC) levels has not been elucidated. Rao et al. (2019) applied a high abundance (10% atm.) of NaH13 CO3 labeling to investigate the incorporation of newly assimilated DIC into the plant organs and NSC of C. acuminata in the short term (24 and 72 h). The hydroponic solution contained 5 mM NaH13 CO3, and the pH was adjusted to 7.5 ± 0.2. Their study showed that the content of major NSC compounds (soluble sugars and starch) substantially increased in stems and leaves, while a slight but significant change was also observed in roots. This meant that root-derived bicarbonate mightily stimulated the NSC pools in the majority of plant organs (Fig. 5.4). Furthermore, the isotope evidence exhibited the most 13 C-enriched NSC in roots compared with shoots, demonstrating that the newly formed photosynthates originating from bicarbonate were incorporated into all NSC pools within a short time. Although the contributions of bicarbonate to different carbon compartments (e.g., organ or compound level; Table 5.3) were all small, its effect on NSC functions (such as regulating plant growth, osmosis, defense, and reproduction) was worthy of attention. Altogether, the results obtained in Rao et al. (2019) revealed the double effects of bicarbonate, which extends our understanding of the postphotosynthetic carbon allocation pattern concerning the assimilation of root-zone DIC.

5.4 Characteristics of the Karst Environment

173

Fig. 5.4 Schematic concept showing the stimulation of bicarbonate on NSC pools in different organs of C. acuminata. Initially, bicarbonate-induced stress leads to a decline in gs and thus reduces photosynthetic CO2 uptake. Then, the stored C reserve is triggered and becomes mobilized NSC in different organs. At the same time, photosynthates supplied from bicarbonate and CO2 , as well as some mobilized NSC, are both transported downward but constrained by bicarbonate to some extent. Finally, the mixture of old NSC, newly fixed photosynthates and stored carbon reserves shapes the size of the new NSC pools across the organs. Box and arrow sizes are roughly proportional to the NSC pool sizes and fluxes among the organs. P means photosynthates. PC and PB represent newly fixed photosynthates supplied from CO2 and bicarbonate, respectively. The direction (dotted arrow) of the bicarbonate stress effect on fluxes or pools are indicated by + (increase) and − (decrease). Reprinted with permission from Rao et al. (2019). Copyright 2019 John Wiley & Sons Press

Table 5.3 Contribution of bicarbonate ( f B ) to the δ13 C of different compartments after 24 and 72 h labeling. The p-values are obtained from multiple comparison analysis; r 2 represents the adjusted coefficients of determination Soluble carbohydrates (%)

Starch (%)

Roots

Organs (%) Stem

Leaves

Roots

Stem

Leaves

Roots

Stem

Leaves

24 h

0.11 (0.01)

0.05 (0.01)

0.02 (0.01)

0.05 (0.01)

0.02 (0.01)

0.01 (0.01)

0.11 (0.01)

0.02 (0.01)

0 (0.00)

72 h

0.14 (0.00)

0.08 (0.01)

0.03 (0.00)

0.06 (0.01)

0.06 (0.01)

0.02 (0.00)

0.24 (0.00)

0.08 (0.01)

0.01 (0.00)

p

0.05

0.14

0.14

0.63

0.06

0.36

0.01

0.01

0.09

r2

0.51

0.33

0.38

0.04

0.47

0.14

0.99

0.76

0.40

5.4 Characteristics of the Karst Environment 5.4.1 Karst and Karstification Karst can be defined as the chemical dissolution and mechanical actions (e.g., erosion, corrosion, and collapse) of highly soluble rocks such as carbonate, marble, and gypsum (Lian et al. 2011; Hartmann et al. 2014; Harmand et al. 2017; Veress 2020) and the resultant landscapes (Ford and Williams 2007). Karstification, a geological terminology derived from the root word “karst”, specifically refers to these chemical and physical processes. In fact, karstification has a very rich connotation according

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5 Root-Derived Inorganic Carbon Assimilation by Plants in Karst …

to various studies, but basically, it concerns the characterization of the dissolution of soluble rock, the underlying mechanisms, and its interaction with the environment. For example, Atapour and Aftabi (2002) concluded the climatological, geochemical, and geomorphological processes of karstification that characterized regions containing soluble rocks and soluble soil and alluvium. Ford and Williams (2007) viewed karst as an open system composed of two closely integrated hydrological and geochemical subsystems. Dubois et al. (2014) stressed the two-stage process involved in karstification and related to hydrology. Karstification requires the presence of both soluble stones and water and carbonic acid (Harmand et al. 2017). When there are only CO2 and water, the solution has an equilibrium mixture of carbonic acid and bicarbonate and carbonate ions, which make up the DIC fraction (Liu et al. 2008): CO2 + H2 O ↔ H2 CO3 ↔ H+ + HCO− 3 ↔ 2H+ + CO2− 3 . The proportion of each species in the solution depends on pH. The equilibrium value of DIC for the CO2 –H2 O system is 0.018 mmol/L at an air temperature of 15 °C and CO2 partial pressure (pCO2 ) of 350 ppmv (Liu et al. 2008). However, when soluble rocks exist, such as limestone and dolostone, the chemical reaction can be shown as CaCO3 /MgCO3 + CO2 + H2 O ↔ Ca2+ /Mg2+ + 2 HCO− 3 . In this case, limestone and dolomite are dissolved away in surface and/ or underground water. The equilibrium value of the DIC for the CaCO3 –CO2 –H2 O system is approximately 65 times larger than that in the CO2 –H2 O system. The underlying mechanisms controlling the chemical dissolution of soluble rocks have been previously described with the theory of turbulent flow by Liu and Dreybrodt (1997). The turbulent flow is controlled by a diffusion boundary layer (DBL) adjacent to the surface of the mineral, through which mass transfer is influenced by molecular diffusion. Thus, they used a rotating disk technique to investigate the effect of DBL thickness and pCO2 of the solution on the dissolution rates of CaCO3 . The experiment showed that the conversion rate of CO2 into bicarbonate increased with pCO2 and DBL thickness. One of their subsequent works suggested that the equilibrium value of DIC for the CaCO3 –CO2 –H2 O system was 15.75 mmol/L in soil water when soil pCO2 reached 105 ppmv (Liu et al. 2008). Therefore, a high soil CO2 concentration is regarded as a driving force of the karstification process. However, when the two indexes continued to increase, the dissolution became rate limiting. They further tested the influence of carbonic anhydrase (CA), a zinc-containing metalloenzyme that catalyzes the interconversion of CO2 into bicarbonate with great efficiency (Moroney et al. 2001), on the dissolution of CaCO3 . The result exhibited an enhancement of CaCO3 dissolution by 1 order of magnitude (Liu and Dreybrodt 1997). In natural conditions, a variety of microorganisms, such as bacteria and fungi, reside on the surface of limestone (Ding and Lian 2008; Lian et al. 2011). These microbes contain extracellular CA and can speed up the corrosion of limestone and dolomite (Ford and Williams 2007; Lian et al. 2011). However, the CaCO3 –CO2 – H2 O system catalyzed by CA will rapidly reach a balance under natural conditions (Lian et al. 2011), which may hamper the continuous dissolution of carbonate. This is because the reaction can only occur when there is flowing water renewing the source of carbon acid (Dubois et al. 2014).

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5.4.2 Karst Hydrochemical and Hydrodynamic Processes The dissolution degree of karst bedrocks depends on a number of factors, of which the availability of water and its model of recharge play an important role (De Waele et al. 2009b). Therefore, studies on the coupling of hydrochemical and hydrodynamic processes are necessary for us to have a better understanding of karst functioning (Lastennet and Mudry 1997). The high strength of chemical dissolution triggered by the attack of CO2 -containing surface water initially results in small fissures, cracks (e.g., stratification joints, bedding planes), and faults (Harmand et al. 2017) and subsequently has a chance to widen the fractures depending on the dissolution rate (Dreybrodt 1992). This process differs greatly from that of sandsones, in which the latter allows the movement of water via lanima flow and has no significant effect on the storage capacity and transmission of groundwater (Ford and Williams 2007). In karst regions, the continuous dissolution driven by the penetration of surface water is likely to develop large groundwater networks over time and act as pathways for water flow (Silva et al. 2017b). The special characteristics of hydrological processes in karst systems thus form the well-known ‘dual hydrological structure’, namely, ground- and underground-linked drainage systems (Ford and Williams 2007). Under these circumstances, rainfall can be transported underground quickly through the fissures and fractures of rockstones, whereas little runoff simply flows across the surface (Peng and Wang 2012). Note that recharge of karst springs, such as storms or snow melt, usually causes a rapid response of the chemistry of water. This is often embodied in changes in the concentrations of dissolved CO2 and carbonates. For example, in temperate climates, the spring water was diluted by the addition of fresh water and exhibited a significant decrease in CO2 concentration, while in summer seasons, the CO2 concentration pronouncedly increased due to enhanced biological processes (e.g., respiration) in the soil and the epikarst (White 2015; Zhao et al. 2015). It was also concluded by Gulley et al. (2015) that the heterogeneous partial pressure of CO2 in karst waters was the dominant mechanism for the spatial distribution of chemical dissolution, cemnentation and macroporosity. Therefore, the variation in hydrochemistry on very short time scales imposes an important influence on karstification. In addition, karst can be regarded as a dissipative system, as the hydrodynamic enegy of water has the potential to remove dissolved carbonate and mechanically erode undissolved particles (Dubois et al. 2014).

5.4.3 Karst Landform The karst landform is a distinctive style of landscape developed on soluble rocks and related to efficient underground drainage (Waltham and Fookes 2003). This landform represents approximately 15% of the Earth’s continental area (Goldscheider et al. 2020), and the largest continuous karst areas are mainly located in Southwest China, Europe, and North America (Yuan 2001; Zhang et al. 2017; Goldscheider

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et al. 2020). Typical karst landforms are usually characterized by sinkholes (dolines), cones (fengcong), towers (fenglin), caves, long dry valleys, deep water tables, large springs, disappearing streams, tube structures, vaults, solution flutes, spongelike (honeycomb), and rillenkarren (Atapour and Aftabi 2002; Waltham and Fookes 2003; Ford and Williams 2007; Yang et al. 2019b). These landforms are the products of different hydrological conditions and developmental stages of karst. Furthermore, local geological features, such as fracture systems, sedimentary and fluvial processes (Atapour and Aftabi 2002; Silva et al. 2017b), and biological, chemical, and climatic conditions, also create suits of karstic characteristics with almost infinite variety (Ford and Williams 1989; De Waele et al. 2009a). For instance, regional climate imposes a strong influence on karst landforms by controlling local precipitation and temperature. For this reason, most mature karst occurs in wet tropical environments, whereas in temperate regions, the chemical weathering of limestone is reduced, and in arid and cold regions, it is largely subdued (Auler and Smart 2003; Waltham and Fookes 2003).

5.4.4 Karst Soil Soils are crumby and porous materials attached to the surface of the earth that are composed of solid (e.g., granular minerals, organic matter, and microorganisms), liquid (water), and gas (air) phases and support the growth of plants. The heterogeneity of karst systems, such as topography (e.g., slope and position), lithology (e.g., composition and structure), climate events (frequency and strength of rainfall), and human activity, influence the attribution and genesis of karst soils (Silva et al. 2017a). In karst regions, soil is generally formed from limestone (in situ dissolution) or derived materials (moved from its original site), as well as decomposition of residual bodies of plants, animals, and microbes under a certain time. The soil formation rate in karst regions with pure carbonate rocks is extremely slow, for example, ranging between 10 and 134.93 t km−2 yr−1 (Li et al. 2020). This is because the dissolution of highly soluble limestone retains a small amount of insoluble particles (residual). The soil depth in karst regions was commonly in the range of 20–40 cm (Yang et al. 2014), which affected community diversity and productivity (Liu et al. 2022). In addition, soil genesis is believed to be polygentic due to the coexistence of properties. When impurities present in limestone or the parent materials receive detritus from elsewhere, the soils in karst systems also contain quartz, clay minerals, and oxides. This phenomenon is also observed in karst regions. Soil properties determine resource availability (e.g., water and nutrients) and play an important role in biological processes and biogeochemical cycles in karst systems. The shallow and highly permeable soils in karst regions have low water content due to rapid infiltration of water through the developed fissures, cracks, and fractures and evaporation of water in the surface layer (Yang et al. 2019a). Studies have shown that in karst areas Climate change and human activity were the main factors influencing soil moisture (Wei et al. 2022). Vegetation types (e.g., grass, shrub,

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177

or tree) and topographic factors (e.g., position, degree or direction of slope) also exert direct or indirect influences on soil water content. Karst soils are characterized by naturally elevated fertility, with high contents of Ca2+ and Mg2+ ions, organic carbon and nitrogen, and high base saturation compared with nonkarst soil (Wen et al. 2016; Li et al. 2017; Silva et al. 2017a) but are limited by phosphorus (Chen et al. 2018). In addition, the high pH of karst soils usually reduces the availability of zinc and ferrous ions. However, most soil carbon is in the mineral-associated fraction, which is unavailable to microbes (Wen et al. 2017). This results in a reduced decomposition of organic matter by microorganisms. These indexes are regarded as the most critical factors reflecting the soil properties, which are strongly influenced by vegetation types (Lu et al. 2014). Limited resource availability has an influence on microbial processes (e.g., microbial respiration and enzymatic activity), which eventually constrains the dissolution of calcium carbonate. Further worthy of note, soil erosion is a serious environmental and ecological problem worldwide, particularly in karst regions, which impedes ecological and economic development in local areas (Gao and Wang 2019). Karst soil is often discontinuous and mixed with outcrops of rocks (Huang et al. 2021b). In natural conditions, soil erosion exhibits high spatial and temporal heterogeneity and is mainly influenced by land use type, slope, rainfall, and so on (Peng and Wang 2012; Gao and Wang 2019). In other cases, the amount of soil loss is very small in karst regions due to well-developed ground and underground drainage systems that enable quick infiltration of rainfall through limestone fissures and fractures (Peng and Wang 2012; Yang et al. 2019a). However, when the karst system is influenced by increasing human activities, for example, land use change (e.g., converting forest to farming and livestock), the strength of soil erosion is significantly enhanced (Gao and Wang 2019; Zhao and Hou 2019).

5.4.5 Karst Habitats and Vegetation Karst habitats are distinct from other topographies in their mountain-shaped carbonate bedrocks covered with thin soils, which are limited in water storage and have high contents of bicarbonate and calcium, high pH, and low nutrients (Yuan 2001; Wang et al. 2004; Cao et al. 2015). In karst areas, shallow soils, together with fissures (filled with surface soil driven by runoff), provide major habitats for plant survival and growth (Yan et al. 2019). Moreover, karst habitats are very fragile and vulnerable to increasing disturbances. Intensive land use, high frequency of drought and heavy precipitation run-off have accelerated the degradation of karst vegetation and even led to karst rocky desertification in past decades (Wang et al. 2004, 2019a; Day 2010; Long et al. 2014; Cao et al. 2015). For plants living in karst habitats, the water demand for maintaining basic functions is often restricted owing to recurring water scarcity (Yu et al. 2015). Drought stress can lead to a decline in stomatal opening and photosynthesis and downregulate metabolic processes (Liu et al. 2010, 2011; Wu et al. 2018). A prolonged drought will result in hydraulic failure and/or

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carbon starvation, increasing the risk of mortality (Sevanto et al. 2014; Rowland et al. 2015). On the other hand, the alkaline soil conditions in karst habitats are unfavorable for plant uptake of nutrients, such as phosphate, ferrous iron, and zinc (Msilini et al. 2009; Du et al. 2011). As a consequence, plants in karst habitats generally show slow growth rates and low productivity (Wang et al. 2004; Jiang et al. 2020). Of course, different vegetation types may differ in their physiological responses to harsh environments. For example, mosses and shrubs usually exhibit higher resistance to moderate or severe drought stress than tress. As shown by Liu et al. (2011), shrubs usually dominate the harsh habitas of karst regions with a higher photosynthetic rate and stomatal conductance, lower specific leaf area, greater intrinsic water use efficiency and thermal dissipation than tree species. Trees are more likely to live in favorable conditions with relatively high soil water content, and their height is less than 3 m in severely degraded habitats. This results in a very common phenomenon in which trees anchor in the fissures of bedrock by developed root systems (EstradaMedina et al. 2013; Nie et al. 2014), revealing more investment of carbohydrates in belowground growth to acquire more resources (Ni et al. 2015). Although plants can adopt tolerance and/or avoidance strategies to cope with environmental stress (Wang et al. 2017; Wu et al. 2018), this does not fully explain why some species grow well in karst habitats. For example, some tree species have heights exceeding 15 m and even form dense forests in subtropical or tropical karst regions (Li and Xiong 2021). It now appears that the survival and growth of karst plants require various strategies to maintain carbon assimilation and biomass production. Hence, other physiological processes of karst plants are suspected to play an important role in regulating their growth under dry and alkaline soil conditions.

5.5 Quantification of Bicarbonate Assimilation by Plants in Karst Habitats 5.5.1 What Can We Learn from Laboratory Experiments? Appropriate methods and models for quantification are critical to understanding plants’ use of soil DIC. According to numerous laboratory experiments, currently available methodological approaches include (i) applying high abundance 13 C labeling and then calculating the ratio of soil DIC fixed in specific tissues or organs to the total carbon gain. The total carbon gain includes soil DIC fixation, apparent photosynthesis measured by commercial infrared gas analyzers, and respiration (Ford et al. 2007; Shimono et al. 2019; Han et al. 2022); (ii) using two-source 13 C (closenatural abundance) labeling in combination with isotope mixing models to determine the contribution of root-derived DIC to leaf total photosynthesis (Wu and Xing 2012; Liang et al. 2020; Simkin et al. 2020; Wu and Wu 2022). These mainflow methods have been successfully used to quantify the actual amount of root-derived DIC fixed

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through or the proportion of DIC contributing to leaf photosynthesis. In field experiments, soil bicarbonate is naturally labeled, and its concentration is as high as those set in laboratory experiments, thus having the potential to be taken up by plant roots and ultimately assimilated in leaf chloroplasts. However, tracing the possibility of soil DIC contributing to leaf photosynthesis is not easy, particularly in natural fields. Both of the abovementioned methods have their own limitations. This experimental procedure of high-abundance 13 C labeling itself had difficulties and sometimes could be problematic. First, it usually requires a sealed environment to prevent the contamination of ambient air and the escape of labels (Ubierna et al. 2009). Second, 13 C labeling is often conducted as a form of solution irrigated to the root zones, which will inevitably change the soil conditions, such as the concentration of DIC and soil water content (also see Enoch and Olesen 1993). Third, the isotope signal of pulse labeling might not be detected due to tree size or wood anatomy (Stutz and Anderson 2021; Tarvainen et al. 2021). Finally, 13 C labeling is expensive, especially when dealing with numerous treatments and replicates. For two-source 13 C (or bidirectional) labeling, the procedures are so complex (replicate a pair of labeling groups and uniform plants) that they cannot be easily handled in field experiments. Specifically, the similar growth state and the systematic discrepancy of δ13 C between the two labeling groups cannot be guaranteed. For these reasons, a new approach needs to be developed to calculate the contribution of soil bicarbonate to leaf total photosynthesis in karst habitats. It has been shown that the natural abundance of 13 C has the potential to address a wide range of ecophysiological and biochemical questions (Göttlicher et al. 2006). For example, naturally occurring δ13 C signals in plant tissues integrate plant–environment interactions over long periods (Yang et al. 2015). In C3 plants, the δ13 C of leaves is mainly controlled by photosynthetic 13 C discrimination (Cernusak et al. 2013), which can be altered by stomatal control and the activity of the carboxylation enzyme Rubisco. Furthermore, when other carbon sources supply leaf photosynthesis, for instance, continuous uptake of DIC from the xylem sap of the host by the mistletoe or utilization of root-derived DIC by some plants (Marshall et al. 1994; Rao and Wu 2017b), the δ13 C of leaves can also be modified. In karst habitats, the high concentration and naturally labeled soil DIC provide us with an opportunity to search for relevant evidence rather than relying on manipunated methods, which could have many side effects on plants. This is because the δ13 C of soil-derived DIC or CO2 is much lower than that of atmospheric CO2 , which will have implications for the carbon isotope composition of photosynthates, depending on the amount of soil DIC or CO2 being photosynthetically assimilated (Brüggemann et al. 2011).

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5.5.2 Potential Carbon Sources for Leaf Photosynthesis in Karst Habitats In addition to atmospheric CO2 , soil CO2 (gas phase), soil DIC (liquid phase), and tissue-respired CO2 (e.g., root or stem respiration) are likely potential substrates for leaf photosynthesis. However, in karst soils, the concentration of soil CO2 is far less than that of root respiration. The concentration gradient from roots to soils will unlikely allow the fixation of soil CO2 by roots (Bloemen et al. 2016). As mentioned in Sect. 5.2.5, the amount of tissue-resipred CO2 entering the xylem sap is too large to be negligible in some species. In this case, the concentration and isotope signal of respiratory CO2 significantly contribute to both of those in the xylem sap, thus influencing the estimation of the contribution of root-derived DIC to leaf total photosynthesis. Furthermore, DIC fixation is also associated with the catalysis of PEPC (Ford et al. 2007; Msilini et al. 2009; Covarrubias and Rombolà 2013). This means that some root-derived DIC is transported to the shoots in the form of malic acid, which is later decarboxylated and then refixed in the carboxylation site of the chloroplast (Rombolà et al. 2005). Altogether, these potential pathways should be clarified and taken into account when quantifying their relative contributions to leaf total photosynthesis.

5.5.3 Theoretical Models to Estimate the Contribution of Soil DIC to Leaf Total Photosynthesis in Karst Habitats Gas-exchange measurement has enabled studies of atmospheric CO2 assimilation through photosynthesis at the leaf, branch or canopy level. In contrast, the internal refixation of tissue-respired and root-derived carbon cannot be directly measured using the conventional method. Furthermore, the limited ability to simultaneously measure multiple uses of carbon sources has constrained our ability to disentangle the participation of xylem-transported DIC from atmospheric CO2 in leaf photosynthesis (but see Stutz and Hanson 2019a, b). Therefore, tracking the fate of root-derived DIC in environment-related scenarios remains a challenge in field conditions. The possibility of xylem transport DIC consumed by leaf photosynthesis has been confirmed by abundant evidence from isotopic and flux techniques. However, this part of carbon gain is not included in gas-exchange measurements; thus, it will cause the underestimation of real photosynthesis (Levy et al. 1999). The combination of the stable isotope technique and gas-exchange system from the soil–plant-air system helps exploit the tissue-respired CO2 and root-derived DIC as possible carbon sources for leaf photosynthesis. Photosynthesis involves complicated gas-diffusion pathways, within which stomata play significant roles in determining how much CO2 diffuses into the intercellular space (i.e., C i /C a , the ratio of intercellular to ambient partial pressure of CO2 ), subsequently affecting leaf δ13 C. When atmospheric CO2 travels into the stomata, heavier 13 C is discriminated more

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than 12 C, and thus 12 CO2 moves faster than 13 CO2 . The robust linear relationship between δ13 C and C i /C a has been observed across various C3 plant species in normal soil conditions (Wingate et al. 2007; Lanigan et al. 2008; Cernusak et al. 2013), making it reliable to predict one value from the other measured value. Therefore, if soil has high DIC in karst habitats and if soil DIC has contributed to leaf photosynthesis, one would expect the observed/measured δ13 C to deviate from the predicted δ13 C from the measured C i /C a using a gas-exchange system. Typically, atmospheric CO2 is regarded as the sole substrate for leaf photosynthesis, and the carbon isotopic composition of photosynthates (δA ) can be predicted with the following equation (Farquhar et al. 1982): δA =

δa − △13 Ccom 1 + △13 Ccom /1000

(5.3)

where δa is the carbon isotope composition of atmospheric CO2 measured on the sampling date and △13 Ccom is the comprehensive discrimination against 13 C during the diffusion of CO2 through the boundary layer, stomatal and mesophyll conductance, as well as the effects of respiration and photorespiration. The current △13 Ccom model is shown as (Ubierna et al. 2018):  1 Ca − Ci a¯ 1−t Ca

  1+t Ci − Cc Cc ' αb Rd αb F b3 − e am + + − f 1−t Ca Ca αe Vc αf Vc

△13 Ccom =

(5.4)

where the variables are defined in Table 5.4, and t and a¯ are a ternary correction factor and the weighted fractionation for CO2 diffusion across the boundary layer and stomata, respectively, which are calculated as (Cernusak et al. 2013): αac E 2gac

(5.5)

ab (C a − Cs ) + as (C s − Ci ) Ca − Ci

(5.6)

t= a=

where αac = 1 + a¯ , E is the transpiration rate, and gac is the combined boundary layer and stomatal conductance to CO2 . In Eq. 5.4, C c is not directly measured by LI 6400, but it could be roughly estimated according to the intrinsic C c /C i relation. Ubierna and Farquhar (2014) assumed that C c /C i ranged from 0.7 to 0.9, von Caemmerer and Evans (1991) reported a value of 0.7 for C c /C i , and Warren et al. (2003) gave a value of 0.8. In this chapter, we use an intermediate value of 0.8 for C c /C i . V c and F are calculated as follows: Vc = A + Rd + F

(5.7)

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Table 5.4 List of symbols used Variable (units)

Definition

A (μmol m−2 s−1 )

Net photosynthetic rate for the fixed atmospheric CO2

ADIC (μmol m−2 s−1 )

Net photosynthetic rate for the fixed soil DIC

a¯ (‰)

Weighted 12 C/13 C fractionation for diffusion across the boundary layer and stomata in series, Eq. 5.6

ab (‰)

12 C/13 C

am (‰)

Summed 12 C/13 C fractionations during dissolution of CO2 and liquid-phase diffusion, am = 1.8‰

fractionation for CO2 diffusion in the boundary layer, ab = 2.9‰

as (‰)

12 C/13 C

fractionation for CO2 diffusion in air, as = 4.4‰

b3'

12 C/13 C

fractionation of RuBisCO, b3' = 29‰ (Evans and von Caemmerer

(‰)

2013)

C

Scaling constant for Γ * , c = 13.49

C a (μmol mol−1 )

CO2 partial pressure in the ambient air

C c (μmol mol−1 )

CO2 partial pressure in the chloroplast

C i (μmol mol−1 )

CO2 partial pressure in the intercellular air

C _R (mM)

Tissue-respired CO2 dissolved in the xylem sap

C s (μmol mol−1 )

CO2 partial pressure in the boundary layer

C _DIC (mM)

Concentration of DIC in the soil solution

E (mol m−2 s−1 )

Transpiration rate

e (‰)

12 C/13 C

fractionation for day respiration, e = 0 (Ubierna et al. 2019)

eb'

12 C/13 C

fractionation associated with the catalyzed dehydration of HCO− 3 ↔

(‰)

CO2 + H2 O, eb' = 9‰ at 25 °C (Mook et al. 1974) F (μmol m−2 s−1 )

Photorespiratory rate

f (‰)

12 C/13 C

f DIC_soil (%)

Fractional contribution of soil DIC to leaf total photosynthesis

fractionation during photorespiration, f = 11‰ (Tcherkez 2006)

f DIC_xylem (%) Fractional contribution of xylem DIC to δWSOM f s/x (%)

Ratio of the concentration of soil DIC to the xylem [CO2 ], f s/x = C _DIC / (C _DIC + C _R )

gac (mol m−2 s−1 )

Combined boundary layer and stomatal conductance to CO2 (continued)

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Table 5.4 (continued) Variable (units)

Definition

gs (mol m−2 s−1 )

Stomatal conductance to CO2

R (kJ J−1 mol−1 )

Molar gas constant, R = 0.008314 kJ J−1 mol−1

Rd (μmol m−2 s−1 )

Nonphotorespiratory CO2 released in the dark

t

Ternary correction coefficient, Eq. 5.5

T leaf (K)

Leaf temperature expressed as absolute temperature

V c (μmol m−2 s−1 )

RuBisCO carboxylation rate for the fixed atmospheric CO2

Vc' (μmol m−2 s−1 )

RuBisCO carboxylation rate for the fixed soil DIC

δ13 C or δ (‰) Carbon isotopic composition in the present study △13 C (‰)

12 C/13 C

ΔH (kJ K−1 mol−1 )

Energy of activation for Γ *, ΔH = 24.46 kJ K−1 mol−1

photosynthetic discrimination, Eq. 5.4, 5.11

Γ * (Pa)

CO2 compensation point in the absence of mitochondrial respiration

α ac

α ac = 1 + a¯

αb

α b = 1 + b3'

αe

αe = 1 + e

αf

αf = 1 + f

[CO2 * ] (mM)

Total quantity of gas-phase CO2 dissolved in water according to Henry’s law

where F=

[ ∗ (A + Rd ) Cc − [ ∗

(5.8)

where Rd is dark respiration and [ * is the CO2 compensation point in the absence of Rd , which is fitted using the following equation (Bernacchi et al. 2002): [∗ = e

△H ∗1000 (c− (R∗(273+T

) lea f )

(5.9)

where c, △H, and R are the scaling constant (13.49), energy of activation (24.46 kJ K−1 mol−1 ), and molar gas constant (0.008314 kJ J−1 mol−1 ), respectively. In addition to atmospheric CO2 , extensive studies have demonstrated that soil DIC can be taken up and photosynthetically fixed by plants. In this case, the carbon isotopic composition of photosynthate (δA' ) originating from the assimilation of soil DIC is given by:

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δA' =

δDIC − eb' − △13 CDIC 1+

△13 CDIC 1000

(5.10)

where δDIC is the carbon isotope composition of soil DIC collected on each sampling date, eb' is the discrimination related to the conversion between bicarbonate and CO2 , and eb' = 9‰ at 25 °C (Mook et al. 1974), accounting for the weak acidity of xylem sap in many species (Levy et al. 1999; Jackson et al. 2003; Jia and Davies 2007; Erda et al. 2014). Note that the fractionation factor of CO2 diffusing in the air is 1.004, whereas in the liquid phase, the fractionation is very low due to the slow diffusion rate of CO2 and thus can be ignored during long-distance transportation in the xylem sap. △13 CDIC is photosynthetic discrimination against 13 C combining the carboxylation of DIC and the effects of respiration and photorespiration, which is expressed as: △13 CDIC = b3' −

αb Rd αb F e − f αe Vc' αf Vc'

(5.11)

where Vc' is the rate of RuBisCO carboxylation for DIC and given by: Vc' = ADIC_ soil + Rd + F

(5.12)

where ADIC_soil is the net photosynthetic rate for soil DIC. Empirically, ADIC was less than 10% of A in many species. Here, we assumed ADIC_soil = A/10. Although this assumption introduces some error, the relatively low uptake of soil DIC will not have a substantial effect on the calculation of the contribution of DIC to leaf total photosynthesis. The expression of discrimination is valid for DIC (Eq. 5.11) because it does not need to go through the boundary layer and intercellular air space, which caused diffusive fractionation for atmospheric CO2 . The diffusion of DIC from leaf veins to the mesophyll chloroplast only occurs in the liquid phase, for example, in the cell membrane, cytosol, and chloroplast membrane. Under this circumstance, no large gradient of DIC concentration and diffusive discrimination are likely to come about, and thus, the carbon isotope discrimination of soil DIC becomes very simple, as shown in Eq. 5.11. Furthermore, if the major form of soil inorganic carbon is CO2 (gas phase), δDIC is calculated with the δ13 C of soil CO2 plus the combined discrimination of CO2 dissolution and diffusion in the water (am ). In reality, CO2 released from respiring cells in woody tissues (C_R ) is also dissolved in xylem sap (Teskey and McGuire 2007). C_R primarily ranges from 0.01 to 10.98 mM and varies with species, height, season, etc. (Levy et al. 1999; Teskey and McGuire 2007; Teskey et al. 2008). The mixture of respiratory CO2 and soil DIC makes the composition of xylem DIC complex. To disentangle the contribution of soil DIC from respiratory CO2 , we propose a ratio of the concentration of DIC (C_DIC ) or soil CO2 to the total DIC in the xylem sap, f s/x = C_DIC /(C_DIC + C_R ). In this chapter, we can directly measure C_DIC and adopt the mean value of 5 mM for C_R . Thus, the carbon isotope composition of the mixed DIC in the xylem sap (δDIC_xylem ) is calculated as follows:

5.6 Isotopic Evidence for Plant Use of Soil DIC in Karst Environments

δDIC_ xylem = f s/x (δDIC − eb' ) + (1 − f s/x )δR

185

(5.13)

where δR is the carbon isotope composition of tissue-respired CO2 dissolved in the xylem sap. We assume that the respiration of living tissues came from recently fixed carbon, such as water-soluble organic matter (WSOM), which will carry the enriched isotopic signal to roots due to postphotosynthetic fractionation (0.8 to −2‰; Bowling et al. 2008; Dubbert et al. 2012) and then result in −1 to −4‰ respiratory fractionation in roots (Gessler et al. 2007; Kodama et al. 2008; Dubbert et al. 2012; Bathellier et al. 2017). Here, we use a common value of −2‰ for postphotosynthetic fractionation and an intermediate value of −2‰ for respiratory fractionation; thus, δR is calculated as δR = δWSOM + 4‰ − am . Considering that both atmospheric CO2 and soil DIC could be used for leaf photosynthesis, the mixture of photosynthates from two substrates determines the value of δWSOM , which is expressed as a two-end-member mixing model:   ' δA · f DIC_ xylem + δA 1 − f DIC_ xylem = δWSOM

(5.14)

where f DIC_xylem is the proportion of xylem DIC contributing to δWSOM , δA' is recalculated with Eq. 5.10 but in which δDIC is replaced by δDIC_xylem . Finally, the contribution of soil DIC to leaf total photosynthesis ( f DIC_soil ) is calculated as follows: f DIC_ soil = f s/x · f DIC_ xylem

(5.15)

5.6 Isotopic Evidence for Plant Use of Soil DIC in Karst Environments 5.6.1 Utilization of Soil DIC by Plants in Karst Habitats: Essential Questions to Be Addressed Two issues need to be addressed in field trials: (1) whether the natural abundance of 13 C signals of soil DIC can be detected in leaves and (2) what the major carbon source supplying the plant roots is. A few studies show that even if the plants were irrigated with 13 CO2 or H13 CO− 3 solution, the carbon isotope signatures are too weak to be detected (Ford et al. 2007; Ubierna et al. 2009). This is probably due to a low concentration of the labeled species dissolved in the solution or DIC in normal soils (usually less than 1 mM), resulting in less CO2 or HCO− 3 being taken up by roots as the diffusion against the concentration gradient from roots to the soil is minimal (Bloemen et al. 2016). However, in karst habitats, the high concentration of soil DIC, reporting more than 5 mM (Rao and Wu 2022), makes it possible for them to be fixed by plants. In addition, labeling experiments usually have a single type of carbon source (Ford et al. 2007; Shimono et al. 2019; Tarvainen et al. 2021),

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while in field conditions, the major form of carbon sources depends on soil pH, humidity, temperature, root respiration, microbial activity, etc. (Shahabi et al. 2005; Poschenrieder et al. 2018).

5.6.2 Characteristics of Soil DIC and CO2 in Karst Habitats Rao and Wu (2022) concurrently measured the concentration of soil DIC and CO2 in the root zones of four tree species (Ligustrum lucidum, Broussonetia papyrifera, Platycarya longipes, and Zelkova serrata) and five shrub species (Viburnum dilatatum, Ampelopsis delavayana, Rosa cymosa, Zanthoxylum armatum, and Rubus biflorus) in karst habitats of Guizhou, southwest China. Their results showed that the average concentrations of DIC (or C_DIC ) in the root zones of four tree species were similar (approximately 9.2 mM), while in shrub species, they exhibited large variations, ranging from 8.1 mM in R. cymosa to 10.5 mM in Z. armatum (Fig. 5.5a). In contrast, the mean concentration of soil CO2 varied from 3919.91 ppm in P. longipes to 5279.83 ppm in B. papyrifera, higher than the mean value of 3454.42 ppm in the studied shrub species. According to Henry’s law, the partial pressure of CO2 over a solution is proportional to the concentration of CO2 in the solution. The calculated quantity of CO2 dissolved in soil water ([CO2 * ]) was in the range of 0.55 to 0.92 mM at pH 7 and 25 °C across nine species, much lower than the mean value of [CO2 * ] in xylem sap (or C_R ) investigated in many species. Thus, it was unlikely that soil CO2 could be fixed by plant roots due to a CO2 gradient from roots to the soil (Bloemen et al. 2016). In addition, the interspecies differences in the δ13 C of soil DIC and CO2 were both less than 2‰ (Fig. 5.5b). The mean values of soil DIC and CO2 among the nine species were −10‰ and −20‰, respectively. In fact, bicarbonate of DIC cannot be directly assimilated through leaf photosynthesis unless it is converted to CO2 . Considering the fractionation of interconversion (9‰ at 25 °C) between HCO− 3 and CO2 , soil DIC can be treated as a depleted 13 C label in comparison to that of atmospheric CO2 . Above all, given the high concentration of soil DIC and its negative isotope signal, the supply of DIC to roots has the potential to affect the carbon isotope composition of photosynthates in the long term, providing a significant amount of DIC uptake and delivery to shoots.

5.6.3 Disprepancy Between Predicted and Measured δ 13 C of Newly Formed Photosynthates Although the results of Rao and Wu (2022) suggested that soil DIC rather than soil CO2 was the main source supplying karst plants, direct evidence is lacking. They further examined whether soil DIC could affect the δ13 C of WSOM (newly formed photosynthates) in leaves. In addition, δ13 C of photosynthates can also be predicted

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Fig. 5.5 Concentrations (a) and δ13 C (b) of soil DIC and CO2 in the root zones of nine species across three altitudes. Reprinted with permission from Rao and Wu (2022). Copyright 2022 MDPI Publisher

with Eq. 5.3, which only accounts for the assimilation of atmospheric CO2 (δA ). In Rao and Wu (2022), δWSOM varied among nine species by approximately 3–4‰ and was lower than δA to different extents (Fig. 5.6), similar to their previous work (Rao and Wu 2017b). The discrepancy between δA and δWSOM (δA − δWSOM ) was species specific. The highest frequency (approximately 30%) of δA − δWSOM was located in the range of 1 to 2‰, while half of the discrepancy was distributed between 2 and 6‰. The isotope measurement error (less than 0.1‰) and diurnal variations in δWSOM (less than 1‰; Dubbert et al. 2012) were both insufficient to explain the systematic discrepancy. This means that there might be other carbon sources with very negative 13 C signals that participate in leaf photosynthesis. The photosynthates originating from xylem-transported DIC (the mixture of soil-derived DIC and tissue-respired CO2 ) carried more 13 C-depleted signal, which could be the only reliable reason for the systematically lower δWSOM than δA .

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Fig. 5.6 δ13 C of photosynthates determined by measuring the isotope signals of leaf WSOM (orange closed circle) or predicted with Eq. 5.3 that only accounted for the assimilation of atmospheric CO2 (green closed circle) of nine species. Reprinted with permission from Rao and Wu (2022). Copyright 2022 MDPI Publisher

5.6.4 Relationship Between δ A − δ WSOM and fDIC_soil The potential contribution of soil DIC ( f DIC_soil ) to leaf total photosynthesis could be quantified with a two-end-member mixing model (Eq. 5.14, 5.15). Again, in the study of Rao and Wu (2022), δA − δWSOM was itimately linked to f DIC_soil among all species (p < 0.001), with a coefficient of determination (R2 ) of 0.99 (Fig. 5.7). That is, the more soil DIC was involved in photosynthesis, the more δA deviated from δWSOM , and the larger the proportion of soil DIC contributed to leaf WSOM. This result further confirmed the speculation that soil DIC was involved in leaf photosynthesis in karst habitats and caused δWSOM to deviate from δA . Fig. 5.7 Relationship between (δA − δWSOM ) and f DIC_soil among nine species across three altitudes. Reprinted with permission from Rao and Wu (2022). Copyright 2022 MDPI Publisher

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5.7 Bicarbonate Assimilation in Karst Habitats 5.7.1 Species-Specific Induced Variation in Plant Use of Soil Bicarbonate in Karst Environments The field trial conducted by Rao and Wu (2022) is a timely work that demonstrates that soil DIC (mainly in the form of bicarbonate) assimilated by karst plants was analogous to those laboratory experiments (including the karst-simulation studies). As illustrated in Fig. 5.8, in tree species, the average values of f DIC_soil in L. lucidum and B. papyrifera were 8.93 and 9.54%, respectively, which were significantly higher than those in P. longipes and Z. serrata (both less than 2.2%). In contrast, the mean values of f DIC_soi within the shrub species varied from 2.48 to 9.99%, with the highest value in Z. armatum and the lowest value in R. biflorus. An assessment of f DIC_soil between two plant life forms, namely, trees and shrubs, showed no clear trend, indicating that the variation in f DIC_soil mainly resulted from interspecies differences. In most species, the values of f DIC_soil were comparable with those reported in laboratory experiments (Wu and Xing 2012; Hang and Wu 2016; Rao and Wu 2017b), suggesting that soil DIC is easily accessible to plants in karst habitats. In their study, nine species were all native, most of which were deciduous trees or shrubs. Although species differed in height, biomass, ages, etc., the measurements and sampling were all conducted with newly expanded leaves and within the root zones of the same soil layer. As leaves and roots control the uptake of atmospheric CO2 and soil DIC, respectively, the species-specific variation in f DIC_soil might be related to leaf gas exchange traits and soil conditions. Correlation analysis revealed that f DIC_soi correlated significantly with A, C i /C a , and WUE i . The f DIC_soil correlates positively with A (p = 0.003) and WUE i (p < 0.001) but negatively with C i /C a (p < 0.001). At first sight, it was surprising that f DIC_soil increased with an increase in

Fig. 5.8 Contribution of soil DIC to leaf total photosynthesis ( f DIC_soil ) in nine species. Capital letters on the error bar indicate significant differences (p < 0.05) among species. N = 4. Reprinted with permission from Rao and Wu (2022). Copyright 2022 MDPI Publisher

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A, which was inconsistent with results reported by their previous study (Rao et al. 2017b). For instance, when plants are confronted with moderate or severe water limitation, it drastically reduces A and gs (Liu et al. 2011) and thus increases the proportion of root or soil-derived DIC to support photosynthesis (Rao et al. 2017b; Wu and Wu 2022). The change in A was not proportional to that of ADIC_soil or E (Rao et al. 2017b), thus promoting the value of f DIC_soil arithmetically. In Rao and Wu (2022), the level of gs indicated mild drought stress in many species. Among these species, higher A corresponded to higher gs and E, which benefits the long-distance transport of DIC in the xylem sap (Bloemen et al. 2013; Stutz and Hanson 2019a). Similarly, a higher WUE i also implied a higher contribution of soil DIC to leaf photosynthesis, although the correlation between E and f DIC_soil was not significant. The negative correlation between f DIC_soil and C i /C a corresponded to the reverse change in f DIC_soil and the contribution of atmospheric CO2 to leaf total photosynthesis (1 − f DIC_soil ) because higher C i /C a usually suggested higher △13 Ccom (Eq. 5.4) and lower A and E (Cernusak et al. 2013). In addition, there was no significant correlation between f DIC_soil and C _DIC and δDIC . This could be explained by less variation of these indicators in comparison to that of leaf gas-exchange parameters among species.

5.7.2 Spatial Variability in Root-Derived Inorganic Carbon Utilization by Plants in Karst Habitats In karst habitats, altitude plays an important role in affecting the soil conditions and microclimate (Rao et al. 2017a), which determines the distribution of plant species (Zhang et al. 2013). Species with different life forms may vary in functional traits, such as root uptake and xylem transport of water and DIC, thus influencing the extent of soil DIC used by plants. In addition, the concentration of soil DIC may change with altitude, which probably acts on the proportion of soil DIC used by leaf photosynthesis. Rao and Wu (2022) investigated the effect of soil DIC at different altitudes on the leaf photosynthesis of three karst plants. In L. lucidum and A. delavayana, f DIC_soil tended to increase from lower to higher altitudes, whereas in V. dilatatum, the pattern was the opposite (Fig. 5.9). The impact of altitude on f DIC_soil was slightly confusing. Altitude did not directly affect f DIC_soil but influences the site-characteristic related microclimate and soil conditions (Zhang et al. 2013; Jucker et al. 2018). For example, altitude constrains the daily mean temperature and vapor pressure deficit (Jucker et al. 2018) and produces considerable variations in soil moisture and nutrient availability (Zhang et al. 2013). However, in the study of Rao and Wu (2022), the air temperature and relative humidity recorded by LI 6400 did not change linearly with altitude but exhibited a high degree of spatial heterogeneity. The reasons might be that the altitude gradient was not large enough, and some occasional factors, e.g., wind, shading, and vegetation coverage, could redistribute these resources, e.g., light and vapor (Burgess et al. 2016; Jucker et al. 2018). Their

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Fig. 5.9 f DIC_soil of one tree species (L. lucidum) and two shrub species (V. dilatatum and A. delavayana) in different altitudes. Capital letters on the error bar indicate significant differences (p < 0.05) among altitudes. N = 4. Reprinted with permission from Rao and Wu (2022). Copyright 2022 MDPI Publisher

previous study reported decreasing patterns for soil water content, organic matter, and some nutrients with altitude in the same area and same season of 2015 (Rao et al. 2017a). However, the increasing trend of f DIC_soil along the altitude gradient in L. lucidum and A. delavayana was opposite to that in V. dilatatum, implying that the differences in soil conditions could not solely explain the variation in f DIC_soil . Therefore, they speculated that all these variabilities were combined to influence the leaf gas exchange and subsequent estimation of f DIC_soil . Future studies may choose a large range of environmental gradients and exclude the influences of some occasional factors.

5.7.3 Temporal Variability in Root-Derived Inorganic Carbon Utilization by Plants in Karst Habitats In karst habitats, species-specific and altitude-induced variations in plant use of bicarbonate have been displayed in the study of Rao and Wu (2022); however, the response of root-derived bicarbonate assimilation to seasonal changes remains unknown. The latest research, also conducted by Rao et al. (2023), was specifically designed to address this question. To compare with their previous studies, two karst-adaptive tree species, P. longipes and C. acuminata, were chosen. Measurements were made on five sampling dates with evenly intervals (April 15, May 24, July 3, August 25, and October 12) throughout the growing season of 2017. The δ13 C of soil CO2 in the root zones of the two species was very sensitive to seasonal change, exhibiting more positive values at the beginning and end of the growing season and relatively negative values in the middle season. In comparison to soil CO2 , the δ13 C of soil DIC showed a declining trend across the growing season, indicating the enhancement of biological activity related

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to the weathering of carbonate rocks. The annual mean values were −8.41 and −7.64‰ for C. acuminata and P. longipes, respectively. At the study site, the δ13 C of atmospheric CO2 also varied with the growing season, ranging from −11.2 to 8.44‰. This was likely due to the seasonal variations in the combustion of fossil fuel. For δ13 C of leaf WSOM, its change pattern was similar to that of soil CO2 . The most negative δ13 C values of leaf WSOM were −30.06‰ for C. acuminata and −28.74‰ for P. longipes in July. Furthermore, the δ13 C of WSOM in P. longipes was larger than that of C. acuminata on the first three sampling dates (p < 0.01). The values of f DIC_soil in the two species fluctuated with the growing season, showing annual mean values of 15.96 ± 0.62% for C. acuminata and 16.23 ± 0.55% for P. longipes (Fig. 5.10a). The change patterns of f DIC_soil in the two species were similar on the first three sampling dates but differed in August. This led to the maximal f DIC_soil of the two species (20.90 ± 1.50% for C. acuminata and 18.68 ± 0.70% for P. longipes) occurring in the same month (May), while the minimal values occurred on different dates (12.00 ± 0.72% for C. acuminata in October and 10.90 ± 0.62% for P. longipes in July). Significant differences in f DIC_soil between the two species were found on the last three sampling dates. Additionally, f DIC_soil was transformed to the photosynthetic flux (ADIC_soil ), as shown in Fig. 5.10b. The seasonal variations in ADIC_soil were similar to those in f DIC_soil in the two species. The annual mean values of ADIC_soil were 0.99 ± 0.08 μmol m−2 s−1 and 1.29 ± 0.07 μmol m−2 s−1 for C. acuminata and P. longipes, respectively. A significant difference between C. acuminata and P. longipes was observed on August 25 and October 12 (p < 0.05). The annual mean values of f DIC_soil in the two species were comparable to the contribution of C. acuminata seedlings exposed to 10 mM HCO− 3 solution in combination with osmotic stress conditions in laboratory experiments (Rao and Wu 2017b) and in similar studies of other karst-adaptable species (Hang and Wu 2016; Wu and Xing 2012). The results indicated the great potential of soil DIC utilization by karstadaptable plants. In contrast, f DIC_soil is usually reported to be less than 2.5% across many species growing in normal soil conditions or hydroponic solution (Levy et al. 1999; Ford et al. 2007; Ubierna et al. 2009; Angert et al. 2012; Bloemen et al. 2013; Stutz and Hanson 2019a). This is because in many studies, the concentration of CO2 or DIC in soil or hydroponic solution is very low. Another reason is that most xylemtransported DIC is lost through transpiration or stem CO2 efflux under nonstress conditions, especially in conifer species (Ubierna et al. 2009; Powers and Marshall 2011; Stutz and Hanson 2019a; Tarvainen et al. 2021). The underlying reasons behind the seasonal change-induced variations in f DIC_soil in the two species lay with their physiological responses to environmental factors. It appeared likely that the high values of f DIC_soil occurred in drought and/or cold seasons, while the low values were shown in wet and/or warm seasons (Fig. 5.10a). This trend seemed not in line with expectations; for instance, the greater f DIC_soil should be in the wetter months. Soil DIC needs transpiration to travel from the roots to the leaves and at high enough rates and concentrations to not be lost entirely to radial diffusion in the stems (Stutz and Hanson 2019a). Nevertheless, f DIC_soil is a relative value determined by both A and ADIC_soil . A simple case was that when the season changed from May (dry spring) to July (wet summer), A increased by 36.48%,

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Fig. 5.10 Seasonal variations in (a) the contribution of soil DIC to leaf total photosynthesis ( f DIC_soil ) and (b) the net photosynthetic rate assimilated from soil DIC (ADIC_soil ) in C. acuminata (filled square and solid line) and P. longipes (open square and dash line), respectively. Values were mean ± 1 SE, N = 10

while E increased by 35.04% in C. acuminata. This resulted in a decline of f DIC_soil by 15.45% from May to July, suggesting that f DIC_soil was not only controlled by transpiration rate. The Pearson correlation analysis showed that f DIC_soil was closely related to A, E, C _DIC , and △13 CDIC in both species (p < 0.001). It is known that A is related to the contribution of atmospheric CO2 to leaf total photosynthesis (1 − f DIC_soil ), E is linked to the flow rate of xylem sap and thus the transport of root-derived DIC (Bloemen et al. 2013; Stutz and Hanson 2019a), C _DIC determines the total amount of xylem-transport DIC being photosynthetically fixed (Stutz and Hanson 2019a), and △13 CDIC is affected by the relative importance of respiration and photorespiration (Eq. 5.11), and finally integrated into the isotope mixing model and the calculation of f DIC_soil . Interestingly, the impacts of A and E on f DIC_soil were opposite between the two species, and it was difficult to find a rational explanation for this inconsistency. Furthermore, there were some other factors influencing the variations in f DIC_soil . For instance, δWSOM was significantly correlated with f DIC_soil in C. acuminata, whereas gs , C i /C a , and △13 Ccom all had a negative relationship with f DIC_soil in P. longipes (p < 0.05). This was because they were all related to the contribution of atmospheric CO2 (1 − f DIC_soil ). Above all, the combined effects of various processes and factors caused the different responses of f DIC_soil to seasonal changes in the two species. While the underlying mechanisms controlling the different

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responses of f DIC_soil to environmental factors were not fully elucidated, and the experiments of this study were only conducted in the growing season of 2017, the study still had some implications for f DIC_soil in the context of global climate change.

5.7.4 Uncertainties Associated with the Estimation of Contribution of Soil DIC Although the estimates of f DIC_soil for the karst-adaptable species are presented in the above two studies (Rao and Wu 2022; Rao et al. 2023), the actual behaviors of soil DIC in the soil–plant-air system may be more complicated. Some uncertainties remaining in the field trials may bias the estimation of f DIC_soil . First, in both Rao and Wu (2022) and Rao et al. (2023), a mean value of 5 mM was assigned for the C_R . However, species differ in C_R due to their different physiological statuses and anatomical characteristics. For example, soil DIC taken up by roots mixes with tissue-respired CO2 (C _R ) in the xylem sap (Teskey and McGuire 2007; Moore et al. 2008; Salomón et al. 2022), where C _R varies within species (Teskey et al. 2008; Werner and Gessler 2011), exhibites durinal or seasonal variations (Teskey and McGuire 2007; Moore et al. 2008; Werner and Gessler 2011; Bloemen et al. 2016), and becomes enriched with an increased distance to the stem base (Teskey and McGuire 2007). These factors cause spatial and temporal variations in the concentration (C_DIC + C_R ) and δ13 C (δDIC_xylem ) of the mixed [CO2 * ] in the xylem sap (Tarvainen et al. 2021), thereby affecting the estimation of f DIC_soil . Sensitivity analysis showed that a variation of 2.50–7.50 mM in C _R led to −3.37 to 5.76% of the change in f DIC_soil in the two species (Fig. 5.11c, d). This variation indicated that consideration of such a mixture is necessary. This uncertainty can be addressed by in situ measurement of xylem [CO2 * ] equipped with a nondispersive infrared CO2 sense (Aubrey and Teskey 2009; Bloemen et al. 2016) and sampling of xylem [CO2 * ] for determination of δ13 C (Ubierna et al. 2009; Powers and Marshall 2011; Tarvainen et al. 2021). Second, these studies assumed that respired CO2 came from newly assimilated carbon and used the value of δWSOM to infer the δ13 C of CO2 respired by living tissues (δR ). Recent studies have shown that vegetation pools respire carbon over a wide range of ages in many species (Gao et al. 2021; Hilman et al. 2021; Huang et al. 2021a; Sierra et al. 2022). For instance, the respired carbon in stems and roots is on average older than 1 year (Muhr et al. 2018; Hilman et al. 2021). When plants are exposed to stress, respiration almost completely relies on the old carbon (Huang et al. 2021b). Therefore, there is a high probability that the investigated species used reserved carbon for respiration under the condition of frequent water limitation in karst habitats. It has been shown that the δ13 C of reserved carbon had a close relationship with that of respired CO2 (Bathellier et al. 2017). Rao et al. (2023) showed that the maximum range of δ13 C in the storage carbon of some karstadaptable species was 4.13‰ during the growing season, which implied a similar or

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Fig. 5.11 Results for sensitivity analysis testing the effects of the total quantity of root-respired CO2 dissolved in the xylem sap (C _R ) and the ratio of chloroplast to the intercellular partial pressure of CO2 (C c /C i ) on the estimation of f DIC_soil in two species. Default input parameters are averaged values obtained from the literatures. During the sensitivity analysis, C c /C i is independently varied from 0.70 to 0.90 with a step change of 0.05 while C _R is held constant (a, C. acuminata; b, P. longipes). Similarly, C _R is independently varied from 2.50 to 7.50 mM with a step change of 1.25 mM while C c /C i is held constant (c, C. acuminata; d, P. longipes)

less variation range of δ13 C for respired CO2 in stems and roots. Some studies also showed that seasonal or annual variability of δ13 C in respired CO2 in stems and roots ranged within 2.0–4.5‰ (Kuptz et al. 2011; Brændholt et al. 2019; Diao et al. 2020). In the study of Rao and Wu (2022), taking L. lucidum as an example, a change of 4.5‰ in δ13 C of leaf WSOM only led to 0.5% of the change in f DIC_soil , indicating a limited influence of the source of carbon used for respiration on the quantification of plant utilization of soil DIC. Third, the determination of C c (CO2 partial pressure in the chloroplast) and involvement of DIC in the anaplerotic reactions also matter. A constant ratio of C c /C i = 0.8 was assumed for the calculation of △13 Ccom in Rao et al. (2023). The empirical value, reported to vary from 0.7 to 0.9, would result in changes of − 4.11 to 3.77% for f DIC_soil in the two species (Fig. 5.11a, b). Currently, the common

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approaches to calculate C c include gas exchange measurement (A-C i curve), chlorophyll fluorescence associated with gas exchange measurement, or photosynthetic carbon discrimination in combination with gas exchange measurement (Pons et al. 2009). Additionally, bicarbonate of soil DIC can be fixed by phosphoenolpyruvate carboxylase in roots and stems (Msilini et al. 2009), causing the accumulation of malate and involvement in anaplerotic fixation (Ford et al. 2007; Sagardoy et al. 2011). However, thus far, there are no data showing the magnitude of organic acids transported from roots to leaves. Collectively, even though some uncertainties remain in the quantification of f DIC_soil , the studies of Rao and Wu (2022) and Rao et al. (2023) provide a benchmark for the determination of f DIC_soil in natural habitats via a natural abundance of 13 C technique. Future works are expected to resolve these uncertainties and thus improve the estimation of f DIC_soil .

5.8 Eco-Physiological and Biogeochemical Significance of Bicarbonate Assimilation 5.8.1 Potential Strategy for Plants Adapting to Karst Environments Drought stress in karst habitats, as indicated by the low stomatal conductance of plants, exerts strong influence on the plant carbon gain from atmospheric CO2 (Liu et al. 2010; Rao and Wu 2017a). In the studies of Rao and Wu (2022) and Rao et al. (2023), the values of gs were very low in many species and throughout the growing season. For example, the highest gs was only 0.13 mol m−2 s−1 for P. longipes in July, which is close to the drought threshold value (0.1 mol m−2 s−1 ) of most angiosperm species (Flexas and Medrano 2002; Zhu et al. 2021). The low gs results in less CO2 available from the atmosphere for carboxylation, which may force plants to direct to other C sources, i.e., from the soil. Isotope evidence of the utilization of soil DIC as a carbon source for leaf photosynthesis (Rao and Wu 2022; Rao et al. 2023) emerges as an adaptation strategy in response to the drought-induced constraint on carbon gain. In comparison to the immediate uptake and carboxylation of atmospheric CO2 , the utilization of soil DIC is rather complicated and time-consuming. The transport of DIC in the xylem sap involves a long distance that relies on the bulk flow of xylem water (Bloemen et al. 2013; Stutz and Hanson 2019a; Salomón et al. 2022). Nevertheless, the advantage is that the xylem sap and stem tissues can build up pools for root-derived DIC (Salomón et al. 2022). It is analogous to the role of NSC that remobilizes and becomes depleted under drought stress (Hartmann and Trumbore 2016; Galiano et al. 2017). The pool size is determined by the length and diameter of the vessels (Teskey and McGuire 2007). Altogether, soil DIC can be continually supplied to the leaves but with a considerable time lag when photosynthesis starts. The estimate of f DIC_soil makes us conscious of the importance of soil DIC for maintaining the physiological functions of plants under the circumstance of water

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limitation. For example, the annual mean value of ADIC_soil was 0.99 μmol m−2 s−1 and 1.29 μmol m−2 s−1 for C. acuminata and P. longipes, respectively (Rao et al. 2023). This level is equivalent to the respiratory flux among many species. The assimilation of soil DIC, similar to NSC, replenishes a certain amount of carbohydrates to meet the basic requirements of living, such as growth, respiration, reproduction, osmotic adjustment, and defense (McDowell 2011; Hartmann and Trumbore 2016; Galiano et al. 2017), and thus relieves the crisis of carbon limitation to some degree during moderate or severe drought stress (Guo et al. 2021). Although the annual mean values of f DIC_soil were similar in the two species, P. longipes exhibited higher A, ADIC_soil , and water use efficiency than C. acuminata. It thus enabled P. longipes to acquire more carbon at a lower cost of water. This is probably one of the reasons that leads to P. longipes becoming one of the few pioneering plants in karst habitats. Above all, the studies of Rao and Wu (2022) and Rao et al. (2023) highlight the important role of soil DIC in modifying plant carbon gain in karst habitats, which is meaningful when considered over the life of a plant (Stutz and Hanson 2019a). Understanding the relationships between plants and environmental factors in karst forest ecosystems will also enable us to apply these findings in vegetation management strategies and restoration of forest communities (Zhang et al. 2013; Cao et al. 2015).

5.8.2 Coupling of Photosynthesis and Karstification and Its Impact on the Terrestrial Carbon Sink Karstification is a geological process occurring in nature that does not produce a net carbon sink. This is because the dissolution of soluble rocks and the precipitation of carbonates is a reversible reaction. However, the incorporation of bicarbonate into the biosphere, for example, photosynthetically fixed inorganic carbon by aquatic or terrestrial plants (Liu et al. 2008; Rao and Wu 2022), will definitely increase carbon sequestration in terrestrial ecosystems. This global-scale carbon flux, as previously recognized as “missing carbon” and not accounted for in the present carbon-cycle model (Liu et al. 2008), is becoming increasingly important, as it may serve as one of the key components for carbon neutralization (Wu and Wu 2022). The reason is that the dissolution of 1 mol of carbonates requires the consumption of 1 mol of CO2 , thereby producing 2 mol of bicarbonate. The assimilation of bicarbonate by plants results in the coupling of photosynthesis and karstification. This process will drive the continuous transport of inorganic carbon from both the lithosphere (e.g., CaCO3 or MgCO3 ) and atmosphere (e.g., CO2 ) or pedosphere (e.g., respiration) to the biosphere (e.g., aquaruc or terrestrial plants) (Fig. 5.12). In this case, the potential of terrestrial ecosystems acting as major carbon sinks for atmospheric CO2 should be recalculated. In a field monitoring experiment, Liu et al. (2008) estimated that aquatic photosynthesis could fix approximately 10.1% of the total anthropogenic CO2 emissions, accounting for 28.6% of the missing carbon sink. For terrestrial

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plants in karst ecosystems, root-derived bicarbonate assimilation usually constitutes an average of 10% of leaf total photosynthesis. As shown by Wu and Wu (2022), the carbon sink capacity of an 8-year woody plant with karstification–photosynthesis coupling of 10% will be twice that without coupling. The carbon sink capacity of a 10year woody plant with a karstification–photosynthesis coupling of 10% is 1.6 times that with a coupling of 5% (1.1010 /1.0510 ); thus, the karst carbon sink capacity is 3.2 times that of the latter. Altogether, the coupling of photosynthesis and karstification is still an important component for global carbon sinks concerning the difficulties in reducing carbon emissions worldwide. In the context of global change, the increasing CO2 level will likely enhance karstification and may also improve the proportion of bicarbonate contributing to photosynthesis.

Fig. 5.12 Carbonic anhydrase is pivotal in karstification–photosynthesis coupling. Carbonic anhydrase (CA) catalyzes the dissolution of carbonate rocks at the rock–soil interface, the hydration of CO2 in the soil solution and soil–atmosphere and soil–vegetation interface, the photosynthesis of plants, etc. Reprinted with permission from Wu and Wu (2022). Copyright 2022 MDPI Publisher

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5.9 Conclusions and Prospects This chapter focuses on whether soil DIC could be assimilated through leaf photosynthesis in karst habitats and whether this process, if taken place, varies between species and exhibits spatial–temporal heterogeneity. In fact, a large number of laboratory experiments have been conducted to elucidate the underlying mechanisms of root uptake of DIC, its long-distance transport in xylem sap, sources and fates, as well as the contribution to plant carbon gain or leaf total photosynthesis. Common approaches, such as high abundance 13 C labeling or bidirectional labeling, may perform well under laboratory controlling conditions but are not applicable to field trials. Therefore, a natural abundance tracer combined with process-based models is developed to solve these problems. This chapter adds to the growing pool of evidence that the δ13 C of recently formed photosynthates in leaves was altered by soil DIC. Large discrepancies between the measured and predicted δ13 C of newly formed photosynthates (δA − δWSOM ) are observed in both laboratory and field experiments. This systematic difference could not be explained by measurement errors, diurnal variations in δWSOM , or flaws in the theory of isotope mixing models but rather by the involvement of soil DIC in foliar photosynthesis. Some studies have confirmed that the contribution of soil DIC to leaf total photosynthesis displays species-specific and altitude-induced variations. Additionally, seasonal change also imposes a significant effect on this contribution, which is mainly related to gas exchange parameters. This chapter highlights the important role of soil DIC in photosynthetic carbon assimilation, which should not be neglected in the study of karst ecology. It also has implications for an investigation into the strategy regulating plant adaptation to adverse environments in karst habitats and its influence on the estimation of global carbon budgets. Future studies are expected to reveal how soil DIC is taken up by plant roots (direct evidence is needed) and to what extent other carbon sources (e.g., malic acids and tissue-respired CO2 ) affect the estimation of root-derived bicarbonate assimilation. In these cases, it requires multidisciplinary knowledge and the capability of using various technologies and instruments, such as gene encoding, isotope labeling, xylem sap monitoring, manipulation of gas-exchange systems and/or laser spectrometers in field trials.

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