rate of photosynthesis in aquatic plants

Sciencing_Icons_Science SCIENCE

Sciencing_icons_biology biology, sciencing_icons_cells cells, sciencing_icons_molecular molecular, sciencing_icons_microorganisms microorganisms, sciencing_icons_genetics genetics, sciencing_icons_human body human body, sciencing_icons_ecology ecology, sciencing_icons_chemistry chemistry, sciencing_icons_atomic & molecular structure atomic & molecular structure, sciencing_icons_bonds bonds, sciencing_icons_reactions reactions, sciencing_icons_stoichiometry stoichiometry, sciencing_icons_solutions solutions, sciencing_icons_acids & bases acids & bases, sciencing_icons_thermodynamics thermodynamics, sciencing_icons_organic chemistry organic chemistry, sciencing_icons_physics physics, sciencing_icons_fundamentals-physics fundamentals, sciencing_icons_electronics electronics, sciencing_icons_waves waves, sciencing_icons_energy energy, sciencing_icons_fluid fluid, sciencing_icons_astronomy astronomy, sciencing_icons_geology geology, sciencing_icons_fundamentals-geology fundamentals, sciencing_icons_minerals & rocks minerals & rocks, sciencing_icons_earth scructure earth structure, sciencing_icons_fossils fossils, sciencing_icons_natural disasters natural disasters, sciencing_icons_nature nature, sciencing_icons_ecosystems ecosystems, sciencing_icons_environment environment, sciencing_icons_insects insects, sciencing_icons_plants & mushrooms plants & mushrooms, sciencing_icons_animals animals, sciencing_icons_math math, sciencing_icons_arithmetic arithmetic, sciencing_icons_addition & subtraction addition & subtraction, sciencing_icons_multiplication & division multiplication & division, sciencing_icons_decimals decimals, sciencing_icons_fractions fractions, sciencing_icons_conversions conversions, sciencing_icons_algebra algebra, sciencing_icons_working with units working with units, sciencing_icons_equations & expressions equations & expressions, sciencing_icons_ratios & proportions ratios & proportions, sciencing_icons_inequalities inequalities, sciencing_icons_exponents & logarithms exponents & logarithms, sciencing_icons_factorization factorization, sciencing_icons_functions functions, sciencing_icons_linear equations linear equations, sciencing_icons_graphs graphs, sciencing_icons_quadratics quadratics, sciencing_icons_polynomials polynomials, sciencing_icons_geometry geometry, sciencing_icons_fundamentals-geometry fundamentals, sciencing_icons_cartesian cartesian, sciencing_icons_circles circles, sciencing_icons_solids solids, sciencing_icons_trigonometry trigonometry, sciencing_icons_probability-statistics probability & statistics, sciencing_icons_mean-median-mode mean/median/mode, sciencing_icons_independent-dependent variables independent/dependent variables, sciencing_icons_deviation deviation, sciencing_icons_correlation correlation, sciencing_icons_sampling sampling, sciencing_icons_distributions distributions, sciencing_icons_probability probability, sciencing_icons_calculus calculus, sciencing_icons_differentiation-integration differentiation/integration, sciencing_icons_application application, sciencing_icons_projects projects, sciencing_icons_news news.

• Share Tweet Email Print
• Home ⋅
• Science ⋅
• Nature ⋅
• Plants & Mushrooms

Photosynthesis in Aquatic Plants

Role of Photosynthesis in Nature

Plants are producers. Instead of consuming food to get energy, they make their own. During the process of photosynthesis, plants take in energy from sunlight and convert it into chemical energy stored in carbohydrates. Photosynthesis involves the same molecules and chemical reactions in land plants and aquatic plants. Floating plants photosynthesize much like plants that grow on land. However, the process presents more of a challenge for aquatic plants if they are fully submerged below the surface of the water.

Photosynthesis Basics

Leaves are the main site for photosynthesis. Leaves contain chloroplasts, which are the organelles in plant cells where photosynthesis occurs. Chloroplasts contain molecules of chlorophyll that absorb visible light, mainly in red and blue wavelengths. Only a few molecules of chlorophyll absorb green wavelengths. As a result, plants appear green because they reflect more green light than they absorb.

Plants use the sugar made during photosynthesis to fuel growth, development, reproduction and repair. The simple sugars produced in photosynthesis bond to from more complex starches such as cellulose that provide structure to plants. In addition to providing a food source for animals and other consumers, photosynthesis also removes carbon dioxide from the environment and replenishes oxygen.

Stages of Photosynthesis

The two stages of photosynthesis are the light dependent and light independent reactions. Light dependent reactions involve the absorption of sunlight and the breakdown of water molecules into oxygen gas, hydrogen ions and electrons. The goal of this stage is to capture light energy and transfer it to the electrons to make energized molecules such as ATP. Oxygen is a waste product of this stage of photosynthesis.

The second stage of photosynthesis, also known as the Calvin cycle, uses the energized molecules created in the first stage to split carbon dioxide molecules taken in from the plant’s environment. The breakdown of carbon dioxide and water molecules in the cell results in the formation of sugar molecules. Specifically, six molecules of carbon dioxide and six molecules of water yield one molecule of glucose, with six molecules of oxygen given off as a by-product.

Floating Plants

Aquatic plants may take in carbon dioxide from the air or water, depending on whether their leaves float or are under water. The leaves of floating plants, such as lotus and water lilies, get direct sunlight. These types of aquatic plants do not require special adaptations to perform photosynthesis. They can take in carbon dioxide from the air and release oxygen into the air. The exposed surfaces of the leaves have a waxy cuticle to mitigate water loss to the atmosphere, like terrestrial plants.

Obtaining Carbon Dioxide

Submerged plants, such as hornwort and sea grasses, use specific strategies to meet the challenges of conducting photosynthesis under water. Gases such as carbon dioxide diffuse much more slowly in water than in air. Plants that are fully submerged have greater difficulty obtaining the carbon dioxide they need. To help ameliorate this problem, underwater leaves lack a waxy coating because carbon dioxide is easier to absorb without this layer. Smaller leaves can more readily absorb carbon dioxide from the water, so submerged leaves maximize their surface to volume ratio. Some species supplement their carbon dioxide intake by extending a few leaves to the surface to absorb carbon dioxide from the air.

Absorbing Sunlight

Adequate sunlight is also hard to come by for submerged plant species. The amount of light energy absorbed by an underwater plant is less than the energy that is available to land plants. Particles in water such as silt, minerals, animal waste and other organic debris reduce the amount of light that enters the water. Chloroplasts in these plants are often situated on the surface of the leaf to maximize exposure to light. As depth below the surface increases, the amount of sunlight available to aquatic plants decreases. Some plant species have anatomical, cellular or biochemical adaptations that allow them to carry out photosynthesis successfully in deep or murky water despite the decreased availability of sunlight.

Other Aquatic Producers

Many organisms other than plants carry out the role of producer in aquatic ecosystems. Some forms of bacteria as well as algae and other protists perform photosynthesis. Colonies of single-celled algae work together to form the macroalga kelp, commonly known as seaweed.

Related Articles

Is algae a decomposer, a scavenger or a producer, what is reduced & oxidized in photosynthesis, key differences between c3, c4 and cam photosynthesis, the three stages of photosynthesis, why is water important to photosynthesis, how do plant cells obtain energy, leaf cell structure, what provides electrons for the light reactions, where does photosynthesis occur in mosses, describe what a photosystem does for photosynthesis, in what type of habitat would you find a protist, how do plants store energy during photosynthesis, what is the relationship between co2 & oxygen in photosynthesis, importance of pigments in photosynthesis, organelles involved in photosynthesis, what is the role of pigments in photosynthesis, explain photosynthesis, how do stomata work in photosynthesis, what is the role of carotenoids in photosynthesis.

• University of Florida Center for Aquatic and Invasive Plants: Photosynthesis
• University of Hawai’i- Exploring Our Fluid Earth: What are Aquatic Plants and Algae
• University of New Hampshire Phycokey: Aquatic Macrophytes

About the Author

A.P. Mentzer graduated from Rutgers University with degrees in Anthropology and Biological Sciences. She worked as a researcher and analyst in the biotech industry and a science editor for an educational publishing company prior to her career as a freelance writer and editor. Alissa enjoys writing about life science and medical topics, as well as science activities for children

Find Your Next Great Science Fair Project! GO

• Skip to primary navigation
• Skip to main content
• Skip to footer

Biology Wise

Photosynthesis in Aquatic Plants

Both terrestrial plants and water plants photosynthesize with the help of light energy to make carbohydrates. Photosynthesis in aquatic plants takes place in the same way as the land plants undergo to produce foods. Read on to know more about how photosynthesis takes place in aquatic plants.

Like it? Share it!

Ability to perform photosynthesis is the main distinguishing feature between green plants and other organisms on Earth. In this chemical process, carbon dioxide and water are combined in presence of light energy to produce carbohydrate and other byproducts. This process of converting carbon dioxide to glucose with the help of radiant energy is observed in cyanobacteria (blue-green algae), some types of algae and all green plants, irrespective of the growing environment.

Photosynthesis in Aquatic Plants and Land Plants

So, is there any difference between photosynthesis in land plants and aquatic plants? Well, the process of producing food with the help of light energy remains the same for both aquatic and land plants. In addition to light, they require the basic raw materials – carbon dioxide (CO 2 ) and water (H 2 O) for synthesis of glucose (C 6 H 12 O 6 ). What is special about food production by plants under water is, deriving these raw materials and light energy from their immediate environment.

The balanced equation of photosynthesis is represented as: 6CO 2 + 12H 2 O + Light → C 6 H 12 O 6 + 6O 2 + 6H 2 O . In case of land plants, the required gases and light energy are available easily. They absorb carbon dioxide from atmospheric air through their stomatal openings (present in upper and lower side of leaves), water from the soil through their root system, and last but not the least, radiant energy from sunlight. Hence, land plants undergo photosynthesis naturally without any special adaptations.

How do Aquatic Plants Photosynthesize?

Since water is available in more than sufficient amounts, the major challenge is to obtain carbon dioxide and light. For the same, majority of these plants show adaptations in some way or the other. Say for example, water lily (lotus) bear floating leaves with waxy surface, which have extra long petioles. The leaves on the surface of water gather light and carbon dioxide for photosynthesis, while the waxy cuticle prevents excess water absorption from the surrounding environment. It also helps in cleaning process and preventing entry of microbes.

Lotus is just one instance, and many water plants have the same feature, i.e., sending their roots at the bottom of the water body for support and leaves protruding at the surface of water. So, absorption of carbon dioxide and light is not an issue for aquatic plants having floating leaves. But, what about the remaining plants that bear submerged leaves? In this aspect, carbon dioxide is obtained from water, which is released during respiration by fish. Also, decomposition of organic matter that takes place in the water ecosystem contributes to carbon dioxide production.

Even though the rate of carbon dioxide dissolution in water is exceptionally low, traces of dissolved carbon dioxide are available to the submerged aquatic plants. As for light requirement, sunlight pass through water is harvested by plants for photosynthesis process. This is specially true for plants that thrive in shallow waters. And for the plant species found in deeper waters, they are adapted to grow under low light conditions. Of course, the rate of photosynthesis is very slow for these plants, and they rely on the dim radiation for manufacturing their own food.

Thus, aquatic plants obtain carbon dioxide and sunlight for photosynthesis process. Last but not the least, availability of usable water is not an issue for freshwater plants for photosynthesis. While for marine plants, they are adapted with waxy stems and leaves. This aids in absorbing water, while preventing the entry of salt to their system. In addition, some marine plants have specialized features to remove salt as soon as possible. All these processes help in regulating the osmotic balance, which otherwise will cause leaching of water and desiccation of plants.

This way, aquatic plants undergo photosynthesis under water. The products of photosynthesis in aquatic plants, basically carbohydrate and oxygen, are used by other organisms living in the same biotic community. And just like animals, plants do require oxygen, but in small amounts. This is obtained from the oxygen released at the time of photosynthesis.

Get Updates Right to Your Inbox

Privacy overview.

• Architecture and Design
• Asian and Pacific Studies
• Business and Economics
• Classical and Ancient Near Eastern Studies
• Computer Sciences
• Cultural Studies
• Engineering
• General Interest
• Geosciences
• Industrial Chemistry
• Islamic and Middle Eastern Studies
• Jewish Studies
• Library and Information Science, Book Studies
• Life Sciences
• Linguistics and Semiotics
• Literary Studies
• Materials Sciences
• Mathematics
• Social Sciences
• Sports and Recreation
• Theology and Religion
• Publish your article
• The role of authors
• Promoting your article
• Abstracting & indexing
• Publishing Ethics
• Why publish with De Gruyter
• How to publish with De Gruyter
• Our book series
• Our subject areas
• Your digital product at De Gruyter
• Contribute to our reference works
• Product information
• Tools & resources
• Product Information
• Promotional Materials
• Orders and Inquiries
• FAQ for Library Suppliers and Book Sellers
• Repository Policy
• Free access policy
• Open Access agreements
• Database portals
• For Authors
• Customer service
• People + Culture
• Journal Management
• How to join us
• Working at De Gruyter
• Mission & Vision
• De Gruyter Foundation
• De Gruyter Ebound
• Our Responsibility
• Partner publishers

Your purchase has been completed. Your documents are now available to view.

Aquatic Photosynthesis

Second edition.

• Paul G. Falkowski and John A. Raven
• X / Twitter

Please login or register with De Gruyter to order this product.

• Language: English
• Publisher: Princeton University Press
• Copyright year: 2007
• Edition: Second
• Audience: Professional and scholarly;College/higher education;
• Main content: 488
• Other: 8 color plates. 10 halftones. 145 line illus. 22 tables.
• Published: October 31, 2013
• ISBN: 9781400849727

An official website of the United States government

The .gov means it’s official. Federal government websites often end in .gov or .mil. Before sharing sensitive information, make sure you’re on a federal government site.

The site is secure. The https:// ensures that you are connecting to the official website and that any information you provide is encrypted and transmitted securely.

• Publications
• Account settings

Preview improvements coming to the PMC website in October 2024. Learn More or Try it out now .

• Advanced Search
• Journal List
• Portland Press Opt2Pay

Emerging approaches to measure photosynthesis from the leaf to the ecosystem

Matthew h. siebers.

1 United States Department of Agriculture, Global Change and Photosynthesis Research Unit, Agricultural Research Service, Urbana, IL 61801, U.S.A.

2 Carl R. Woese Institute for Genomic Biology, University of Illinois at Urbana-Champaign, Urbana, IL 61801, U.S.A.

3 Departments of Plant Biology and Crop Sciences, University of Illinois at Urbana-Champaign, Urbana, IL 61801, U.S.A.

Nuria Gomez-Casanovas

Katherine meacham-hensold, caitlin e. moore.

4 Center for Advanced Bioenergy and Bioproducts Innovation, University of Illinois at Urbana-Champaign, Urbana, IL 61801, U.S.A.

5 Institute for Sustainability, Energy & Environment, University of Illinois at Urbana-Champaign, Urbana, IL 61801, U.S.A.

6 School of Agriculture and Environment, The University of Western Australia, Crawley, WA 6009, Australia

Carl J. Bernacchi

Measuring photosynthesis is critical for quantifying and modeling leaf to regional scale productivity of managed and natural ecosystems. This review explores existing and novel advances in photosynthesis measurements that are certain to provide innovative directions in plant science research. First, we address gas exchange approaches from leaf to ecosystem scales. Leaf level gas exchange is a mature method but recent improvements to the user interface and environmental controls of commercial systems have resulted in faster and higher quality data collection. Canopy chamber and micrometeorological methods have also become more standardized tools and have an advanced understanding of ecosystem functioning under a changing environment and through long time series data coupled with community data sharing. Second, we review proximal and remote sensing approaches to measure photosynthesis, including hyperspectral reflectance- and fluorescence-based techniques. These techniques have long been used with aircraft and orbiting satellites, but lower-cost sensors and improved statistical analyses are allowing these techniques to become applicable at smaller scales to quantify changes in the underlying biochemistry of photosynthesis. Within the past decade measurements of chlorophyll fluorescence from earth-orbiting satellites have measured Solar Induced Fluorescence (SIF) enabling estimates of global ecosystem productivity. Finally, we highlight that stronger interactions of scientists across disciplines will benefit our capacity to accurately estimate productivity at regional and global scales. Applying the multiple techniques outlined in this review at scales from the leaf to the globe are likely to advance understanding of plant functioning from the organelle to the ecosystem.

Introduction

The terrestrial biosphere consists of an assemblage of diverse ecosystems. Its complexity is illustrated with a diversity of plants with distinct canopy structures subject to changing environmental conditions. Life on earth relies on the energy captured by these ecosystems through photosynthesis, which accounts for the single largest flux associated with the global carbon cycle [ 1 ]. Photosynthesis varies among plant functional types (e.g. C3 vs. C4) and over a wide range of spatial and temporal scales associated with changes in light, temperature, water and nutrients [ 2 , 3 ]. Global climate change driven by anthropogenic activities is having profound impacts on terrestrial ecosystems, with global temperatures rising faster than worst-case predictions [ 4 ]. Increasing agricultural demands associated with a growing population requires a doubling of crop yields by 2050 to keep up with demands [ 5 ], yet current rates of improvement fall short of this goal [ 6 , 7 ], which is likely to suffer with continued global warming [ 8–11 ].

Photosynthesis is a highly complex and relatively inefficient process, yet it is a critical component of the biosphere. Understanding photosynthetic responses over a range of spatial and temporal scales is needed to understand current and to predict future global carbon cycling. This understanding will also lead to improving photosynthesis, which can lead to higher productivity to meet growing agricultural demands [ 12 , 13 ]. These goals can only be achieved through the ability to measure photosynthesis over time and space, yet photosynthesis is difficult to measure directly. This is due to the multiple processes that are represented by the exchange of CO 2 between plants/ecosystems and the surrounding air. For example, at the leaf scale CO 2 is removed from the air by photosynthesis but this is partially countered by photorespiration and respiration, both of which release CO 2 [ 14 , 15 ]. The combined fluxes of these three processes represents net carbon assimilation ( A ) and partitioning this net flux into the component fluxes is challenging [ 16 ]. Scaling beyond the leaf only presents additional challenges. At canopy or ecosystem scales, respiration from non-photosynthetic tissues and heterotrophic organisms also release CO 2 , which combined with A provide measures of Net Ecosystem Exchange (NEE; Table 1 ). In this review, we outline the current and emerging approaches to measure photosynthesis at multiple scales and address the challenges and opportunities at each scale ( Figure 1 ). We begin with a focus on the well-established and widely used gas exchange techniques and follow with more recent approaches made available through recent technological advances.

(A) Leaf-level gas exchange with one measured representative photosynthetic CO 2 response curve. (B) Canopy photosynthesis chamber situated over a soybean field with representative diurnal Net Ecosystem Productivity (NEP) data (Image Credit: Anthony DiGrado). (C) Ecosystem-scale eddy covariance system situated over sorghum with representative Net Ecosystem Exchange (NEE; negative values signify downward flux from atmosphere toward land surface) partitioned into Gross Primary Productivity (GPP) and Ecosystem Respiration (ER). (D) leaf hyperspectral point sensor being used on the model crop tobacco and representative spectral reflectance measurements. (E) A hyperspectral imaging sensor measuring plots of the model crop tobacco and an example hypercube showing the visible surface and spectral information for each pixel with depth of image. (F) aircraft and satellite depicted over the earth surface and a map of GPP (public domain image courtesy of GeoEye/NASA SeaWIFS project). Other than where indicated, all images were taken by authors.

Table 1

In general, photosynthesis is defined as the process why which plants capture light energy and atmospheric CO 2 to synthesize complex carbohydrates. Photosynthesis supports the production of food, fiber, wood, grain fed to livestock, and fuel for humanity and regulates the concentration of CO 2 in the atmosphere. Quantifying global terrestrial photosynthesis is essential to understanding the global CO 2 cycle in a changing environment and the climate system.

Gas exchange

The fundamentals of gas exchange at any scale are relatively similar and require the ability to measure gas concentrations in air surrounding and the flow rate in which the air interacts with photosynthetic tissue. In addition to these measurements, numerous assumptions, corrections, and parameterizations are required to fully exploit the power of this technique [ 16 , 17 ]. Gas exchange methods have been applied at scales ranging from the organelle (e.g. [ 18 , 19 ]) to the whole ecosystem/region [ 20 ] to provide a basic understanding of how leaves, plants, and ecosystems function and respond to their environment ( Figure 1 ). Historically, gas exchange measurements were limited to enclosed sampling chambers, ranging from sections of leaves to whole plant canopies, where the rate of CO 2 exchange was measured over time. With the advent of micrometeorological techniques, gas exchange measurements at large scales (e.g. whole ecosystems) were developed that removed the need for enclosures ( Table 2 ). Despite errors, uncertainties and challenges associated with gas exchange, the various techniques are the current ‘gold standard’ by which emerging techniques are compared. This section provides an overview of gas exchange measurements at the leaf to ecosystem scales as a baseline in the understanding of emerging techniques.

Table 2

The global eddy covariance network, called FLUXNET ( https://fluxnet.org/about/ ), includes measurement sites linked across regional networks in North, Central and South America, Europe, Asia, Africa, and Australia.

Leaf scale gas exchange

Knowledge of leaf photosynthetic physiology stems from the development and application of leaf-level gas exchange systems [ 16 ]. Gas exchange technology has matured to the point where commercial gas exchange systems are widely available from many vendors. In addition to providing the key variables necessary to assess leaf scale carbon assimilation, these systems now provide the opportunity to precisely control the environmental conditions surrounding the photosynthetic tissue and to measure more than just carbon assimilation, including but not limited to transpiration, intercellular CO 2 concentrations, and stomatal conductance. Gas exchange techniques have been used for decades and most recent advancements have focused on improvements in accuracy, precision, usability, environmental control, and reduction in time to stable measurements. Despite the ease with which leaf level gas exchange can be measured, the importance of understanding gas exchange theory to ensure proper measurement and analysis cannot be overstated.

Gas exchange systems are the most commonly utilized technique for leaf scale photosynthetic measurements. While systems provide measures of A , various techniques can be applied to separate fluxes of photosynthesis, photorespiration, and respiration. However, many challenges exist with gas exchange that limit the wide application of the technique. These include cost, usability, data processing requirements, and time needed for ensuring quality measurements. Off-the-shelf gas exchange systems cost tens of thousands of dollars and require frequent maintenance that challenges their widespread use. Most gas exchange systems limit the area of measurement to, at most, several cm 2 , which presents issues related to scaling photosynthesis beyond a small section of one leaf. Typical measurements of in situ gas exchange require a minimum of 2–3 min to allow for both the system and the leaf to stabilize. Using these systems to measure beyond a simple survey of gas exchange, for example to measure light response or CO 2 response curves of A , requires substantially more time for each leaf. Recent techniques that exploit improved instrument precision can reduce the time for some measurements but generally at the expense of accuracy, and often require more advanced post-processing [ 21 ].

Canopy and ecosystem scale gas exchange

Scaling gas exchange measurements to the canopy or whole ecosystem presents significantly more challenges than at the leaf level, yet there are also more options ( Table 2 ). Canopy chambers work in much the same way as leaf chambers, although at a larger scale. The general principle follows that of leaf-level measurements, although chambers are required to be much larger to encompass multiple plants and the potential is greater for errors associated with leaks or pressure fluctuations [ 17 ]. Canopy chambers have been extensively used to measure CO 2 fluxes for a wide range of vegetation types and their strengths lie in their ability to address small-scale spatial variability ( Table 2 ). Furthermore, canopy chambers have been used both as a measurement and treatment system in global change studies to impose treatments as open-top chambers and acting as sample chambers when enclosed (e.g. [ 22 ]). Canopy chambers, however, can be limited in sampling frequency and spatial integration ( Table 2 ) while also having a profound impact on the canopy microclimate.

Micrometeorological approaches to gas exchange lack the need for chambers but require large spatial areas (>4 Ha) and a sensor suite that can measure the upward/downward movement of air coupled with the gas concentrations in the air [ 20 ]. The dominant micrometeorological technique, eddy covariance (EC), provides near-continuous measurements of NEE integrated over large spatial areas, called the flux footprint, with minimal disturbance ( Table 2 ) [ 20 ]. Air flow over a canopy consists of numerous rotating eddies. Measuring the speed and CO 2 concentrations of the eddies moving air upward and downward, provides the basic data needed to calculate fluxes of the footprint, which varies with wind speed and direction [ 23 ]. EC requires several important considerations to ensure the NEE data are robust and reliable [ 24 ], including ensuring sufficient atmospheric turbulence [ 23 ], applying corrections to exclude data fluxes extending beyond the area of measurements [ 25 , 26 ], and ensuring all measured fluxes follow the laws of thermodynamics [ 27 , 28 ] ( Table 3 ). Because of inevitable gaps in data collection associated with field instrumentation, gap filling strategies are used to complete the time-series of flux data ( Table 4 ). In addition to NEE, EC can apply to any measurable component of the atmosphere provided high temporal resolution sensors (≥10 Hz) exist (e.g. water vapor, methane, etc.). A global EC flux network, called FLUXNET, provides data from over 900 sites globally, allowing for a link between ecosystem and global NEE. This network provides unprecedented insights into environmental and biological drivers of ecosystem NEE [ 3 , 20 , 29–33 ]. Among other purposes, the long-term measurements of NEE from this network have improved understanding of ecosystem responses to climate and land-use change [ 34 ], and the data are essential to validate remote sensing and modeling products that scale to regions and the globe [ 35 , 36 ].

Table 3

Some of these challenges include ensuring sufficient atmospheric turbulent conditions are met [ 23 ], applying footprint corrections to exclude data when a significant portion of fluxes occur outside the ecosystem region of interest [ 25 , 26 ], and quantifying energy balance closure at the site [ 27 , 28 ]. Improving the robustness of NEE estimates from flux towers is an area of active research in the flux community, and one which will lead to greater understanding of ecosystem photosynthesis across a diversity of biomes.

Table 4

Good reliability of annual sum of the net CO 2 exchange refers to methods that ranked the best based on a several statistical metrics to predict annual fluxes as reported in References [ 122 , 123 ]. These statistical metrics include Root Mean Square Error, Bias Error and the annual CO 2 flux sum among others and were evaluated by comparing the filled NEE data with the observed values.

Whether using chamber-based or micrometeorological approaches, measured NEE provides an opportunity to explore changes in ecosystem-scale gas exchange at high temporal frequency. Photosynthesis at the ecosystem scale is generally defined as gross primary productivity (GPP), which is only one component of NEE. GPP is derived as the difference between measured NEE and modeled ecosystem respiration (ER; Table 5 ). Obtaining GPP from NEE involves modeling ER using temperature and light response functions; a process typically referred to as flux partitioning [ 24 , 32 , 37 , 38 ]. Flux partitioning allows for the investigation over time of GPP and ER in response to a variety of conditions [ 39–41 ]. A challenge with flux partitioning is introduced by the inhibitory effect of light on leaf respiration rates, known as the Kok effect [ 42 ]. In the light, autotrophic respiration can be significantly lower than at night resulting in GPP estimation errors when ignored [ 43 ].

Table 5

Both methods assume that any difference between daytime and nighttime Reco is due to temperature alone.

Recent micrometeorological approaches have attempted to measure GPP using a sulfur-containing analog of CO 2 , carbonyl sulfide (COS) that acts as natural ‘tracer’ molecule for GPP. This molecule enters a leaf in the same manner as CO 2 and is broken down by the enzyme carbonic anhydrase. Because of this, COS ‘uptake’ should scale with GPP, removing the need for partitioning NEE into the GPP and respiration components [ 44 ]. Studies using this method are showing promising insights with GPP estimated using CO 2 vs. COS measurements agreeing within 15% in forests and crops [ 45 ]. Another study that investigated variability in COS uptake and release in forests found agreement to within 3.5% between the two methods when GPP was high [ 46 ]. These results suggest an opportunity to use indirect methods for assessing GPP at larger scales, although recent work also suggests that photosynthetic tissues are not the only sink for COS [ 46–48 ].

Remote and proximal sensing

Obtaining photosynthetic carbon uptake measurements using gas exchange systems is laborious resulting in efforts to replace this technique with other high-throughput methods. There exists a rapid growth in plant phenotyping greenhouses with the goal of automated measurement capabilities [ 49 ] at scales ranging from leaf to globe ( Figure 1 ). Even with the most modern technologies, direct monitoring of leaf or plant level gas exchange would require substantial effort and resources. Thus, there are emerging technologies that provide means to infer plant responses to their growth environments that overcome the limitation of gas exchange [ 50–53 ]. Commercial sensors are available that provide information about plant canopy architecture and volume, which is important to infer growth over time [ 54 ], yet disentangling the underlying factors that lead to this growth requires physiological understanding. In the field, plot-level estimations of photosynthetic traits have been successfully estimated using a variety of platforms [ 55–57 ]. However, there needs to be improvements to the precision, accuracy, repeatability, and data pipeline before we can use these methods to estimate photosynthesis. Nonetheless, these new methods have a large potential impact on leaf to canopy understanding of plant physiology, ecosystem functioning and improving breeding efforts to maximize crop yields. In this section, we will discuss emerging technologies to monitor photosynthesis using spectral reflectance or fluorescence techniques. We will first outline the tools used for these approaches followed by a description of how these tools are being used.

Hyperspectral approaches to measure photosynthesis

Hyperspectral analysis is a non-destructive means of analysis that uses light reflected from vegetation to infer leaf, plant, canopy, or ecosystem performance. At the leaf and single-plant level, spectral sensors funnel light reflected from vegetation through a holographic diffraction grating, which separates light by wavelength across the electromagnetic spectrum [ 58 ]. Hyperspectral imaging data is in three ‘cubed’ dimensions with spectral wavelength (z) across spatial co-ordinates (x,y). Depending on the size of a single-pixel hyperspectral cameras can image vegetation from the whole plant to ecosystem scale [ 58 ].

Reflected light has become a powerful tool to characterize plant traits, including photosynthesis, given the varying response of light to leaf structure and pigment content at different wavelengths. In the near infrared (770–1300 nm), differences in chlorophyll and plant nitrogen content indicate a variety of vegetation stressors such as nutrient deficiency [ 59 , 60 ], plant disease status [ 61 , 62 ], and ozone damage [ 63 ], while the short wave infrared (SWIR1; 1300–2500 nm) indicates plant water status based traits [ 64 , 65 ]. In the past, discrete spectral reflectance indices were used as proxies for crop status [ 66 ]. However, computational and technological advances make it possible to derive photosynthetic capacities (maximum rate of carboxylation for C3 and C4 plants, V cmax and V max , respectively; and maximum rate of electron transport, J max ) and make predictions about photosynthetic performance scaling from the leaf [ 67–71 ] to the plot [ 72 , 73 ] and ecosystem scales [ 74 ].

One significant advance is the commercial availability of high-resolution fiber optic leaf clip-attachments. Hyperspectral radiometers typically contain a radiometrically calibrated light source and standardized white and dark reference panels for calibration. Leaf-level reflective intensity is compared with the reference material. Computer models (discussed later) are then used to correlate portions of the leaf's reflective spectrum with traditional measurements of gas exchange. Hyperspectral data can provide significant information about leaf photosynthesis at a fraction of the time compared with gas exchange [ 67–71 , 75 ]. These measurements can offer insight for upscaling to the plot level using field push carts [ 76 ] or drones mounted with hyperspectral cameras for breeding and research trials.

In addition to the hyperspectral methods mentioned above, recently handheld multispectral tools (e.g. FluroPen, Photo Systems Instruments, Drásov, Czech Republic; MultispeQ, PHOTOSYNQ INC. East Lancing MI, U.S.A.; and LI-600, LiCOR Biosciences Lincoln NE, U.S.A.) are used to monitor fluorescence and other parameters associated with leaves. Compared with hyperspectral leaf clips or fluorescence chambers sold with gas exchange units, these leaf tools can be used to more quickly and inexpensively screen for the vitality of photosynthetic systems under biotic and abiotic stresses (e.g. [ 77 ]). Furthermore, these tools provide opportunity, in some cases, to specify wavebands of interest for specific phenotypes that can extend beyond photosynthetic measurements.

Inspired by the successful leaf-level estimations of photosynthetic capacities, hyperspectral imaging (HSI) techniques are increasingly applied to canopy-scale measurements [ 73 , 78 ]. Imaging hyperspectral spectrometers provide more spatial information than a leaf-clip portable radiometer. Because of this, these sensors are being utilized to reveal variability in photosynthetic traits of interest across leaves, plants, and/or over large geographic areas [ 72 , 74 ]. These HSI sensors can scan individual plants in a few seconds [ 79 ] or provide analysis spanning several km 2 if mounted on aircraft or Earth-orbiting satellites [ 80 , 81 ]. Compared with point-based portable radiometers, these HSI sensors result in the accumulation of large amounts of data that need to be processed in an innovative way.

To link reflectance spectra to photosynthetic physiological parameters, data processing pipelines must be tailored to specific sensing platforms. These data pipelines are critical to applications such as field phenotyping in a high-throughput manner. For leaf-level estimations of photosynthetic variables using reflectance spectra, great efforts have been made to select statistical techniques that can provide the best predictive power [ 75 ]. Partial Least Square Regression (PLSR) [ 82 ] is currently the most common technique used to relate reflectance spectra to photosynthesis associated parameters [ 68 , 71 ] due to its ability to reduce tens to hundreds of spectral bands to just a few orthogonal principle components (also known as latent variables). There are also other machine learning algorithms such as Artificial Neural Network (ANN)-based regression and Least Absolute Shrinkage and Selection Operator (LASSO) that have been used to estimate photosynthesis [ 83 ]. The availability of these machine learning and empirical algorithms also poses a dilemma regarding the most effective approach. Collectively harnessing the strengths of individual empirical or machine learning algorithms through regression stacking shows promise [ 72 ] although further studies are needed to test its effectiveness across more plant species. For estimations of photosynthesis using reflectance spectra at the plot and ecosystem levels, further data processing steps are necessary to account for spurious variations in reflectance caused by sun-target-sensor geometry, canopy structure, leaf scattering, atmospheric contaminations, and background soil [ 75 ]. These steps are required to ensure that only reflectance data associated with photosynthesis are used for estimations. Although Radiative Transfer Models (RTMs) such as PROSAIL [ 84 ] are developed to remove those spurious variations, few of them can be directly used in the proximal sensing setting [ 85 ]. However, these RTMs provide an alternative way to reduce hyperspectral data into several meaningful leaf traits, such as chlorophyll concentration, that can serve as a proxy for photosynthesis. For example, RTMs-inverted traits were shown to explain up to 60% of variation in photosynthetic physiology in a crop species [ 72 ].

Remote-sensing products that measure GPP are traditionally based on the Light-Use Efficiency (LUE) concept of ecosystem modeling [ 86 ] and empirical models that rely on the relationships between remote sensing-derived variables and GPP [ 87–90 ]. These methods provide reasonable estimates of GPP compared with measured EC fluxes, however, new emerging spectral sensing technologies including Solar-Induced chlorophyll Fluorescence (SIF) are providing potential for estimating GPP at the ecosystem scale [ 91–93 ]. A fraction of solar radiation absorbed by chlorophyll is emitted as fluorescence, hence SIF is more physiologically based than other traditional remote sensing products [ 94 ] as it is a direct product of the photosynthetic process [ 95–97 ]. While pulse amplitude modulated chlorophyll fluorescence has long been used to measure photochemical efficiencies and heat dissipation in individual leaves [ 98 ], this should not be confused with SIF, which relies on measuring of the radiance chlorophyll fluorescence from an ecosystem.

Passive SIF measurements were first applied at the satellite scale ( Table 6 ) [ 99 ] to assess regional and global scale patterns of SIF alongside GPP [ 91–93 ] and is now being implemented at flux towers across multiple ecosystem types to determine the physiological and structural relationship between SIF and photosynthesis at this scale [ 100–103 ]. Likewise, the near-infrared radiance of vegetation index (NIR v ) has shown promising accuracy at detecting photosynthetic variability at the hourly scale over crop and forest system [ 104 , 105 ]. Therefore, both SIF and NIR v should enable real-time monitoring of productivity and stress.

Table 6

SIF measurement was first applied at the satellite scale [ 99 ] to assess regional and global scale patterns of SIF alongside GPP [ 91–93 ]. Currently, it is being implemented at flux towers across multiple ecosystem types to determine the physiological and structural relationship between SIF and photosynthesis at this scale [ 100–103 ]. For comparison, the EC method has a spatial resolution between hundred meters and several kilometers, and a continuous temporal resolution (half-hour) with a fine spatial coverage at the ecosystem and landscape scales.

The relationship between SIF and GPP is primarily dominated by absorbed photosynthetic active radiation (APAR) [ 106 , 107 ], implying that the correlation between SIF and GPP is the highest when photosynthesis is primarily light-limited [ 108 , 109 ]. However, GPP is also controlled by environmental factors other than light, and recent insights suggest that SIF responded to environmental stresses in a similar way as GPP, encouraging the application of SIF to estimate photosynthesis [ 94 ]. A relationship between SIF and GPP was similar among ecosystems although the relationship was stronger for grasslands than forests, savannas and croplands, and for C4 grasslands and crops than C3 ecosystems [ 94 ]. This quasi-universal relationship indicates that SIF could be a valuable tool for inferring GPP of the land surface. More collaborative studies between the EC and remote sensing communities are needed to evaluate why the relationship between SIF and GPP varies among ecosystems and under differing environmental conditions to improve the ability of SIF products to estimate ecosystem GPP robustly to scale regionally and globally.

Much progress has been made to understand the relationship between SIF and GPP but many challenges remain [ 109–111 ]. Higher spatial and temporal resolution SIF measurements are needed to coincide with the continuous GPP measurements [ 112 ]. Promising solutions to these challenges would be to develop remote sensing approaches that can cross-calibrate and blend multi-source SIF and reflectance measurements for a consistent record in both spatial and temporal domains. For example, combining satellite SIF with satellite reflectance was used to generate a spatially and temporally continuous SIF dataset [ 113 ]. Another solution is to improve SIF sensor designs to facilitate measurements at a much higher spatial and temporal resolutions. For example, the Fluorescence Imaging Spectrometer (FLORIS) onboard the Fluorescence EXplorer (FLEX) satellite can provide SIF at a better spatial resolution than its predecessors ( Table 6 ) [ 114 ] and the newly launched Orbiting Carbon Observatory 3 instrument (OCO-3) allow for more coverage globally at higher definition [ 115 ].

Interestingly, much of the work on remote sensing has initiated with large-scale measurements, yet there is a tremendous need to increase throughput of measurements at leaf and plot scales, particularly for application in high throughput phenotyping facilities. Whether these techniques are fully scalable remains uncertain, yet the opportunity for multidisciplinary research has advanced the versatility of the tools outlined in this review beyond their original users. Moving forward, simplifying data collection through ‘turn-key’ sensors and standardizing data analysis pipelines for the variety of techniques outlined here are certain to advance understanding of plant function from molecular to global scale.

• Monitoring Photosynthesis at every scale, from leaf to ecosystem, is an important task given the challenges of climate change and growing human populations.
• In the past 5 years there have been significant improvements to the technology and computational tools used to measure photosynthesis at every scale. And new facilities and equipment are being used around the world to monitor photosynthesis.
• Hyperspectral imaging at the leaf, and canopy scale paired with improved computational modeling allows for rapid estimates of important biochemical parameters.
• Micrometeorological approaches to estimate Gross Primary Productivity have been improved by the uses of sulfur tracing elements.
• Monitoring Solar Induced Fluorescence is a promising satellite-based method that should enable real-time monitoring of global ecosystem productivity.

Acknowledgements

This work is supported by funding from Global Change and Photosynthesis Research Unit of the USDA Agricultural Research Service. Any opinions, findings, and conclusions or recommendations expressed in this publication are those of the author(s) and do not necessarily reflect the views of the U.S. Department of Agriculture. Mention of trade names or commercial products in this publication is solely for the purpose of providing specific information and does not imply recommendation or endorsement by the U.S. Department of Agriculture. USDA is an equal opportunity provider and employer.

Abbreviations

Competing interests.

The authors declare that there are no competing interests associated with the manuscript.

Author Contribution

M.H.S., N.G.-C., and C.J.B. conceived the outline, all authors contributed to the organization and writing of the manuscript, M.H.S. and N.G-C. Edited the manuscript, and C.J.B. supervised the project.

Photosynthesis in Aquatic Plants

Cite this chapter.

• J. A. Raven  

Part of the book series: Springer Study Edition ((SSE,volume 100))

1133 Accesses

5 Citations

To address the topic of the ecophysiology of photosynthesis in aquatic plants in the space allotted is a daunting task, and the coverage must of necessity be very selective. Since this Volume is honoring Professor Dr. Lange, I shall emphasize those aspects which interface with his work, and since the rest of the Volume deals with terrestrial plants, I shall emphasize more generally the comparison with terrestrial plants. My third objective is to address particularly those aspects which most interest me and which are, or could be, growth-points in research.

This is a preview of subscription content, log in via an institution to check access.

Access this chapter

• Available as PDF
• Read on any device
• Instant download
• Own it forever
• Compact, lightweight edition
• Dispatched in 3 to 5 business days
• Free shipping worldwide - see info

Tax calculation will be finalised at checkout

Purchases are for personal use only

Institutional subscriptions

Unable to display preview.  Download preview PDF.

Alberte RS (1989) Physiological and cellular features of Prochloron . In: Lewin RA, Cheng L (eds) Prochloron : a microbial enigma. Chapman and Hall, New York, pp 31-52

Chapter   Google Scholar  

Andrews JH (1991) Comparative ecology of microorganisms and macroorganisms. Springer, Berein Heidelberg New York

Book   Google Scholar  

Babin M, Levasseur M, Michaud D, Legendre L (1992) Effect of angular distribution of light on photosynthesis at the scale of a phytoplankton cell. In: Maestrini S (ed) Poster abstracts of symposium on measurement of primary production from the molecular to the global scale. ICES, Copenhagen, p 1

Google Scholar  

Berg M (1983) Random walks in biology. Princeton University Press, Princeton

Bertsch A (1966) CO 2 -Gaswechsel und Wasserhaushalt der aerophilen Grünalge Apactococcus lobatus . Planta 70: 46–62

Article   Google Scholar  

Boston HL, Farmer AM, Madsen TD, Adams MS, Hurley JP (1991) Light-harvesting caratenoids in two deep-water bryophytes. Photosynthetica 25: 61–66

CAS   Google Scholar  

Britting SA (1992) Effect of emergence on the physiology and biochemistry of a high intertidal alga, Endocladia muricata (Post & Rupt) J Ag (Cryptonemiates, Rhodophyta). PhD dissertation, University of California, Los Angeles

Cooper LW, De Niro MJ (1989) Detection of heavy isotopes of oxygen and hydrogen in tissue water of intertidal plants: implications for water economy. Mar Biol 101: 397–400

Article   CAS   Google Scholar  

Cowan IR, Lange OL, Green TGA (1992) Carbon-dioxide exchange in lichens: determination of transport and carboxylation reactions. Planta 187: 282–284

Den Hartog C, Segal C (1964) A new classification of water plant communities. Acta Bot Neerl 13: 367–393

Dring MJ (1981) Chromatic adaptation of photosynthesis in benthic marine algae: an examination of its ecological significance using a theoretical model. Limnol Oceanogr 26: 271–284

Fawley MW (1991) Disjunct distribution of the xanthophyll loroxanthin in the green algae. J Phycol 27: 544–548

Fujiward S, Iwahashi H, Someya J, Nishikawd S (1993) Structure and cotranscription of the plastid-encoded rbcL and rbcS genes of Pleuroehrysis carterae (Prymnesiophyta). J Phycol 29: 347–355

Goericke R, Repeta DJ (1992) The pigments of Prochlorococcus marinus : the presence of divinyl chlorophyll a and b in a marine procaryote. Limnol Oceanogr 37: 425–433

Greene RM, Gerard VA (1990) Effects of high-frequency light fluctuation on growth and photoacclimation of the red alga Chondus crispus . Mar Biol 105: 337–344

Guillard RRL, Keller MD, O’Kelly CJ, Floyd GL (1991) Pycnococcus provasolii gen. et sp. nov., a coccoid prasinoxanthin-containing phytoplankton from the western North Atlantic and Gulf of Mexico. J Phycol 27: 39–47

Karentz D, Cleaver JE, Mitchell DL (1991) Cell survival characteristics and molecular responses of phytoplankton to ultraviolet B radiation. J phycol 27: 326–341

Kirk JTO (1985) Light and photosynthesis in aquatic ecosystems. Cambridge University Press, Cambridge

Kyle DJ, Osmond CB, Arntzen CJ (eds) (1987) Photoinhibition. Elsevier, Amsterdam

Lange OL (1989) Ecophysiology and photosynthesis: performance of poikilohydric and homoiohydric plants. In: Greuter W, Zimmer B (eds) Proceedings of the XIV International Botanical Congress. Koeltz, Königstein/Taunus, pp 357–383

Lange OL, Schulze ED, Koch W (1970) Experimental-ökologische Untersuchungen an Flechter der Negev-Wüste II CO 2 -Gaswechsel und Wasserhaushalt von Ramalina maciformis (De) Bory am natürlichen Standort während der sommerlichen Trockenperiode. Flora (Jena) 159: 38–62

Lichtlé C, Spilar A, Dural JC (1992) Immunogold localization of light-harvesting and photosystem I complexes in the thylakoids of Fucus serratus (Phaeophyceae). Protoplasma 166: 99–106

Littler MM, Littler DS, Blair SM, Norris JN (1985) Deepest known plant life discovered on an unchanted seamount. Science 227: 57–59

Article   PubMed   CAS   Google Scholar  

Littler MM, Littler DS, Blair SM, Norris JN (1986) Deep water plant communities from an uncharted seamount off San Salvador Island, Bahamas: distribution, abundance and primary productivity. Deep-Sea Res 33: 881–892

Luther H (1947) Vorschlag zu einer ökologischen Grundeinteilung der Hydrophyten. Acta Bot Fenn 44: 1–15

Maberly SC (1985a) Photosynthesis by Fontinalis antipyretica . I. Interaction between photon irradiance, concentration of carbon dioxide and temperature. New Phytol 100: 127–140

Maberly SC (1985b) Photosynthesis by Fontinalis antipyretica . II. Assessment of environmental factors limiting photosynthesis and production. New Phytol 100: 141–155

Maberly SC, Madsen TV (1990) Contribution of air and water in the carbon balance of Fucus spiralis . Mar Ecol Prog Ser 62: 175–183

Madsen TV, Maberly SC (1990) A comparison of air and water as environments for photosynthesis by the intertidal alga Fucus vesiculosus (Phaeophyta). J Phycol 26: 24–30

Mitchell JG (1991) The influence of cell size on marine bacterial mobility and energetics. Microb Ecol 22: 227–238

Newman JR (1991) Carbon assimilation by freshwater aquatic macrophytes. PhD Thesis, University of Dundee

Palenik B, Haselkorn R (1992) Multiple evolutionary origins of prochlorophytes, the chlorophyll b -containing prokaryotes. Nature 355: 265–267

Pienaar RN (1980) Chrysophytes. In: Cox ER (ed) Phytoflagellates. Elsevier, New York, pp 213–242

Pyszniak AM, Gibbs SP (1992) Immunocytochemical localization of photosystem I and the fucoxanthin-chorophyll a/c light-harvesting complex in the diatom Phaeodactylum tricornutum . Protoplasma 166: 208–217

Ramus J (1978) Seaweed anatomy and photosynthetic performance: the ecological significance of light guides, heterogenous absorption and multiple scatter. J Phycol 14: 352–362

Raunkaier C (1934) The life forms of plants and statistical plant geography. Clarendon Press, Oxford

Raven JA (1981) Nutritional strategies of submerged benthic plants: the acquisition of C, N and P by rhizophytes and haptophytes. New Phytol 88: 1–30

Raven JA (1984a) Energetics and transport in aquatic plants. Liss, New York

Raven JA (1984b) A cost-benefit analysis of photon absorption by photosynthetic unicells. New Phytol 98: 593–625

Raven JA (1986a) Physiological consequences of extremely small size for autotrophic organisms in the sea. In: Platt T, Li WKW (eds) Photosynthetic picoplankton. Can Bull Fish Aquat Sci 214: 1–70

Raven JA (1986b) Evolution of life forms. In: Givnish TV (ed) On the economy of plant form and function. Cambridge University Press, Cambridge, pp 421–492

Raven JA (1987) Biochemistry, biophysics and physiology of chlorophyll b-containing algae: implications for taxonomy and phylogeny. Prog Phycol Res 5: 1–122

Raven JA (1989) Fight or flight: the economics of repair and avoidance of photoinhibition of photosynthesis. Funct Ecol 3: 5–19

Raven JA (1991a) Responses of aquatic photosynthetic organisms to increased solar UV-B. J Photochem Photobiol B: Biology 9: 239–244

Raven JA (1991b) Implications of inorganic C utilization: ecology, evolution and geochemistry. Can J Bot 68: 905–924

Raven JA (1991c) Physiology of inorganic C acquisition and implications for resource use efficiency by marine phytoplankton: relation to increased CO 2 and temperature. Plant Cell Environ 14: 779–794

Raven JA (1992a) Energy and nutrient acquisition by autotrophic symbioses. Symbiosis 14: 33–60

Raven JA (1992b) How benthic macroalgae cope with flowing freshwater: resource acquisition and retention. J Phycol 28: 133–146

Raven JA (1993a) Comparative aspects of chrysophyte nutrition with emphasis on carbon, phosphorus and nitrogen. In: Sandgren CD, Smol JP, Kristiansen J (eds) Chrysophyte algae: ecology, phylogeny and development. Cambridge University Press, Cambridge (in press)

Raven JA (1993b) Carbon: a phycocentric view. In: Evans GT, Fasham MJR (eds) Towards & Model of Ocean Biogeochemical Cycles. Springer, Berlin Heidelberg New York, pp 123–152

Raven JA, Johnston AM (1991a) Photosynthetic inorganic carbon assimilation by Prasiola stipitata (Prasiolales, Chlorophyta) under emersed and submersed conditions: relationship to the taxonomy of Prasiola . Br Phycol J 26: 247–247

Raven JA, Johnston AM (1991b) Carbon assimilation mechanisms. Implications for intensive culture of seaweeds. In: Garcia-Reina G, Pedersen M (eds) Seaweed cellular biotechnology, physiology and intensive cultivation. Las Palmas de Gran Canada, Espana, pp 151–166

Raven JA, Johnston AM (1991c) Mechanisms of inorganic carbon acquisition in marine phytoplankton and their implications for the use of other resources. Limnol Oceanogr 36: 1701–1714

Raven JA, Richardson K (1984) Dinophyte flagella: a cost-benefit analysis. New Phytol 98: 259–276

Raven JA, Richardson K (1986) Marine environments. In: Baker NR, Long SP (eds) Photosynthesis in contrasting envioronments. Elsevier, Amsterdam, pp 337–398

Raven JA, Samuelsson G (1986) Repair of photoinhibity damage in Anacystis nidulans 625 ( Synechococcus 6301): relation to catalytic capacity for, and energy supply to, protein synthesis, and implications for μ max and the efficiency of light-limited growth. New Phytol 103: 625–643

Raven JA, Smith FA, Smith SE (1980) Ions and osmoregulation. In: Rains DW, Valentine RC, Hollaender A (eds) Genetic engineering of osmoregulation: impact on plant productivity for food, chemicals and energy. Plenum Press, New York, pp 101–118

Raven JA, Handley LL, MacFarlane JJ, McInroy S, McKenzie L, Richards JH, Samuelsson G (1988) The role of root CO 2 uptake and CAM in inorganic C acquisition by plants of the isoetid life form. A review, with new data on Eriocaulon decangulare . New Phytol 108: 125–148

Raven JA, Johnston AM, Surif MB (1989) The photosynthetic apparatus as a phyletic character. In: Green JC, Leadbeater BSC, Diver WL (eds) The chromophyte algae. Problems and perspectives. Oxford Science Publications, Oxford, pp 63–84

Raven JA, Johnston AM, MacFarlane JJ (1990a) Carbon metabolism. In: Sheath RG, Cole KM (eds) The biology of the red algae. Cambridge University Press, New York, pp 171–202

Raven JA, Johnston AM, Handley LL, McInroy SG (1990b) Transport and assimilation of inorganic carbon by Lichina pygmaea under emersed and submersed conditions. New Phytol 114: 407–417

Reiskind JB, Bowes G (1991) The role of phosphoenolpyruvate carboxykinase in a marine macroalga with C 4 -like photosynthetic characteristics. Proc Natl Acad Sci USA 88: 2993–2887

Rowan KS (1989) Photosynthetic pigments of algae. Cambridge University Press, Cambridge

Rugg DA, Norton TA (1987) Pelvetia canaliculata , a high-shore seaweed that shuns the sea. In: Crawford RMM (ed) Plant life in aquatic and amphibious habitats. Blackwell, Oxford, pp 347–358

Sand-Jensen K, Pedersen MF, Nielsen SL (1992) Photosynthetic use of inorganic carbon among primary and secondary water plants in streams. Freshwater Biol 27: 283–293

Stebbins GL, Hill GJC (1980) Did multicellular plants invade the land? Am Nat 115: 342–353

Surif MB, Raven JA (1990) Photosynthetic gas exchange under emersed conditions in eulittoral and normally submersed members of the Fucales and the Laminariales: interpretation in relation to C isotope ratio and N and water use efficiency. Oecologia 82: 68–80

Thomas TE, Turpin DH, Harrison PJ (1987) Desiccation enhanced nitrogen uptake rates in intertidal seaweeds. Mar Biol 94: 293–298

Thomsen HA (1986) A survey of the smallest eucaryotic organisms of the marine phytoplankton. In: Platt TL, Li WKW (eds) Photosynthetic picoplankton. Can Bull Fish Aquat Sci 214: 121–158

Urbach E, Robertson DL, Chisholm SW (1992) Multiple evolutionary origins of prochlorophytes within the cyanobacterial radiation. Nature 355: 267–270

Wilhelm C, Wiedemann I (1991) Evidence of protein-bound chlorophyll c 3 in a light-harvesting protein isolated from the flagellate alga Prymnesium parvum (Prymnesiophyceae). Photosynthetica 25: 249–255

Willemoes M, Monas E (1991) Relationship between growth irradiance and xanthophyll cycle pool in the diatom Nitzschia palea . Physiol Plant 83: 459–456

Download references

You can also search for this author in PubMed   Google Scholar

Editor information

Editors and affiliations.

Lehrstuhl für Pflanzenökologie, Universität Bayreuth, Postfach 101251, Bayreuth, D-95447, Germany

Ernst-Detlef Schulze

Department of Range Science and Ecology Center, Utah State University, Logan, UT, 84322-5230, USA

Martyn M. Caldwell

Rights and permissions

Reprints and permissions

Copyright information

© 1995 Springer-Verlag Berlin Heidelberg

About this chapter

Raven, J.A. (1995). Photosynthesis in Aquatic Plants. In: Schulze, ED., Caldwell, M.M. (eds) Ecophysiology of Photosynthesis. Springer Study Edition, vol 100. Springer, Berlin, Heidelberg. https://doi.org/10.1007/978-3-642-79354-7_15

Download citation

DOI : https://doi.org/10.1007/978-3-642-79354-7_15

Publisher Name : Springer, Berlin, Heidelberg

Print ISBN : 978-3-540-58571-8

Online ISBN : 978-3-642-79354-7

eBook Packages : Springer Book Archive

Share this chapter

Anyone you share the following link with will be able to read this content:

Sorry, a shareable link is not currently available for this article.

Provided by the Springer Nature SharedIt content-sharing initiative

• Publish with us

Policies and ethics

• Find a journal
• Track your research

Thank you for visiting nature.com. You are using a browser version with limited support for CSS. To obtain the best experience, we recommend you use a more up to date browser (or turn off compatibility mode in Internet Explorer). In the meantime, to ensure continued support, we are displaying the site without styles and JavaScript.

• View all journals
• Explore content
• About the journal
• Publish with us
• Sign up for alerts
• Review Article
• Published: 09 April 2024

Plant responses to changing rainfall frequency and intensity

• Andrew F. Feldman   ORCID: orcid.org/0000-0003-1547-6995 1 , 2 ,
• Xue Feng   ORCID: orcid.org/0000-0003-1381-3118 3 , 4 ,
• Andrew J. Felton 5 ,
• Alexandra G. Konings   ORCID: orcid.org/0000-0002-2810-1722 6 ,
• Alan K. Knapp   ORCID: orcid.org/0000-0003-1695-4696 7 ,
• Joel A. Biederman 8 &
• Benjamin Poulter   ORCID: orcid.org/0000-0002-9493-8600 1  

Nature Reviews Earth & Environment volume  5 ,  pages 276–294 ( 2024 ) Cite this article

1065 Accesses

71 Altmetric

Metrics details

• Carbon cycle
• Climate-change ecology

Regardless of annual rainfall amount changes, daily rainfall events are becoming more intense but less frequent with anthropogenic warming. Larger rainfall events and longer dry spells have complex and sometimes opposing effects on plant photosynthesis and growth, challenging abilities to understand broader consequences on the carbon cycle. In this Review, we evaluate global plant responses to rainfall regimes characterized by fewer, larger rainfall events across evidence from field experiments, satellites and models. Plant function responses vary between −28% and 29% (5th to 95th percentile) under fewer, larger rainfall events, with the direction of response contingent on climate; productivity increases are more common in dry ecosystems (46% positive; 20% negative), whereas responses are typically negative in wet ecosystems (28% positive; 51% negative). Contrasting responses in dry and wet ecosystems are attributed to nonlinear plant responses to soil moisture driven by several ecohydrological mechanisms. For example, dry ecosystem plants are more sensitive to large rainfall pulses compared with wet ecosystem plants, partly driving dry ecosystem positive responses to fewer, larger rainfall events. Knowledge gaps remain over optimal rainfall frequencies for photosynthesis, the relative dominance of rainfall pulse and dry spell mechanisms and the disproportionate role of extreme rainfall pulses on plant function.

This is a preview of subscription content, access via your institution

Access options

Access Nature and 54 other Nature Portfolio journals

Get Nature+, our best-value online-access subscription

24,99 € / 30 days

cancel any time

Subscribe to this journal

Receive 12 digital issues and online access to articles

92,52 € per year

only 7,71 € per issue

Buy this article

• Purchase on Springer Link
• Instant access to full article PDF

Prices may be subject to local taxes which are calculated during checkout

Similar content being viewed by others

Stomatal responses of terrestrial plants to global change

Xingyun Liang, Defu Wang, … David S. Ellsworth

Contrasting responses of woody and grassland ecosystems to increased CO2 as water supply varies

Yude Pan, Robert B. Jackson, … Yiqi Luo

Responses of plant diversity to precipitation change are strongest at local spatial scales and in drylands

Lotte Korell, Harald Auge, … Tiffany M. Knight

Data availability

CMIP6 rainfall projections can be obtained from https://cds.climate.copernicus.eu . Observation-based rainfall data sets from MERRA, CPC and GPCC can be obtained from https://gmao.gsfc.nasa.gov/reanalysis/MERRA-2/data_access , https://psl.noaa.gov/data/gridded/data.cpc.globalprecip.html and https://psl.noaa.gov/data/gridded/data.gpcc.html , respectively. FLUXNET observations can be downloaded from https://fluxnet.org .

Ruehr, S. et al. Evidence and attribution of the enhanced land carbon sink. Nat. Rev. Earth Environ. 4 , 518–534 (2023).

Article   CAS   Google Scholar  

Friedlingstein, P. et al. Global Carbon Budget 2022. Earth Syst. Sci. Data 14 , 4811–4900 (2022).

Article   Google Scholar  

Yang, Y. et al. Evapotranspiration on a greening Earth. Nat. Rev. Earth Environ. 4 , 626–641 (2023).

Green, J. K. et al. Regionally strong feedbacks between the atmosphere and terrestrial biosphere. Nat. Geosci. 10 , 410–414 (2017).

Forzieri, G., Alkama, R., Miralles, D. G. & Cescatti, A. Satellites reveal contrasting responses of regional climate to the widespread greening of Earth. Science 360 , 1180–1184 (2018).

Spracklen, D. V., Arnold, S. R. & Taylor, C. M. Observations of increased tropical rainfall preceded by air passage over forests. Nature 489 , 282–285 (2012).

Nemani, R. R. et al. Climate-driven increases in global terrestrial net primary production from 1982 to 1999. Science 300 , 1560–1563 (2003).

Humphrey, V. et al. Sensitivity of atmospheric CO 2 growth rate to observed changes in terrestrial water storage. Nature 560 , 628–631 (2018).

Madani, N., Kimball, J. S., Jones, L. A., Parazoo, N. C. & Guan, K. Global Analysis of bioclimatic controls on ecosystem productivity using satellite observations of solar-induced chlorophyll fluorescence. Remote Sens. 9 , 530 (2017).

Poulter, B. et al. Contribution of semi-arid ecosystems to interannual variability of the global carbon cycle. Nature 509 , 600–603 (2014).

Ahlström, A. et al. The dominant role of semi-arid ecosystems in the trend and variability of the land CO 2 sink. Science 348 , 895–900 (2015).

Green, J. K. et al. Large influence of soil moisture on long-term terrestrial carbon uptake. Nature 565 , 476–479 (2019).

Reichstein, M. et al. Climate extremes and the carbon cycle. Nature 500 , 287–295 (2013).

Ciais, P. et al. Europe-wide reduction in primary productivity caused by the heat and drought in 2003. Nature 437 , 529–533 (2005).

Reyer, C. P. O. et al. A plant’s perspective of extremes: terrestrial plant responses to changing climatic variability. Glob. Chang. Biol. 19 , 75–89 (2013).

Sala, O. E., Parton, W. J., Joyce, L. A. & Lauenroth, W. K. Primary production of the central grassland region of the United States. Ecology 69 , 40–45 (1988).

Biederman, J. A. et al. CO 2 exchange and evapotranspiration across dryland ecosystems of southwestern North America. Glob. Chang. Biol. 23 , 4204–4221 (2017).

Ukkola, A. M. et al. Annual precipitation explains variability in dryland vegetation greenness globally but not locally. Glob. Chang. Biol. 27 , 4367–4380 (2021).

Pendergrass, A. G. & Knutti, R. The uneven nature of daily precipitation and its change. Geophys. Res. Lett. 45 , 11,980–11,988 (2018).

Fowler, H. J. et al. Anthropogenic intensification of short-duration rainfall extremes. Nat. Rev. Earth Environ. 2 , 107–122 (2021).

Kannenberg, S. A., Bowling, D. R. & Anderegg, W. R. L. Hot moments in ecosystem fluxes: high GPP anomalies exert outsized influence on the carbon cycle and are differentially driven by moisture availability across biomes. Environ. Res. Lett. 15 , 054004 (2020).

Trenberth, K. E. Changes in precipitation with climate change. Clim. Res. 47 , 123–138 (2011).

Pendergrass, A. G., Knutti, R., Lehner, F., Deser, C. & Sanderson, B. M. Precipitation variability increases in a warmer climate. Sci. Rep. 7 , 1–9 (2017). This study estimates daily rainfall frequency and intensity trends, both observed and model projected, and compares with trends with annual rainfall totals.

Giorgi, F., Raffaele, F. & Coppola, E. The response of precipitation characteristics to global warming from climate projections. Earth Syst. Dyn. 10 , 73–89 (2019). An investigation of daily rainfall frequency and intensity-projected trends within CMIP5 experiments.

Holdrege, M. C., Kulmatiski, A., Palmquist, K. A. & Beard, K. H. Precipitation intensification increases shrub dominance in arid, not mesic, ecosyst. Ecosystems 26 , 568–584 (2023).

Thomey, M. L. et al. Effect of precipitation variability on net primary production and soil respiration in a Chihuahuan Desert grassland. Glob. Chang. Biol. 17 , 1505–1515 (2011).

Short Gianotti, D. J., Akbar, R., Feldman, A. F., Salvucci, G. D. & Entekhabi, D. Terrestrial evaporation and moisture drainage in a warmer climate. Geophys. Res. Lett. 47 , e2019GL086498 (2020).

McColl, K. A. et al. Global characterization of surface soil moisture drydowns. Geophys. Res. Lett. 44 , 3682–3690 (2017).

Zhang, F., Biederman, J. A. & Dannenberg, M. P. Five decades of observed daily precipitation reveal longer and more variable drought events across much of the Western United States. Geophys. Res. Lett. 48 , e2020GL092293 (2021).

Feldman, A. F., Short Gianotti, D. J., Trigo, I. F., Salvucci, G. D. & Entekhabi, D. Land–atmosphere drivers of landscape-scale plant water content loss. Geophys. Res. Lett. 47 , e2020GL090331 (2020).

Grossiord, C. et al. Plant responses to rising vapor pressure deficit. N. Phytol. 226 , 1550–1566 (2020).

Ross, I. et al. How do variations in the temporal distribution of rainfall events affect ecosystem fluxes in seasonally water-limited Northern Hemisphere shrublands and forests? Biogeosciences 9 , 1007–1024 (2012).

Liu, J. et al. Impact of temporal precipitation variability on ecosystem productivity. Wiley Interdiscip. Rev. Water 7 , e1481 (2020).

Knapp, A. K. et al. Rainfall variability, carbon cycling, and plant species diversity in a mesic grassland. Science 298 , 2202–2205 (2002). One of the first field experiments evaluating effects of fewer, larger rainfall events on soil and plants at a Kansas site where many rainfall manipulation experiments have been performed.

Liu, W. J. et al. Repackaging precipitation into fewer, larger storms reduces ecosystem exchanges of CO 2 and H 2 O in a semiarid steppe. Agric. For. Meteorol. 247 , 356–364 (2017).

Fay, P. A., Kaufman, D. M., Nippert, J. B., Carlisle, J. D. & Harper, C. W. Changes in grassland ecosystem function due to extreme rainfall events: implications for responses to climate change. Glob. Chang. Biol. 14 , 1600–1608 (2008).

Heisler-White, J. L., Blair, J. M., Kelly, E. F., Harmoney, K. & Knapp, A. K. Contingent productivity responses to more extreme rainfall regimes across a grassland biome. Glob. Chang. Biol. 15 , 2894–2904 (2009).

Piao, S. et al. Characteristics, drivers and feedbacks of global greening. Nat. Rev. Earth Environ. 1 , 14–27 (2020).

Baudena, M., Boni, G., Ferraris, L., von Hardenberg, J. & Provenzale, A. Vegetation response to rainfall intermittency in drylands: results from a simple ecohydrological box model. Adv. Water Resour. 30 , 1320–1328 (2007).

Feng, X., Porporato, A. & Rodriguez-Iturbe, I. Changes in rainfall seasonality in the tropics. Nat. Clim. Chang. 3 , 811–815 (2013).

Su, J., Zhang, Y. & Xu, F. Divergent responses of grassland productivity and plant diversity to intra- annual precipitation variability across climate regions: a global synthesis. J. Ecol. 111 , 1–14 (2023). Synthesized plant responses to changing rainfall frequency and intensity from field experiments across the globe.

Knapp, A. K. et al. Consequences of more extreme precipitation regimes for terrestrial ecosystems. Bioscience 58 , 811–821 (2008). Uses a soil water bucket model to provide a framework for why plants in dry and wet ecosystems would have opposing responses to fewer, larger wet days.

Wang, L. et al. Dryland productivity under a changing climate. Nat. Clim. Chang. 12 , 981–994 (2022).

Lian, X. et al. Multifaceted characteristics of dryland aridity changes in a warming world. Nat. Rev. Earth Environ. 2 , 232–250 (2021).

Huxman, T. E. et al. Precipitation pulses and carbon fluxes in semiarid and arid ecosystems. Oecologia 141 , 254–268 (2004).

Held, I. M. & Soden, B. J. Robust responses of the hydrological cycle to global warming. J. Clim. 19 , 5686–5699 (2006).

Lee, J.-Y. et al. in Climate Change 2021 — The Physical Science Basis: Contribution of Working Group I to the Sixth Assessment Report of the Intergovernmental Panel on Climate Change Ch. 4 (eds Masson-Delmotte, V. et al.) 553–672 (Cambridge Univ. Press, 2023).

Byrne, M. P. & O’Gorman, P. A. The response of precipitation minus evapotranspiration to climate warming: why the ‘wet-get-wetter, dry-get-drier’ scaling does not hold over land. J. Clim. 28 , 8078–8093 (2015).

Zaitchik, B. F., Rodell, M., Biasutti, M. & Seneviratne, S. I. Wetting and drying trends under climate change. Nat. Water 1 , 502–513 (2023).

Alexander, L. V. et al. Global observed changes in daily climate extremes of temperature and precipitation. J. Geophys. Res. Atmos. 111 , 1–22 (2006).

Ziese, M. et al. GPCC full data daily version 2022 at 1.0°: daily land-surface precipitation from rain-gauges built on GTS-based and historic data. GPCC https://doi.org/10.5676/DWD_GPCC/FD_D_V2022_100 (2022).

Reichle, R. H. et al. Land surface precipitation in MERRA-2. J. Clim. 30 , 1643–1664 (2017).

Chen, M. et al. Assessing objective techniques for gauge-based analyses of global daily precipitation. J. Geophys. Res. Atmos. 113 , 1–13 (2008).

Google Scholar  

Pendergrass, A. G. What precipitation is extreme? Science 360 , 1072–1073 (2018).

O’Gorman, P. A. & Schneider, T. The physical basis for increases in precipitation extremes in simulations of 21st-century climate change. Proc. Natl Acad. Sci. USA 106 , 14773–14777 (2009).

Pendergrass, A. G. & Gerber, E. P. The rain is askew: two idealized models relating vertical velocity and precipitation distributions in a warming world. J. Clim. 29 , 6445–6462 (2016).

Pfahl, S., O’Gorman, P. A. & Fischer, E. M. Understanding the regional pattern of projected future changes in extreme precipitation. Nat. Clim. Chang. 7 , 423–427 (2017).

Lu, J., Vecchi, G. A. & Reichler, T. Expansion of the Hadley cell under global warming. Geophys. Res. Lett. 34 , 2–6 (2007).

Vecchi, G. A. & Soden, B. J. Global warming and the weakening of the tropical circulation. J. Clim. 20 , 4316–4340 (2007).

Donat, M. G., Lowry, A. L., Alexander, L. V., O’Gorman, P. A. & Maher, N. More extreme precipitation in the world’s dry and wet regions. Nat. Clim. Chang. 6 , 508–513 (2016).

Donat, M. G., Angélil, O. & Ukkola, A. M. Intensification of precipitation extremes in the world’s humid and water-limited regions. Environ. Res. Lett. 14 , 065003 (2019).

Fischer, E. M., Beyerle, U. & Knutti, R. Robust spatially aggregated projections of climate extremes. Nat. Clim. Chang. 3 , 1033–1038 (2013).

Miranda, J., Padilla, F. M., Lázaro, R. & Pugnaire, F. I. Do changes in rainfall patterns affect semiarid annual plant communities? J. Veg. Sci. 20 , 269–276 (2009).

Zhang, J. et al. Moderately prolonged dry intervals between precipitation events promote production in Leymus chinensis in a semi-arid grassland of Northeast China. BMC Plant. Biol. 21 , 1–11 (2021).

Grant, K., Kreyling, J., Beierkuhnlein, C. & Jentsch, A. Importance of seasonality for the response of a mesic temperate grassland to increased precipitation variability and warming. Ecosystems 20 , 1454–1467 (2017).

Ma, Y., Zhang, T. & Liu, X. Effect of intensity of small rainfall simulation in spring on annuals in Horqin Sandy Land, China. Environ. Earth Sci. 74 , 727–735 (2015).

Padilla, F. M. et al. Effects of extreme rainfall events are independent of plant species richness in an experimental grassland community. Oecologia 191 , 177–190 (2019).

Schuster, M. J., Smith, N. G. & Dukes, J. S. Responses of aboveground C and N pools to rainfall variability and nitrogen deposition are mediated by seasonal precipitation and plant community dynamics. Biogeochemistry 129 , 389–400 (2016).

Arca, V., Power, S. A., Delgado-Baquerizo, M., Pendall, E. & Ochoa-Hueso, R. Seasonal effects of altered precipitation regimes on ecosystem-level CO 2 fluxes and their drivers in a grassland from Eastern Australia. Plant Soil 460 , 435–451 (2021).

Zhang, F. et al. Precipitation temporal repackaging into fewer, larger storms delayed seasonal timing of peak photosynthesis in a semi‐arid grassland. Funct. Ecol. 36 , 646–658 (2021).

Holdrege, M. C., Beard, K. H. & Kulmatiski, A. Woody plant growth increases with precipitation intensity in a cold semiarid system. Ecology 102 , 1–11 (2021).

Fay, P. A. et al. Relative effects of precipitation variability and warming on tallgrass prairie ecosystem function. Biogeosciences 8 , 3053–3068 (2011).

Kulmatiski, A. & Beard, K. H. Woody plant encroachment facilitated by increased precipitation intensity. Nat. Clim. Chang. 3 , 833–837 (2013).

Berry, R. S. & Kulmatiski, A. A savanna response to precipitation intensity. PLoS ONE 12 , 1–18 (2017).

Williams, K. J., Wilsey, B. J., McNaughton, S. J. & Banyikwa, F. F. Temporally variable rainfall does not limit yields of Serengeti grasses. Oikos 81 , 463 (1998).

Xie, Y., Li, Y., Xie, T., Meng, R. & Zhao, Z. Impact of artificially simulated precipitation pattern change on the growth and morphology of Reaumuria soongarica seedlings in Hexi Corridor of China. Sustain 12 , 2439 (2020).

Good, S. P. & Caylor, K. K. Climatological determinants of woody cover in Africa. Proc. Natl Acad. Sci. USA 108 , 4902–4907 (2011).

Zhao, H., Jia, G., Xu, X., Zhang, A. & Wang, H. Divergent effects of intensified precipitation on primary production in global drylands. Sci. Total Environ. 892 , 164736 (2023).

Case, M. F. & Staver, A. C. Soil texture mediates tree responses to rainfall intensity in African savannas. N. Phytol. 219 , 1363–1372 (2018).

Ritter, F., Berkelhammer, M. & Garcia-Eidell, C. Distinct response of gross primary productivity in five terrestrial biomes to precipitation variability. Commun. Earth Environ. 1 , 34 (2020).

Fang, J. et al. Precipitation patterns alter growth of temperate vegetation. Geophys. Res. Lett. 32 , 1–5 (2005).

Xu, X. et al. Tree cover shows strong sensitivity to precipitation variability across the global tropics. Glob. Ecol. Biogeogr. 27 , 450–460 (2018).

Zhang, Y. et al. Extreme precipitation patterns and reductions of terrestrial ecosystem production across biomes. J. Geophys. Res. Biogeosci. 118 , 148–157 (2013).

Zhang, W., Brandt, M., Tong, X., Tian, Q. & Fensholt, R. Impacts of the seasonal distribution of rainfall on vegetation productivity across the Sahel. Biogeosciences 15 , 319–330 (2018).

Guo, Q. et al. Contrasting responses of gross primary productivity to precipitation events in a water-limited and a temperature-limited grassland ecosystem. Agric. For. Meteorol. 214–215 , 169–177 (2015).

D’Onofrio, D., Sweeney, L., von Hardenberg, J. & Baudena, M. Grass and tree cover responses to intra-seasonal rainfall variability vary along a rainfall gradient in African tropical grassy biomes. Sci. Rep. 9 , 1–10 (2019).

Porporato, A., Daly, E. & Rodriguez-Iturbe, I. Soil water balance and ecosystem response to climate change. Am. Nat. 164 , 625–632 (2004). This study uses a minimalist process model to predict and attribute plant responses to fewer, larger rainfall events at the Knapp et al. (2002) Kansas field experiment.

Zhang, D. H., Li, X. R., Zhang, F., Zhang, Z. S. & Chen, Y. L. Effects of rainfall intensity and intermittency on woody vegetation cover and deep soil moisture in dryland ecosystems. J. Hydrol. 543 , 270–282 (2016).

Nordbotten, J. M., Rodriguez-Iturbe, I. & Celia, M. A. Stochastic coupling of rainfall and biomass dynamics. Water Resour. Res. 43 , 1–7 (2007).

Xu, X., Medvigy, D. & Rodriguez-Iturbe, I. Relation between rainfall intensity and savanna tree abundance explained by water use strategies. Proc. Natl Acad. Sci. USA 112 , 12992–12996 (2015).

Guan, K. et al. Continental-scale impacts of intra-seasonal rainfall variability on simulated ecosystem responses in Africa. Biogeosciences 11 , 6939–6954 (2014).

Medvigy, D., Wofsy, S. C., Munger, J. W. & Moorcroft, P. R. Responses of terrestrial ecosystems and carbon budgets to current and future environmental variability. Proc. Natl Acad. Sci. USA 107 , 8275–8280 (2010).

Peng, S. et al. Precipitation amount, seasonality and frequency regulate carbon cycling of a semi-arid grassland ecosystem in Inner Mongolia, China: a modeling analysis. Agric. For. Meteorol. 178–179 , 46–55 (2013).

Fay, P. A. et al. Altered rainfall patterns, gas exchange, and growth in grasses and forbs. Int. J. Plant Sci. 163 , 549–557 (2002).

Griffin-Nolan, R. J., Slette, I. J. & Knapp, A. K. Deconstructing precipitation variability: rainfall event size and timing uniquely alter ecosystem dynamics. J. Ecol. 109 , 3356–3369 (2021).

Rodríguez-Iturbe, I., D’Odorico, P., Porporato, A. & Ridolfi, L. On the spatial and temporal links between vegetation, climate, and soil moisture. Water Resour. 35 , 3709–3722 (1999).

Laio, F., Porporato, A., Fernandez-Illescas, C. P. & Rodriguez-Iturbe, I. Plants in water-controlled ecosystems: active role in hydrologic processes and response to water stress II. Probabilistic soil moisture dynamics. Adv. Water Resour. 24 , 707–723 (2001).

Daly, E., Porporato, A. & Rodriguez-Iturbe, I. Coupled dynamics of photosynthesis, transpiration, and soil water balance. Part II: stochastic analysis and ecohydrological significance. J. Hydrometeorol. 5 , 559–566 (2004).

Daly, E., Porporato, A. & Rodríguez-Iturbe, I. Coupled dynamics of photosynthesis, transpiration, and soil water balance. Part I: upscaling from hourly to daily level. J. Hydrometeorol. 5 , 546–558 (2004).

Seneviratne, S. I. et al. Investigating soil moisture–climate interactions in a changing climate: a review. Earth Sci. Rev. 99 , 125–161 (2010).

Porporato, A., Laio, F., Ridolfi, L. & Rodriguez-Iturbe, I. Plants in water-controlled ecosystems: active role in hydrologic processes and response to water stress III. Vegetation water stress. Adv. Water Resour. 24 , 725–744 (2001).

Guan, K. et al. Simulated sensitivity of African terrestrial ecosystem photosynthesis to rainfall frequency, intensity, and rainy season length. Environ. Res. Lett. 13 , 025013 (2018).

Nippert, J. B., Knapp, A. K. & Briggs, J. M. Intra-annual rainfall variability and grassland productivity: can the past predict the future? Plant Ecol. 184 , 65–74 (2006).

Noy-Meir, I. Desert ecosystems: environment and producers. Annu. Rev. Ecol. Syst. 4 , 25–52 (1973). This paper poses the ‘ pulse reserve paradigm ’ to describe dryland plant responses to individual rainfall events.

Felton, A. J., Slette, I. J., Smith, M. D. & Knapp, A. K. Precipitation amount and event size interact to reduce ecosystem functioning during dry years in a mesic grassland. Glob. Chang. Biol. 26 , 658–668 (2020).

Fay, P. A., Carlisle, J. D., Knapp, A. K., Blair, J. M. & Collins, S. L. Productivity responses to altered rainfall patterns in a C 4-dominated grassland. Oecologia 137 , 245–251 (2003).

Post, A. K. & Knapp, A. K. How big is big enough? Surprising responses of a semiarid grassland to increasing deluge size. Glob. Chang. Biol. 27 , 1157–1169 (2021).

Robinson, T. M. P. & Gross, K. L. The impact of altered precipitation variability on annual weed species. Am. J. Bot. 97 , 1625–1629 (2010).

Wilcox, K. R., von Fischer, J. C., Muscha, J. M., Petersen, M. K. & Knapp, A. K. Contrasting above- and belowground sensitivity of three Great Plains grasslands to altered rainfall regimes. Glob. Chang. Biol. 21 , 335–344 (2015).

Slette, I. J., Blair, J. M., Fay, P. A., Smith, M. D. & Knapp, A. K. Effects of compounded precipitation pattern intensification and drought occur belowground in a mesic grassland. Ecosystems 25 , 1265–1278 (2021).

Zeppel, M. J. B., Wilks, J. V. & Lewis, J. D. Impacts of extreme precipitation and seasonal changes in precipitation on plants. Biogeosciences 11 , 3083–3093 (2014).

Huang, H., Calabrese, S. & Rodriguez-Iturbe, I. J. Variability of ecosystem carbon source from microbial respiration is controlled by rainfall dynamics. Proc. Natl Acad. Sci. USA 118 , e2115283118 (2021).

Berg, A., Sultan, B. & De Noblet-Ducoudré, N. What are the dominant features of rainfall leading to realistic large-scale crop yield simulations in West Africa? Geophys. Res. Lett. 37 , 1–6 (2010).

Harper, C. W., Blair, J. M., Fay, P. A., Knapp, A. K. & Carlisle, J. D. Increased rainfall variability and reduced rainfall amount decreases soil CO 2 flux in a grassland ecosystem. Glob. Chang. Biol. 11 , 322–334 (2005).

Felton, A. J. et al. Climate disequilibrium dominates uncertainty in long-term projections of primary productivity. Ecol. Lett. 25 , 2688–2698 (2022).

Knapp, A. K. et al. Past, present, and future roles of long-term experiments in the LTER network. Bioscience 62 , 377–389 (2012).

Beier, C. et al. Precipitation manipulation experiments — challenges and recommendations for the future. Ecol. Lett. 15 , 899–911 (2012).

Adler, P. B., White, E. P. & Cortez, M. H. Matching the forecast horizon with the relevant spatial and temporal processes and data sources. Ecography 43 , 1729–1739 (2020).

Smith, M. D., Knapp, A. K. & Collins, S. L. A framework for assessing ecosystem dynamics in response to chronic resource alterations induced by global change. Ecology 90 , 3279–3289 (2009).

Feng, X., Dawson, T. E., Ackerly, D. D., Santiago, L. S. & Thompson, S. E. Reconciling seasonal hydraulic risk and plant water use through probabilistic soil–plant dynamics. Glob. Chang. Biol. 23 , 3758–3769 (2017).

Throop, H. L., Reichmann, L. G., Sala, O. E. & Archer, S. R. Response of dominant grass and shrub species to water manipulation: an ecophysiological basis for shrub invasion in a Chihuahuan Desert Grassland. Oecologia 169 , 373–383 (2012).

Javadian, M. et al. Thermography captures the differential sensitivity of dryland functional types to changes in rainfall event timing and magnitude. N. Phytol. 240 , 114–126 (2023).

Reynaert, S. et al. Risk of short-term biodiversity loss under more persistent precipitation regimes. Glob. Chang. Biol. 27 , 1614–1626 (2021).

Jackson, R. B., Banner, J. L., Jobbagy, E. G., Pockman, W. T. & Wall, D. H. Ecosystem carbon loss with woody plant invasion of grasslands. Nature 418 , 623–626 (2002).

Gherardi, L. A. & Sala, O. E. Enhanced precipitation variability decreases grass- and increases shrub-productivity. Proc. Natl Acad. Sci. USA 112 , 12735–12740 (2015).

Schreiner-McGraw, A. P. et al. Woody plant encroachment has a larger impact than climate change on dryland water budgets. Sci. Rep. 10 , 1–9 (2020).

Hao, Y. et al. Climate-induced abrupt shifts in structural states trigger delayed transitions in functional states. Ecol. Indic. 115 , 106468 (2020).

Harrison, S. P., Gornish, E. S. & Copeland, S. Climate-driven diversity loss in a grassland community. Proc. Natl Acad. Sci. USA 112 , 8672–8677 (2015).

Barbeta, A., Ogaya, R. & Peñuelas, J. Dampening effects of long-term experimental drought on growth and mortality rates of a Holm oak forest. Glob. Chang. Biol. 19 , 3133–3144 (2013).

Jones, S. K., Collins, S. L., Blair, J. M., Smith, M. D. & Knapp, A. K. Altered rainfall patterns increase forb abundance and richness in native tallgrass prairie. Sci. Rep. 6 , 1–10 (2016).

Zhang, Y., Keenan, T. F. & Zhou, S. Exacerbated drought impacts on global ecosystems due to structural overshoot. Nat. Ecol. Evol. 5 , 1490–1498 (2021).

Jump, A. S. et al. Structural overshoot of tree growth with climate variability and the global spectrum of drought-induced forest dieback. Glob. Chang. Biol. 23 , 3742–3757 (2017).

Collins, S. L. et al. Stability of tallgrass prairie during a 19-year increase in growing season precipitation. Funct. Ecol. 26 , 1450–1459 (2012).

Keenan, T. F. et al. Increase in forest water-use efficiency as atmospheric carbon dioxide concentrations rise. Nature 499 , 324–327 (2013).

Donohue, R. J., Roderick, M. L., McVicar, T. R. & Farquhar, G. D. Impact of CO 2 fertilization on maximum foliage cover across the globe’s warm, arid environments. Geophys. Res. Lett. 40 , 3031–3035 (2013).

Zhang, Y. et al. Increasing sensitivity of dryland vegetation greenness to precipitation due to rising atmospheric CO 2 . Nat. Commun. 13 , 4875 (2022).

Roby, M. C., Scott, R. L. & Moore, D. J. P. High vapor pressure deficit decreases the productivity and water use efficiency of rain-induced pulses in semiarid ecosystems. J. Geophys. Res. Biogeosci. 125 , 1–14 (2020).

Katul, G. G., Oren, R., Manzoni, S., Higgins, C. & Parlange, M. B. Evapotranspiration: a process driving mass transport and energy exchange in the soil–plant–atmosphere–climate system. Rev. Geophys. 50 , 1–25 (2012).

Reynolds, J. F., Kemp, P. R., Ogle, K. & Fernández, R. J. Modifying the ‘pulse-reserve’ paradigm for deserts of North America: precipitation pulses, soil water, and plant responses. Oecologia 141 , 194–210 (2004).

Fensham, R. J., Butler, D. W. & Foley, J. How does clay constrain woody biomass in drylands? Glob. Ecol. Biogeogr. 24 , 950–958 (2015).

Wei, L. et al. Experimental investigation of relationship between infiltration rate and soil moisture under rainfall conditions. Water 14 , 1–11 (2022).

Penna, D., Tromp-Van Meerveld, H. J., Gobbi, A., Borga, M. & Dalla Fontana, G. The influence of soil moisture on threshold runoff generation processes in an alpine headwater catchment. Hydrol. Earth Syst. Sci. 15 , 689–702 (2011).

Western, A. W. & Grayson, R. B. The Terrawarra data set: soil moisture patterns, soil characteristics, and hydrological flux measurements. Water Resour. Res. 34 , 2765–2768 (1998).

Vereecken, H. et al. Soil hydrology in the Earth system. Nat. Rev. Earth Environ. 3 , 573–587 (2022).

Lian, X., Zhao, W. & Gentine, P. Recent global decline in rainfall interception loss due to altered rainfall regimes. Nat. Commun. 13 , 7642 (2022).

Thomey, M. L., Collins, S. L., Friggens, M. T., Brown, R. F. & Pockman, W. T. Effects of monsoon precipitation variability on the physiological response of two dominant C 4 grasses across a semiarid ecotone. Oecologia 176 , 751–762 (2014).

Lauenroth, W. K. & Bradford, J. B. Ecohydrology of dry regions of the United States: water balance consequences of small precipitation events. Ecohydrology 5 , 46–53 (2012).

Demaria, E. M. C., Hazenberg, P. & Meles, M. B. Intensification of the North American Monsoon rainfall as observed from a long‐term high‐density gauge network geophysical research letters. Geophys. Res. Lett. 46 , 6839–6847 (2019).

Pockman, W. T. & Small, E. E. The influence of spatial patterns of soil moisture on the grass and shrub responses to a summer rainstorm in a Chihuahuan desert ecotone. Ecosystems 13 , 511–525 (2010).

McDonald, A. K., Kinucan, R. J. & Loomis, L. E. Ecohydrological interactions within banded vegetation in the northeastern Chihuahuan Desert, USA. Ecohydrology 2 , 66–71 (2009).

Schreiner-Mcgraw, A. P., Ajami, H. & Vivoni, E. R. Extreme weather events and transmission losses in arid streams. Environ. Res. Lett . 14 , 084002 (2019).

CAS   Google Scholar  

Scott, R. L. & Biederman, J. A. Critical zone water balance over 13 years in a semiarid savanna. Water Resour. Res. 55 , 574–588 (2019).

Nippert, J. B. & Holdo, R. M. Challenging the maximum rooting depth paradigm in grasslands and savannas. Funct. Ecol. 29 , 739–745 (2015).

Potts, D. L. et al. Antecedent moisture and seasonal precipitation influence the response of canopy-scale carbon and water exchange to rainfall pulses in a semi-arid grassland. N. Phytol. 170 , 849–860 (2006).

He, Z., Zhao, W., Liu, H. & Chang, X. The response of soil moisture to rainfall event size in subalpine grassland and meadows in a semi-arid mountain range: a case study in northwestern China’s Qilian Mountains. J. Hydrol. 420–421 , 183–190 (2012).

Tumber-Dávila, S. J., Schenk, H. J., Du, E. & Jackson, R. B. Plant sizes and shapes above and belowground and their interactions with climate. N. Phytol. 235 , 1032–1056 (2022).

Jackson, R. B. et al. A global analysis of root distributions for terrestrial biomes. Oecologia 108 , 389–411 (1996).

Raz-Yaseef, N., Yakir, D., Rotenberg, E., Schiller, G. & Cohen, S. Ecohydrology of a semi-arid forest: partitioning among water balance components and its implications for predicted precipitation changes. Ecohydrology 3 , 143–154 (2010).

Fravolini, A. et al. Precipitation pulse use by an invasive woody legume: the role of soil texture and pulse size. Oecologia 144 , 618–627 (2005).

Zhu, X. et al. Soil coarsening alleviates precipitation constraint on vegetation growth in global drylands. Environ. Res. Lett. 17 , 11 (2022).

Ogle, K., Wolpert, R. L. & Reynolds, J. F. Reconstructing plant root area and water uptake profiles. Ecology 85 , 1967–1978 (2004).

Schwinning, S., Sala, O. E., Loik, M. E. & Ehleringer, J. R. Thresholds, memory, and seasonality: understanding pulse dynamics in arid/semi-arid ecosystems. Oecologia 141 , 191–193 (2004).

Brodribb, T. J. & Cochard, H. Hydraulic failure defines the recovery and point of death in water-stressed conifers. Plant Physiol. 149 , 575–584 (2009).

Bassiouni, M., Good, S. P., Still, C. J. & Higgins, C. W. Plant water uptake thresholds inferred from satellite soil moisture. Geophys. Res. Lett. 47 , e2020GL087077 (2020).

Jonard, F., Feldman, A. F., Gianotti, D. J. S. & Entekhabi, D. Observed water- and light-limitation across global ecosystems. Biogeosciences 19 , 5575–5590 (2022).

Short Gianotti, D. J., Rigden, A. J., Salvucci, G. D. & Entekhabi, D. Satellite and station observations demonstrate water availability’s effect on continental-scale evaporative and photosynthetic land surface dynamics. Water Resour. Res. 55 , 540–554 (2019).

Feldman, A. F., Short Gianotti, D. J., Trigo, I. F., Salvucci, G. D. & Entekhabi, D. Satellite-based assessment of land surface energy partitioning–soil moisture relationships and effects of confounding variables. Water Resour. Res. 55 , 10657–10677 (2019).

Schwinning, S. & Sala, O. E. Hierarchy of responses to resource pulses in arid and semi-arid ecosystems. Oecologia 141 , 211–220 (2004).

Famiglietti, C. A., Michalak, A. M. & Konings, A. G. Extreme wet events as important as extreme dry events in controlling spatial patterns of vegetation greenness anomalies. Environ. Res. Lett. 16 , 074014 (2021).

Novick, K. A. et al. Confronting the water potential information gap. Nat. Geosci. 15 , 158–164 (2022).

Feldman, A. F., Short Gianotti, D. J., Konings, A. G., Gentine, P. & Entekhabi, D. Patterns of plant rehydration and growth following pulses of soil moisture availability. Biogeosciences 18 , 831–847 (2021).

Post, A. K. & Knapp, A. K. Plant growth and aboveground production respond differently to late-season deluges in a semi-arid grassland. Oecologia 191 , 673–683 (2019).

Huxman, T. E. et al. Response of net ecosystem gas exchange to a simulated precipitation pulse in a semi-arid grassland: the role of native versus non-native grasses and soil texture. Oecologia 141 , 295–305 (2004).

Raz-Yaseef, N., Yakir, D., Schiller, G. & Cohen, S. Dynamics of evapotranspiration partitioning in a semi-arid forest as affected by temporal rainfall patterns. Agric. For. Meteorol. 157 , 77–85 (2012).

Klein, T., Cohen, S. & Yakir, D. Hydraulic adjustments underlying drought resistance of Pinus halepensis . Tree Physiol. 31 , 637–648 (2011).

Feldman, A. F., Chulakadabba, A., Short Gianotti, D. J. & Entekhabi, D. Landscape-scale plant water content and carbon flux behavior following moisture pulses: from dryland to mesic environments. Water Resour. Res. 57 , e2020WR027592 (2021).

Potts, D. L., Barron-Gafford, G. A. & Scott, R. L. Ecosystem hydrologic and metabolic flashiness are shaped by plant community traits and precipitation. Agric. For. Meteorol. 279 , 107674 (2019).

Guo, Q. et al. Responses of gross primary productivity to different sizes of precipitation events in a temperate grassland ecosystem in Inner Mongolia, China. J. Arid Land 8 , 36–46 (2016).

Williams, C. A., Hanan, N., Scholes, R. J. & Kutsch, W. Complexity in water and carbon dioxide fluxes following rain pulses in an African savanna. Oecologia 161 , 469–480 (2009).

Parton, W., Morgan, J., Smith, D. & Grosso, S. D. E. L. Impact of precipitation dynamics on net ecosystem productivity. Glob. Chang. Biol. 18 , 915–927 (2012).

Scott, R. L., Biederman, J. A., Hamerlynck, E. P. & Barron-Gafford, G. A. The carbon balance pivot point of southwestern U.S. semiarid ecosystems: insights from the 21st century drought. J. Geophys. Res. Biogeosci. 120 , 2612–2624 (2015).

Pastorello, G., Trotta, C. & Canfora, E. The FLUXNET2015 dataset and the ONEFlux processing pipeline for eddy covariance data. Sci. Data 7 , 225 (2020).

Post, A. K. & Knapp, A. K. The importance of extreme rainfall events and their timing in a semi-arid grassland. J. Ecol. 108 , 2431–2443 (2020).

Collins, S. L. et al. A multiscale, hierarchical model of pulse dynamics in arid-land ecosystems. Annu. Rev. Ecol. Evol. Syst. 45 , 397–419 (2014).

Feldman, A. F. et al. Moisture pulse-reserve in the soil-plant continuum observed across biomes. Nat. Plants 4 , 1026–1033 (2018). An evaluation of plant sensitivity to individual rainfall pulses and their soil moisture thresholds using satellite measurements across the tropics.

Harris, B. L. et al. Satellite-observed vegetation responses to intraseasonal precipitation variability. Geophys. Res. Lett. 49 , 1–11 (2022).

Roman, D. T. et al. The role of isohydric and anisohydric species in determining ecosystem-scale response to severe drought. Oecologia 179 , 641–654 (2015).

Ignace, D. D., Huxman, T. E., Weltzin, J. F. & Williams, D. G. Leaf gas exchange and water status responses of a native and non-native grass to precipitation across contrasting soil surfaces in the Sonoran Desert. Oecologia 152 , 401–413 (2007).

Craine, J. M. et al. Timing of climate variability and grassland productivity. Proc. Natl Acad. Sci. USA 109 , 3401–3405 (2012).

Hao, Y. B. et al. Aboveground net primary productivity and carbon balance remain stable under extreme precipitation events in a semiarid steppe ecosystem. Agric. For. Meteorol. 240–241 , 1–9 (2017).

Brown, R. F., Sala, O. E., Sinsabaugh, R. L. & Collins, S. L. Temporal effects of monsoon rainfall pulses on plant available nitrogen in a Chihuahuan desert grassland. J. Geophys. Res. Biogeosci. 127 , e2022JG006938 (2022).

Sitch, S. et al. Evaluation of ecosystem dynamics, plant geography and terrestrial carbon cycling in the LPJ dynamic global vegetation model. Glob. Chang. Biol. 9 , 161–185 (2003).

Zhang, F. et al. Using high frequency digital repeat photography to quantify the sensitivity of a semi-arid grassland ecosystem to the temporal repackaging of precipitation. Agric. For. Meteorol. 338 , 109539 (2023).

Paschalis, A. et al. Rainfall manipulation experiments as simulated by terrestrial biosphere models: where do we stand? Glob. Chang. Biol. 26 , 3336–3355 (2020).

McColl, K. A. et al. The global distribution and dynamics of surface soil moisture. Nat. Geosci. 10 , 100–104 (2017).

Cherwin, K. & Knapp, A. Unexpected patterns of sensitivity to drought in three semi-arid grasslands. Oecologia 169 , 845–852 (2012).

Cochard, H., Bréda, N. & Granier, A. Whole tree hydraulic conductance and water loss regulation in Quercus during drought: evidence for stomatal control of embolism? Ann. For. Sci. 53 , 197–206 (1996).

Medlyn, B. E. et al. Reconciling the optimal and empirical approaches to modelling stomatal conductance. Glob. Chang. Biol. 17 , 2134–2144 (2011).

Berdugo, M. et al. Global ecosystem thresholds driven by aridity. Science 367 , 787–790 (2020).

North, G. B. & Nobel, P. S. Changes in hydraulic conductivity and anatomy caused by drying and rewetting roots of agave deserti (Agavaceae). Am. J. Bot. 78 , 906 (1991).

McDowell, N. G. et al. Mechanisms of woody-plant mortality under rising drought, CO 2 and vapour pressure deficit. Nat. Rev. Earth Environ. 3 , 294–308 (2022).

Choat, B. et al. Global convergence in the vulnerability of forests to drought. Nature 491 , 752–755 (2012).

Gherardi, L. A. & Sala, O. E. Effect of interannual precipitation variability on dryland productivity: a global synthesis. Glob. Chang. Biol. 25 , 269–276 (2019).

Konings, A. G. & Gentine, P. Global variations in ecosystem-scale isohydricity. Glob. Chang. Biol. 23 , 891–905 (2017).

Still, C. J., Berry, J. A., Collatz, G. J. & DeFries, R. S. Global distribution of C3 and C4 vegetation: carbon cycle implications. Global Biogeochem. Cycles 17 , 1006 (2003).

Yu, K. & D’Odorico, P. Climate, vegetation, and soil controls on hydraulic redistribution in shallow tree roots. Adv. Water Resour. 66 , 70–80 (2014).

Larsen, L., Thomas, C., Eppinga, M. & Coulthard, T. Exploratory modeling: extracting causality from complexity. Eos Trans. Am. Geophys. Union 95 , 285–286 (2014).

Weltzin, J. F. et al. Assessing the response of terrestrial ecosystems to potential changes in precipitation. Bioscience 53 , 941 (2003).

Kröel-Dulay, G. et al. Field experiments underestimate aboveground biomass response to drought. Nat. Ecol. Evol. 6 , 540–545 (2022).

Novick, K. A. et al. The increasing importance of atmospheric demand for ecosystem water and carbon fluxes. Nat. Clim. Chang. 6 , 1023–1027 (2016).

Lundholm, J. T. & Larson, D. W. Experimental separation of resource quantity from temporal variability: seedling responses to water pulses. Oecologia 141 , 346–352 (2004).

Ogle, K. & Reynolds, J. F. Plant responses to precipitation in desert ecosystems: integrating functional types, pulses, thresholds, and delays. Oecologia 141 , 282–294 (2004).

Orth, R. When the land surface shifts gears. AGU Adv. 2 , 2019–2022 (2021).

Hsu, J. S., Powell, J. & Adler, P. B. Sensitivity of mean annual primary production to precipitation. Glob. Chang. Biol. 18 , 2246–2255 (2012).

Maurer, G. E., Hallmark, A. J., Brown, R. F., Sala, O. E. & Collins, S. L. Sensitivity of primary production to precipitation across the United States. Ecol. Lett. 23 , 527–536 (2020).

Feldman, A. F., Gianotti, D. J. S., Trigo, I. F., Salvucci, G. D. & Entekhabi, D. Observed landscape responsiveness to climate forcing. Water Resour. Res. 58 , e2021WR030316 (2022).

Dong, J., Akbar, R., Feldman, A., Gianotti, D. S. & Entekhabi, D. Land surfaces at the tipping-point for water and energy balance coupling. Water Resour. Res. 59 , e2022WR032472 (2023).

Fu, Z. et al. Critical soil moisture thresholds of plant water stress in terrestrial ecosystems. Sci. Adv. 7827 , 1–13 (2022).

Fu, Z. et al. Uncovering the critical soil moisture thresholds of plant water stress for European ecosystems. Glob. Chang. Biol. 28 , 2111–2123 (2022).

Jensen, J. On the convex functions and inequalities between mean values. Acta Math. 30 , 175–193 (1906).

Heisler-White, J. L., Knapp, A. K. & Kelly, E. F. Increasing precipitation event size increases aboveground net primary productivity in a semi-arid grassland. Oecologia 158 , 129–140 (2008).

Guan, K., Sultan, B., Biasutti, M., Baron, C. & Lobell, D. B. What aspects of future rainfall changes matter for crop yields in West Africa? Geophys. Res. Lett. 42 , 8001–8010 (2015).

Seyfried, M. S. et al. Ecohydrological control of deep drainage in arid and semiarid regions. Ecology 86 , 277–287 (2005).

Sala, O. E., Gherardi, L. A. & Peters, D. P. C. Enhanced precipitation variability effects on water losses and ecosystem functioning: differential response of arid and mesic regions. Clim. Change 131 , 213–227 (2015).

Berkelhammer, M., Stefanescu, I. C., Joiner, J. & Anderson, L. High sensitivity of gross primary production in the Rocky mountains to summer rain. Geophys. Res. Lett. 44 , 3643–3652 (2017).

Nielsen, U. N. & Ball, B. A. Impacts of altered precipitation regimes on soil communities and biogeochemistry in arid and semi-arid ecosystems. Glob. Chang. Biol. 21 , 1407–1421 (2015).

Biederman, J. A. et al. Terrestrial carbon balance in a drier world: the effects of water availability in southwestern North America. Glob. Chang. Biol. 22 , 1867–1879 (2016).

Siteur, K. et al. How will increases in rainfall intensity affect semiarid ecosystems? Water Resour. Res. 50 , 5980–6001 (2014).

Li, L. et al. Nonlinear carbon cycling responses to precipitation variability in a semiarid grassland. Sci. Total Environ. 781 , 147062 (2021).

Swemmer, A. M., Knapp, A. K. & Snyman, H. A. Intra-seasonal precipitation patterns and above-ground productivity in three perennial grasslands. J. Ecol. 95 , 780–788 (2007).

Fu, Z. et al. Atmospheric dryness reduces photosynthesis along a large range of soil water deficit. Nat. Commun. 13 , 989–999 (2022).

Rigden, A. J., Mueller, N. D., Holbrook, N. M., Pillai, N. & Huybers, P. Combined influence of soil moisture and atmospheric evaporative demand is important for accurately predicting US maize yields. Nat. Food 1 , 127–133 (2020).

Kannenberg, S. A. et al. Quantifying the drivers of ecosystem fluxes and water potential across the soil–plant–atmosphere continuum in an arid woodland. Agric. For. Meteorol. 329 , 109269 (2023).

Konings, A. G., Williams, A. P. & Gentine, P. Sensitivity of grassland productivity to aridity controlled by stomatal and xylem regulation. Nat. Geosci. 10 , 284–288 (2017).

Zscheischler, J. et al. A few extreme events dominate global interannual variability in gross primary production. Environ. Res. Lett. 9 , 035001 (2014).

Zscheischler, J. et al. Short-term favorable weather conditions are an important control of interannual variability in carbon and water fluxes. J. Geophys. Res. Biogeosci. 121 , 2186–2198 (2016).

Smith, M. D. The ecological role of climate extremes: current understanding and future prospects. J. Ecol. 99 , 651–655 (2011).

Smith, M. D. An ecological perspective on extreme climatic events: a synthetic definition and framework to guide future research. J. Ecol. 99 , 656–663 (2011).

Knapp, A. K. et al. Characterizing differences in precipitation regimes of extreme wet and dry years: implications for climate change experiments. Glob. Chang. Biol. 21 , 2624–2633 (2015).

Sala, O. E. & Lauenroth, W. K. Small rainfall events: an ecological role in semiarid regions. Oecologia 53 , 301–304 (1982).

Rezaei, E. E. et al. Climate change impacts on crop yields. Nat. Rev. Earth Environ. 4 , 831–846 (2023).

Donohue, R. J., McVicar, T. R. & Roderick, M. L. Climate-related trends in Australian vegetation cover as inferred from satellite observations, 1981–2006. Glob. Chang. Biol. 15 , 1025–1039 (2009).

Haverd, V., Ahlström, A., Smith, B. & Canadell, J. G. Carbon cycle responses of semi-arid ecosystems to positive asymmetry in rainfall. Glob. Chang. Biol. 23 , 793–800 (2017).

Copernicus Climate Change Service Climate Data Store. CMIP6 climate projections. C3S https://doi.org/10.24381/cds.c866074c (2021).

Eyring, V. et al. Overview of the Coupled Model Intercomparison Project Phase 6 (CMIP6) experimental design and organization. Geosci. Model Dev. 9 , 1937–1958 (2016).

Fatichi, S., Ivanov, V. Y. & Caporali, E. Investigating interannual variability of precipitation at the global scale: is there a connection with seasonality? J. Clim. 25 , 5512–5523 (2012).

Hajek, O. L. & Knapp, A. K. Shifting seasonal patterns of water availability: ecosystem responses to an unappreciated dimension of climate change. N. Phytol. 233 , 119–125 (2022).

Paschalis, A., Fatichi, S., Katul, G. G. & Ivanov, V. Y. Cross-scale impact of climate temporal variability on ecosystem water and carbon fluxes. J. Geophys. Res. Biogeosci. 120 , 641–660 (2015).

Guo, Q. et al. Spatial variations in aboveground net primary productivity along a climate gradient in Eurasian temperate grassland: effects of mean annual precipitation and its seasonal distribution. Glob. Chang. Biol. 18 , 3624–3631 (2012).

Chou, C. et al. Increase in the range between wet and dry season precipitation. Nat. Geosci. 6 , 263–267 (2013).

Good, S. P., Guan, K. & Caylor, K. K. Global patterns of the contributions of storm frequency, intensity, and seasonality to interannual variability of precipitation. J. Clim. 29 , 3–15 (2016).

Download references

Acknowledgements

The research of A.F.F. was supported by an appointment to the NASA Postdoctoral Program at the NASA Goddard Space Flight Center, administered by Oak Ridge Associated Universities under contract with NASA. A.F.F. was partly supported by a NASA Terrestrial Ecology scoping study for a dryland field campaign. A.G.K. was supported by NSF DEB 1942133 and by the Alfred P. Sloan Foundation. The authors acknowledge the World Climate Research Programme, which, through its Working Group on Coupled Modelling, coordinated and promoted CMIP6. The authors thank the climate modelling groups for producing and making available their model output, the Earth System Grid Federation (ESGF) for archiving the data and providing access and the multiple funding agencies who support CMIP6 and ESGF. This work used eddy covariance data acquired and shared by the FLUXNET community, including AmeriFlux. The FLUXNET eddy covariance data processing and harmonization was carried out by the ICOS Ecosystem Thematic Center, AmeriFlux Management Project and Fluxdata project of FLUXNET, with the support of CDIAC, and the OzFlux, ChinaFlux and AsiaFlux offices.

Author information

Authors and affiliations.

Biospheric Sciences Laboratory, NASA Goddard Space Flight Center, Greenbelt, MD, USA

Andrew F. Feldman & Benjamin Poulter

Earth System Science Interdisciplinary Center, University of Maryland, College Park, MD, USA

Andrew F. Feldman

Department of Civil, Environmental and Geo-Engineering, University of Minnesota, Minneapolis, MN, USA

Saint Anthony Falls Laboratory, University of Minnesota, Minneapolis, MN, USA

Department of Land Resources and Environmental Sciences, Montana State University, Bozeman, MT, USA

Andrew J. Felton

Department of Earth System Science, Stanford University, Stanford, CA, USA

Alexandra G. Konings

Department of Biology and Graduate Degree Program in Ecology, Colorado State University, Fort Collins, CO, USA

Alan K. Knapp

USDA Agricultural Research Service Southwest Watershed Research Center, Tucson, AZ, USA

Joel A. Biederman

You can also search for this author in PubMed   Google Scholar

Contributions

A.F.F. led the Review and wrote the initial draft. X.F., A.J.F., A.G.K, A.K.K, J.A.B. and B.P. all contributed substantial edits to the initial outline, manuscript and figures. All authors contributed equally to the generation of the central ideas in the manuscript and initial figure concepts.

Corresponding author

Correspondence to Andrew F. Feldman .

Ethics declarations

Competing interests.

The authors declare no competing interests.

Peer review

Peer review information.

Nature Reviews Earth & Environment thanks Madelon Case, Rene Orth, Yuting Yang and Yao Zhang for their contribution to the peer review of this work.

Additional information

Publisher’s note Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.

Supplementary information

Supplementary information.

An evolutionary adaptation of photosynthesis occurring mainly in some grass and crop species under which photosynthesis is more efficient because photorespiration is largely avoided.

An evolutionary adaptation of photosynthesis occurring mainly in plant species in arid environments that allows them to save water by only exchanging gases with the atmosphere at night.

The species types and their relative abundance within a defined ecosystem, here referring specifically to plants.

A thermodynamic equation that describes the nonlinear increase of saturation vapour pressure, or the capacity of air to hold water, with increases in air temperature.

Some of the largest organized circulations of air in the atmosphere of the Earth that contribute substantially to weather and climate patterns of the Earth.

Rainfall that is captured and stored by vegetation, even briefly, such that it is prevented from infiltrating into the soil or running off of the ground surface.

A commonly used satellite-based vegetation index that estimates greenness at the top of the vegetation canopy based on satellite measurements in the infrared portion of the electromagnetic spectrum.

The annual cyclic nature of plant functioning, specifically referring to its periodic increase and decrease in functioning during similar months of each year.

A general indicator based on how much water is available for plants to use towards essential plant functions such as photosynthesis and transpiration.

The rainfall rate or rainfall depth over a defined time period. The rainfall rate is often defined hourly across hydrological sciences, although it is designated to be daily in this Review.

The distribution of the root volume of a plant across the soil depth.

Vegetation metrics derived from satellite measurements that typically span large spatial extents, including vegetation areal cover, greenness, height, photosynthetic capacity, water content and others.

An empirical relationship between decreasing soil moisture and decline in plant functions such as photosynthesis or transpiration.

Rights and permissions

Springer Nature or its licensor (e.g. a society or other partner) holds exclusive rights to this article under a publishing agreement with the author(s) or other rightsholder(s); author self-archiving of the accepted manuscript version of this article is solely governed by the terms of such publishing agreement and applicable law.

Reprints and permissions

About this article

Cite this article.

Feldman, A.F., Feng, X., Felton, A.J. et al. Plant responses to changing rainfall frequency and intensity. Nat Rev Earth Environ 5 , 276–294 (2024). https://doi.org/10.1038/s43017-024-00534-0

Download citation

Accepted : 23 February 2024

Published : 09 April 2024

Issue Date : April 2024

DOI : https://doi.org/10.1038/s43017-024-00534-0

Share this article

Anyone you share the following link with will be able to read this content:

Sorry, a shareable link is not currently available for this article.

Provided by the Springer Nature SharedIt content-sharing initiative

Quick links

• Explore articles by subject
• Guide to authors
• Editorial policies

Sign up for the Nature Briefing newsletter — what matters in science, free to your inbox daily.

Teacher Resource Center

Pasco partnerships.

2024 Catalogs & Brochures

• Photosynthesis of Aquatic Plants

Students use an optical dissolved oxygen sensor and a photosynthesis tank to study the photosynthetic rate of aquatic plants under different light conditions.

Supports NGSS Disciplinary Core Idea LS2.B

Grade Level: High School

Subject: Biology • Environmental Science

Student Files

Teacher Files

Sign In to your PASCO account to access teacher files and sample data.

Featured Equipment

Wireless Optical Dissolved Oxygen Sensor

This sensor is perfect for the live or long-term monitoring of dissolved oxygen concentration and saturation, in both the lab and field.

Photosynthesis Tank

Designed to work with PASCO sensors, the Photosynthesis Tank is an ideal apparatus for studying photosynthesis and respiration in aquatic life.

Many lab activities can be conducted with our Wireless , PASPORT , or even ScienceWorkshop sensors and equipment. For assistance with substituting compatible instruments, contact PASCO Technical Support . We're here to help. Copyright © 2020 PASCO

Source Collection: Lab #12

Essential Biology Teacher Lab Manual

More experiments.

Advanced Placement

• Photosynthesis

High School

• Photosynthesis and Respiration with Algae Beads
• Energy Content of Food
• Soil pH (PASPORT sensors)

Measuring the rate of photosynthesis

• Share to Facebook
• Share to Twitter
• Share via Email

Without photosynthesis life as we know it would not exist. It’s worth a moment’s reflection…

There would be no biology without photosynthesis. Plant biomass is the food and fuel for all animals. Plants are the primary producers. These amazing organisms are capable of capturing the energy of sunlight and fixing it in the form of potential chemical energy in organic compounds. The organic compounds are constructed from two principle raw materials; carbon dioxide and water (which is a source of hydrogen). These compounds are stable and can be stored until required for life processes. Hence animals, fungi and non-photosynthetic bacteria depend on these for the maintenance of life.

But how can we measure the rates at which photosynthesis takes place?

The quantities are mind boggling. A hectare (e.g. a field 100 m by 100 m) of wheat can convert as much as 10,000 kg of carbon from carbon dioxide into the carbon of sugar in a year, giving a total yield of 25,000 kg of sugar per year.

There is a total of 7000 x 109 tonnes of carbon dioxide in the atmosphere and photosynthesis fixes 100 x 109 tonnes per year. So 15% of the total carbon dioxide in the atmosphere moves into photosynthetic organisms each year.

What are the different methods of measuring the rate of photosynthesis?

There are a few key methods to calculate the rate of photosynthesis. These include:

• Measuring the uptake of CO2
• Measuring the production of O2
• Measuring the production of carbohydrates
• Measuring the increase in dry mass

As the equation for respiration is almost the reverse of the one for photosynthesis, you will need to think whether these methods measure photosynthesis alone or whether they are measuring the balance between photosynthesis and respiration.

Measuring photosynthesis via the uptake of carbon dioxide

Using ‘immobilised algae’ – It’s easy and accurate to measure the rate of photosynthesis and respiration using immobilised algae in hydrogen carbonate indicator solution – known as the ‘algal balls’ technique. Read the full protocol on  using immobilsed algae to measure photosynthesis .

Using an IRGA – Uptake of CO2 can be measured with the means of an IRGA (Infra-Red Gas Analyser) which can compare the CO2 concentration in gas passing into a chamber surrounding a leaf/plant and the CO2 leaving the chamber.

Using a CO2 monitor – More simply, you could put a plant in a plastic bag and monitor the CO2 concentration in the bag using a CO2 monitor. Naturally, the soil and roots must NOT be in the bag (as they respire). Alternatively, you could place some Bicarbonate Indicator Solution in the bag with the plant and watch the colour change. This would best be done with a reference colour chart to try to make the end-point less subjective. This could give a comparison between several plants. There are difficulties with this method, as I’m sure you can appreciate. The leaf area of the plants should be measured so you can compensate for plant size. Atmospheric air is only 400ppm CO2, so there is not much CO2 to monitor and the plant will soon run out of CO2 to fix.

Measuring photosynthesis via the production of oxygen

Oxygen can be measured by counting bubbles evolved from pondweed, or by using the Audus apparatus to measure the amount of gas evolved over a period of time. To do this, place Cabomba pondweed in an upside down syringe in a water bath connected to a capillary tube (you can also use Elodea, but we find Cabomba more reliable). Put the weed in a solution of NaHCO3 solution. You can then investigate the amount of gas produced at different distances from a lamp. Read a full protocol on how to  investigate photosynthesis using pondweed .

Measuring photosynthesis via the production of carbohydrates

There is a crude method where a disc is cut out of one side of a leaf (using a cork borer against a rubber bung) and weighed after drying. Some days (or even weeks later), a disk is cut out of the other half of the leaf, dried and weighed. Increase in mass of the disc is an indication of the extra mass that has been stored in the leaf. This is very simple to do and enables you to investigate plants growing in the wild. However, you can probably think of several inaccuracies in this method.

Measuring photosynthesis via the increase in dry mass

Dry mass is often monitored by the technique of ‘serial harvests’ where several plants are harvested, dried to constant weight and weighed – this is repeated over the duration of the experiment. If you harvest several plants and record how much mass they have accumulated you will have an accurate measure of the surplus photosynthesis over and above the respiration that has taken place. As with most methods, you need several plants so you have replicate measurements and you can find an average and a standard deviation if necessary.

Investigating the light-dependent reaction in photosynthesis

The rate of decolourisation of DCPIP in the Hill Reaction is a measure of the rate of the light-requiring stages of photosynthesis

• Photosynthesis

Related content

Teaching resources.

• 'Algal balls' - Photosynthesis using algae wrapped in jelly balls
• A-level set practicals - factors affecting rates of photosynthesis
• Demonstrating Oxygen Evolution during Photosynthesis using Pondweed
• Investigating Photosynthesis with the SAPS / NCBE Photosynthesis Kit

REVIEW article

Underwater photosynthesis of submerged plants – recent advances and methods.

• 1 The Freshwater Biological Laboratory, Department of Biology, University of Copenhagen, Hillerød, Denmark
• 2 Institute of Advanced Studies, The University of Western Australia, Crawley, WA, Australia
• 3 School of Plant Biology, The University of Western Australia, Crawley, WA, Australia

We describe the general background and the recent advances in research on underwater photosynthesis of leaf segments, whole communities, and plant dominated aquatic ecosystems and present contemporary methods tailor made to quantify photosynthesis and carbon fixation under water. The majority of studies of aquatic photosynthesis have been carried out with detached leaves or thalli and this selectiveness influences the perception of the regulation of aquatic photosynthesis. We thus recommend assessing the influence of inorganic carbon and temperature on natural aquatic communities of variable density in addition to studying detached leaves in the scenarios of rising CO 2 and temperature. Moreover, a growing number of researchers are interested in tolerance of terrestrial plants during flooding as torrential rains sometimes result in overland floods that inundate terrestrial plants. We propose to undertake studies to elucidate the importance of leaf acclimation of terrestrial plants to facilitate gas exchange and light utilization under water as these acclimations influence underwater photosynthesis as well as internal aeration of plant tissues during submergence.

Knowledge of plant and environmental factors determining photosynthesis by submerged plants is essential for understanding aquatic plant ecophysiology and ecosystem productivity, as well as submergence tolerance of terrestrial plants. Following the pioneering studies by Arens (1933) and Steemann Nielsen (1946) on the use of dissolved inorganic carbon (DIC) for photosynthesis of aquatic plants, numerous studies on the regulatory role of light and DIC for underwater photosynthesis of aquatic plants have been conducted. Particularly, the use of DIC by aquatic plants has fascinated researchers and been reviewed several times (e.g., Madsen and Sand-Jensen, 1991 ; Maberly and Madsen, 2002 ; Raven and Hurd, 2012 ) because this process is important for growth and survival and the uptake mechanisms are very different from those of terrestrial, amphibious, and floating leaved plants exposed to atmospheric air (definitions of these life forms and examples of species are in Sculthorpe (1967) ). Since the physical conditions differ markedly between water and air, we have often been approached by researchers asking for practical advice, unavailable in the literature, before engaging in work with underwater photosynthesis. Thus, this review serves to offer the background and a practical guide for measurements of carbon fixation by plants when under water.

Moreover, a growing number of researchers are interested in tolerance of terrestrial plants during flooding (Figure 1 A). Torrential rains sometimes result in overland floods that inundate terrestrial plants ( Vervuren et al., 2003 ) and with the current projection on climate change, the frequency of such flooding events are expected to increase ( Parry et al., 2007 ). We therefore predict that research on underwater photosynthesis will extend greatly beyond its current focus on aquatic plants as natural wetlands and many crops will become submerged in future flooding events. Researchers engaging in underwater photosynthesis should be aware that physical restrictions on light availability and gas exchange are much more profound under water than on land ( Sand-Jensen and Krause-Jensen, 1997 ). Moreover, the aquatic sources and mechanisms of inorganic carbon use are complex, difficult to study, and often challenging to fully understand ( Madsen and Sand-Jensen, 1991 ; Raven and Hurd, 2012 ).

Figure 1. Completely submerged terrestrial vegetation (A), white flakes of CaCO 3 on leaves of a pondweed ( Potamogeton lucens ) (B) and an incubator with a vertically rotating wheel holding vials with leaf segments for measurements of underwater net photosynthesis (C) or dark respiration when in complete darkness . The bubbles on the submerged mosses (A) are obvious signs that underwater photosynthesis takes place with O 2 evolution causing the bubble formation. Moreover, the submerged grasses possess superhydrophobic self cleansing leaf surfaces that retain a thin gas film when under water, evident as a silvery reflecting surface. In (B) , high pH on the adaxial leaf surfaces following extensive underwater photosynthesis has resulted in precipitation of CaCO 3 (See “Challenges Under Water – Reduced Gas Diffusion and Light Penetration”). Photos: a shallow puddle on Öland, Sweden (A) , the bicarbonate rich (1.8 mmol DIC L −1 ) Lake Slåen, Denmark (B) and the custom built wheel incubator by Ray Scott at the University of Western Australia (C) ; photos by Ole Pedersen.

Photosynthesis provides sugars and O 2 . The importance of underwater photosynthesis to internal O 2 status (Figure 1 B), including via internal long-distance transport into roots growing in anoxic sediments, has been demonstrated for aquatic species (e.g., Borum et al., 2005 ; Sand-Jensen et al., 2005 ; Holmer et al., 2009 ; Pedersen et al., 2011 ) and terrestrial wetland plants when completely submerged ( Pedersen et al., 2006 ; Winkel et al., 2011 , 2013 ). By contrast, during the night submerged plants rely on O 2 uptake from the surrounding water to sustain their respiration and belowground organs can suffer from O 2 deficiency.

The majority of studies on photosynthesis by submerged aquatic plants have been carried out on detached leaves and algal thalli. These may experience very different environmental conditions than entire communities of submerged plants or plant dominated ecosystems, which have rarely been examined (e.g., Sand-Jensen et al., 2007 ; Christensen et al., 2013 ). We thus recommend undertaking studies on communities and ecosystems because they may reveal very different principles of regulation of greater relevance for the ecology and natural performance of submerged aquatic plants, as well as survival of terrestrial species during overland floods.

With the present review, we describe the general background and the recent advances in underwater photosynthesis of phytoelements (shoots, excised thalli, or leaf portions), communities, and plant dominated aquatic ecosystems and present contemporary methods tailor made to quantify photosynthesis and carbon fixation under water.

Underwater Photosynthesis

Challenges under water – reduced gas diffusion and light penetration.

The 10 4 -fold lower diffusion coefficient of gases in water, compared with in air, presents a major challenge to submerged plants ( Armstrong, 1979 ; Maberly and Madsen, 2002 ). Diffusive boundary layers (DBL) develop on all surfaces and their thickness adjacent to leaves in water is of the same order of magnitude as those for leaves in air ( Vogel, 1994 ; Raven and Hurd, 2012 ). Although the transport distance for gases across the DBL is similar, the much lower diffusion coefficient in water results in a 10 4 -fold lower flux for the same concentration gradient and thus the DBL constitutes a much larger proportion of the total resistance to gas exchange for leaves under water than in air ( Maberly and Madsen, 2002 ). The “bottleneck effect” of the DBL on underwater gas exchange was demonstrated in a study of four submerged aquatic species, where the DBL accounted for 90% of the total resistance to carbon fixation ( Black et al., 1981 ). Hence, inorganic carbon limitation of photosynthesis is a much more common and prominent feature for aquatic than terrestrial leaves ( Stirling et al., 1997 ). On average, the underwater photosynthesis increased threefold in a study of 14 submerged freshwater species at saturating relative to ambient supply of DIC supply ( Nielsen and Sand-Jensen, 1989 ). The immediate acclimation of photosynthesis of five fast growing annual terrestrial species by doubling of atmospheric CO 2 was 1.6- to 2.1-fold while the average increase of relative growth rate over 56 day was 1.25-fold and only significant for one of the five species ( Stirling et al., 1998 ).

The restricted gas exchange under water has resulted in evolution of a suite of adaptive features in aquatic leaves and macroalgal thalli to reduce the influence of DBL on the exchange of O 2 and CO 2 , including the supplementary use of HCO 3 - (bicarbonate) ( Sculthorpe, 1967 ; Maberly, 1990 ; Colmer et al., 2011 ). In seawater and in many freshwaters, the pool of HCO 3 - is several fold higher than of CO 2 and thus presents an attractive alternative to CO 2 . Use of HCO 3 - has evolved many times among unicellular algae, macroalgae, and angiosperms in freshwater and marine environments ( Raven and Hurd, 2012 ) and can involve direct uptake into the cells or external conversion to CO 2 in the DBL catalyzed by surface bound carbonic anhydrase and/or extrusion of protons in acids bands (charophytes; Lucas and Smith, 1973 ) or lower leaf surfaces in e.g., species of Potamogeton and Elodea ( Prins et al., 1980 ) often with precipitation of CaCO 3 on the alkaline upper leaf surfaces (Figure 1 B). While the DBL reduces the direct HCO 3 - flux to the leaf surface, the “stagnant” layer is required to forming high CO 2 concentrations by acid titration of HCO 3 - ( Helder, 1985 ). Use of HCO 3 - is prominent for marine macroalgae and seagrasses, freshwater charophytes, and in 50% of 80 tested species of freshwater angiosperms ( Sand-Jensen and Gordon, 1984 ; Maberly and Madsen, 2002 ), but the ability is absent among mosses and pteridophytes. Also 12 amphibious species alternating between emergent and submerged forms in Danish lowland streams relied solely on CO 2 use although high HCO 3 - concentrations may still benefit photosynthesis by stabilizing pH and permitting rapid uncatalyzed replenishment of the CO 2 consumed ( Maberly, 1990 ). The proportion of HCO 3 - users among angiosperm species in lakes increases significantly with alkalinity and, thus, availability of HCO 3 - ( Maberly and Madsen, 2002 ) in accordance with the increasing advantage of HCO 3 - use for photosynthesis and growth. Assuming for simplicity a 10-fold higher apparent preference for CO 2 than HCO 3 - by leaves in alkaline water containing 0.015 mmol L −1 CO 2 and 1.5 mmol L −1 HCO 3 - , the supply rate of HCO 3 - would be 10-fold higher than that of CO 2 . In softwater containing only 0.15 mmol L −1 HCO 3 - the supply rate of the two carbon species would be the same. Terrestrial plant species lack these adaptive features for aquatic life, and when underwater their leaves show dramatically reduced net photosynthesis ( Sand-Jensen et al., 1992 ; Nielsen, 1993 ) and dark respiration ( Colmer and Pedersen, 2008 ; Pedersen et al., 2009 ). Thus, 13 terrestrial species submerged in CO 2 rich stream water were unable to use HCO 3 - and median rates of underwater net photosynthesis were sevenfold lower than of 10 permanently submerged stream plants and the terrestrial species were unable to support substantial growth ( Sand-Jensen et al., 1992 ; Figure 2 ).

Figure 2. Rates of underwater net photosynthesis (nmol O 2 g −1 DM s −1 ) in ambient [(A), 90–400 μmol CO 2 L −1 ] and CO 2 enriched water [(B), 800 μmol CO 2 L −1 ] and final pH from a pH drift experiment; Section “pH Drift Approach to Establish CO 2 Compensation Points” (C) in four groups of life forms defined in Sculthorpe (1967) . The box whisker plot indicates the range of the observations (bars), 50% of the observations (boxes) and the median (horizontal line). The amount of free CO 2 present at median final pH of the four groups was 6.20, 6.00, 4.80, and 0.04 μmol L −1 , respectively. Leaves of aquatic plants species ( n = 10), heterophyllous amphibious plants ( n = 5), homophyllous amphibious plants ( n = 7) were all produced under water while leaves of terrestrial plant species were formed in air ( n = 12). Letters indicate significant differences ( P < 0.05); Tukey test. Data from Sand-Jensen et al. (1992) , with measurements taken at 15°C.

The extraction capacity of the DIC pool is only some 1–4% for obligate CO 2 users while it is typically 40–70% among HCO 3 - users; 16 of 19 species ( Madsen and Sand-Jensen, 1991 ). This is because of the ability of HCO 3 - users to continue photosynthesizing despite very high external pH and low DIC in the water. In vegetation rich water bodies of high pH, HCO 3 - users can eventually out compete all obligate CO 2 users ( Sand-Jensen et al., 2010 ). Submerged aquatic plants unable to use HCO 3 - typically have final pH’s in the external medium of the order of 8.6–9.8 in alkaline solutions and final CO 2 concentrations equivalent to CO 2 compensation points of 2–10 μmol L −1 , while active HCO 3 - users typically have final pH’s of 9.8–11.0 and final CO 2 concentrations mostly below 0.3 μmol L −1 ( Sand-Jensen et al., 1992 ). For a large collection of stream plants, median final CO 2 values among the supposedly obligate CO 2 users were 6.0 μmol L −1 for homophyllous and 4.8 μmol L −1 for heterophyllous amphibious plants, within the typical range of CO 2 compensation points, while the median final CO 2 concentration for the putative HCO 3 - users was only 0.04 μmol L −1 reflecting the supplementary use of HCO 3 - (Figure 2 ). Heterophyllous amphibious species form morphologically and anatomically distinct leaf types under water as compared to in air ( Sculthorpe, 1967 ). The underwater leaf forms are an acclimation that enhances underwater gas exchange ( Sand-Jensen et al., 1992 ; Colmer et al., 2011 ).

Photosynthesis of submerged aquatic plants and flooded terrestrial plants can also be severely limited by light ( Kirk, 1994 ). In water, light is exponentially attenuated with depth following the equation: I Z = I 0 ( 1 - f ) e - Z ε ; where I z is the available irradiance at a given depth ( z ), I 0 is the irradiance at the surface, and ε is the attenuation coefficient. The proportional reflection and back scattering at the water surface (f) is variable but typically about 0.1 such that the proportion of down welling irradiance is 0.9 ( Kirk, 1994 ). The attenuation coefficient of pure water averaged across the photosynthetic spectrum is about 0.03 m −1 , so in ultra clear water such as oligotrophic oceanic water, rooted plants could grow as deep as 70 m with 10% of surface irradiance still being available, which happens to be the approximate lower depth limit of seagrasses ( Duarte, 1991 ). In most cases, however, colored dissolved organic matter (CDOM), pigments in planktonic algae and suspended particles, together reduce light penetration much more profoundly ( Staehr et al., 2012b ). Because freshwaters compared with marine waters are typically richer in nutrients, phytoplankton, CDOM exported from land and particles suspended from shallow sediments, attenuation coefficients typically range from 0.3 to 10 m −1 and thus have lower depth limits of rooted plants from 7 m to only 0.2 m ( Middelboe and Markager, 1997 ). Flooding after heavy rain is commonly associated with erosion, high particle loads and high release of CDOM from inundated terrestrial soils. Flooded terrestrial plants can, therefore, experience extreme shading corresponding to attenuation coefficients between 1 and 8 m −1 ( Parolin, 2009 ) making light limitation also of terrestrial plants a prominent feature during flooding events ( Colmer et al., 2011 ).

Underwater Photosynthesis in Submerged Aquatic Plants and Recent Advances

The net process of photosynthesis (Eq. 1) is often described simply as the fixation of CO 2 (or HCO 3 - in water; Eq. 2) catalyzed by several enzymes, including Rubisco, driven by light and resulting in production of organic matter, O 2 and OH − :

The rate of the process can be determined by the production of O 2 and new organic matter (e.g., by isotopic tracing with 13 C and 14 C) and the consumption of CO 2 , HCO 3 - , or more generally the pool of DIC: CO 2 , HCO 3 - and CO 3 2 - . Photosynthesis is an alkalinization process as reflected by the release of OH − in Eq. 2 and the equivalent consumption of CO 2 and protons in Eq. 1 (i.e., CO 2 + H 2 O ↔ H + + HCO 3 - ) such that photosynthesis can be determined by the pH increase when converted to DIC consumption accounting for the buffer capacity [mainly due to carbonate alkalinity (CA), See, The CO 2 Equilibria in Water].

In charophytic macroalgae, use of HCO 3 - in photosynthesis can be closely coupled stochiometrically to carbonate precipitation ( McConnaughey, 1991 ):

This process is pH neutral because conversion of HCO 3 - to CO 3 2 - generates the necessary proton for conversion of HCO 3 - to CO 2 for assimilation. Thus, HCO 3 - is equally divided between production of organic matter and CaCO 3 and the photosynthetic quotient (PQ: mol O 2 evolved mol −1 DIC consumed) is only about 0.5 compared with the typical value of 1.0 in Eqs 1 and 2. The active processes are apparently active antiport of H + and Ca 2+ in the acid bands and Ca 2+ extrusion in the alkaline bands resulting in gradual accumulation of CaCO 3 from inside the carbonate layer ( McConnaughey, 1991 ). Carbonate precipitation closely coupled to photosynthesis is also found in coralline red algae, several green algae, and numerous freshwater angiosperms developing polar leaves with high pH and carbonate precipitation being confined to the upper leaf surfaces ( Raven and Hurd, 2012 ). However, it remains to be tested whether active Ca 2+ extrusion is involved in angiosperm use of HCO 3 - as in charophytes ( Prins et al., 1980 ). Even though photosynthesis and carbonate precipitation may not be closely coupled, the alkalinization process in Eqs 1 and 2 may still result in carbonate precipitation on leaf surfaces (Figure 1 B) or in the surrounding water because of increase of pH and CO 3 2 - , though with a variable ratio to the fixation of CO 2 in organic matter. Consequently, O 2 evolution is a more reliable measure of underwater photosynthesis, while DIC use and production of organic matter and carbonate are essential parameters in the analysis of plant growth and carbon dynamics in ecosystems and on regional and global scales ( McConnaughey et al., 1994 ; Cole et al., 2007 ). While coupled calcification photosynthesis leads to extensive drawdown of DIC and sediment accumulation of organic carbon and carbonates, carbonate formation per se generates H + tending to reduce pH and increase CO 2 . During geological periods of intense formation of coral reefs, CO 2 concentrations are suggested to have increased in the ocean and the atmosphere ( Opdyke and Walker, 1992 ). The coupled photosynthesis calcification process has three major ecological or physiological implications: (i) in coral and coralline algae carbonates are directly used to build up the structural skeleton, (ii) in all phototrophs, surface precipitates will protect them against grazing animals, and (iii) calcification will prevent alkalinization during intensive photosynthesis which could otherwise have led to such high pH levels that tissues are damaged and photosynthesis is severely inhibited.

The photosynthetic capacity under optimum light, temperature, and DIC conditions varies among species and within species depending on their investment in active transport processes and catalytic machinery. On dry mass basis, maximum rates of photosynthesis of detached leaves of submerged aquatic plants from lakes typically vary 25-fold and dark respiration only fourfold between slow-growing, oligotrophic isoetid species and fast growing, eutrophic elodeid species ( Nielsen and Sand-Jensen, 1989 ). Photosynthetic rates per unit dry mass increase significantly with reduced leaf thickness, higher relative surface area, and higher concentrations of pigments and nitrogen in structural and catalytic proteins, including Rubisco ( Madsen et al., 1993 ). Because metabolism on a dry mass basis increases with declining leaf thickness, photosynthesis expressed per surface area only varies eightfold among species ( Nielsen and Sand-Jensen, 1989 ). A comparison with terrestrial leaves characterizes the aquatic leaves by their lower chlorophyll and Rubisco concentrations and lower photosynthetic rates per surface area mainly due to the thin leaves of most aquatic species. This finding has been interpreted by Maberly and Madsen (2002) as a result of selection of submerged plants to match the low rates of carbon influx predominantly because of high transport resistance. Thin submerged leaves with chloroplasts in epidermal layers will also increase the cost effectiveness of light use and can also be regarded as a particular advantage in a low light aquatic environment with no risk of desiccation damage to the epidermal layers.

Realized rates of underwater photosynthesis for a given plant tissue varies from zero at compensation levels for light and DIC to maximum rates at saturating light and DIC. Light and DIC levels required to saturate photosynthesis increase with temperature and are highly dependent on the extent of self shading and, therefore, the scale of analysis of either detached leaves, individuals or populations (See Underwater Photosynthesis – Approaches and Methods). On a daily level, light limitation takes place early in the morning (low light but plenty of CO 2 , Figure 3 ) and co-limitation of both light and CO 2 takes place late in the afternoon where also CO 2 is low (Figure 3 ). On a seasonal level, light limitation is present from late autumn to early spring outside the tropical regions. Populations in deep or turbid waters and dense populations with high self shading face permanent light limitation. Photosynthetic rates at saturating light and DIC increase with temperature due to stimulation of enzyme activity up to an optimum level depending on the adaptation and acclimation of the species but are usually located between 25 and 32°C for temperate submerged aquatic plants ( Santamaría and van Vierssen, 1997 ). The temperature exponent gradually declines and reaches zero as temperature approaches the optimum temperature and it turns negative above the optimum due to increasing influence of photorespiration ( Long, 1991 ) and risk of damage of macromolecules, membranes, and structural organization of membrane proteins ( Johnson et al., 1974 ; Lambers et al., 2008 ). Respiration continues to increase up to higher temperatures than photosynthesis resulting in proportionally greater losses of organic matter and optima for growth being located at lower temperature than optima for photosynthesis ( Olesen and Madsen, 2000 ; Pilon and Santamaría, 2001 ).

Figure 3. Diurnal fluctuations in surface irradiance and water temperature (A), O 2 partial pressure (B), and CO 2 concentration and pH (C) in the water of a vegetation rich pond in Western Australia . The pond was densely vegetated by Meionectes brownii which in late afternoon had extracted CO 2 from the water down to below air equilibrium (<15 μmol L −1 ). During the night, total system respiration consumed O 2 (pO 2 dropped to 2.5 kPa early in the morning) and CO 2 rose to 550 μmol L −1 . Data extracted from Rich et al. (2013) .

Ninety nine percent of all studies of aquatic photosynthesis have been carried out with detached leaves or thalli and this selectiveness influences the perception of the regulation of aquatic photosynthesis ( Sand-Jensen and Krause-Jensen, 1997 ). The influence of light, DIC, and temperature on underwater photosynthesis show mutual interdependencies and are, moreover, strongly dependent on the spatial scale. From detached phytoelements to closed communities, light compensation points typically increase three- to eightfold and light saturation levels increase from 200 to 400 μmol m −2 s −1 to more than the maximum irradiances at noon of about 1500 μmol m −2 s −1 (Table 1 ; Sand-Jensen et al., 2007 ). The stimulation of photosynthesis in alkaline water by rising CO 2 concentrations from 20 (close to air equilibrium) to 250 μmol L −1 (more than 10-fold above air equilibrium) is about ninefold for detached leaves and only 1.9- to 2.5-fold for dense communities of freshwater CO 2 users while for efficient HCO 3 - users the CO 2 stimulation is only about twofold for individual leaves and insignificant for dense communities (Table 2 ; Sand-Jensen et al., 2007 ). Open communities of less self shading take an immediate position between detached individual leaves and dense communities. Profound self shading and light limitation of photosynthesis in dense aquatic communities imply that the influence of temperature and inorganic carbon supply is smaller than observed for well illuminated phytoelements. The full scale influence of temperature and CO 2 on community photosynthesis is confined to tissues in the upper part of the canopy receiving irradiances above light saturation.

Table 1 . Photosynthetic parameters for thallus segments and communities of Fucus serratu s of varying leaf area index (LAI) .

Table 2 . Increase (x-fold) of maximum gross production of O 2 at high (250 μmol L − 1 ) relative to low (20 μmol L − 1 ) CO 2 concentration in alkaline water (5000 μmol L − 1 DIC) of leaves and freshwater plant communities at two densities (LAI; 2 or 10 m 2 m − 2 ) .

Up scaling of metabolic analyses from communities of submerged aquatic plants to entire ecosystems dominated by rooted plants have only been done in a few instances. Kelly et al. (1983) studied a shallow, densely vegetated stream (Gryde Stream, Denmark) by open water O 2 measurements and confirmed that incoming irradiance was the main determinant of daily and seasonal variations of underwater photosynthesis which was only light saturated for a few hours at noon on clear summer days. The high CO 2 concentrations (typically 10-fold air equilibrium) in lowland streams is a prerequisite for the high photosaturated rates and strong light dependency of submerged plants in general and CO 2 users in particular ( Sand-Jensen and Frost-Christensen, 1998 ). With natural CO 2 concentrations close to air equilibrium, as observed in most lakes and ponds and in streams in the afternoon after several hours of planktonic photosynthesis ( Sand-Jensen and Frost-Christensen, 1998 ; Christensen et al., 2013 ), CO 2 plays a stronger regulatory role for photosynthesis particularly in open plant stands of low self shading ( Sand-Jensen and Frost-Christensen, 1998 ). Moreover, the species rich group of terrestrial plants in lowland streams would be unable to survive if the water had not been greatly supersaturated as their CO 2 compensation points resemble or exceed the CO 2 concentrations at air equilibrium ( Sand-Jensen and Frost-Christensen, 1999 ). Recent use of open water measurements of O 2 and pH in shallow, alkaline ponds dominated entirely by charophytes documents that high biomass densities in late summer are attained by sustained slow growth over the preceding 3 months at very low nutrients concentrations in the water, and that daily photosynthesis is mostly limited by light (Figure 4 ) and only briefly by DIC at high pH (>9.5), and with virtually no CO 2 available in the afternoon ( Christensen et al., 2013 ). Only submerged aquatic plants capable of using HCO 3 - and concentrating CO 2 internally at the site of Rubisco can thrive in this environment ( Sand-Jensen et al., 2010 ). Plant species forming dense communities in shallow ponds must also be able to tolerate substantial diurnal variations in temperature (e.g., 18–32°C) and O 2 (hypoxic to twice air equilibrium) ( Christensen et al., 2013 ). Daily photosynthesis and respiration were high in the pond and closely interrelated showing that newly produced organic matter was mostly rapidly respired by plants and bacteria.

Figure 4. Relationship between daily surface irradiance and community gross primary production in a shallow pond in Öland, Sweden over a daily time course of 8 days with naturally variable light conditions (each data point represents 1 day) . The dominant submerged aquatic plant in the pond was, Chara virgata (data extracted from Christensen et al., 2013 ). Pearson correlation = 0.96, P < 0.001.

Overall, the analyses of individual phytoelements, communities, and ecosystems confirm that the relative roles of light and DIC for determining photosynthesis are closely interrelated and highly dependent on plant density and species affinities for CO 2 and HCO 3 - . Maximum photosynthetic rates under light and inorganic carbon saturation are quite variable both between and within species depending on selected strategies and the investment in catalytic machinery coupled to supply of resources (e.g., nutrients). While photosaturated photosynthetic rates are strongly dependent on species, acclimation, and temperature, light limited rates are rather temperature independent and relatively similar among species ( Frost-Christensen and Sand-Jensen, 1992 ). As the importance of light limitation for community photosynthesis increases in dense plant stands, the influence of species, temperature, and DIC supply decline and enable us to predict community photosynthesis primarily from the overall distribution and absorptance of light in the canopy ( Binzer and Sand-Jensen, 2002a ; b; Binzer et al., 2006 ).

Recent Advances in Underwater Photosynthesis in Terrestrial Wetland Plants

Terrestrial wetland plants grow in waterlogged soils and/or sediments with shallow standing water, so that a large proportion of the shoot is in contact with air. So, aerial photosynthesis predominates but these plants can experience episodes of complete submergence during floods. Although much more tolerant of submergence than non-wetland terrestrial species, submergence is regarded as a serious abiotic stress for terrestrial wetland plants, but species (and genotypes within a species) differ markedly in submergence tolerance ( Bailey-Serres and Voesenek, 2008 ; Colmer and Voesenek, 2009 ). The impeded gas exchange under water restricts respiration and photosynthesis (See Challenges Under Water – Reduced Gas Diffusion and Light Penetration); photosynthesis can also be limited by low light when submerged (See Challenges Under Water – Reduced Gas Diffusion and Light Penetration and Underwater Photosynthesis in Submerged Aquatic plants and Recent Advances). Thus, submergence disrupts energy metabolism of terrestrial plant species as a result of a reduced O 2 supply (at least during the night, in some tissues) and/or diminished carbohydrate status because of the restricted photosynthesis when under water.

Terrestrial wetland species lack most of the adaptive leaf features for inorganic carbon acquisition for photosynthesis as described in Section “Underwater Photosynthesis in Submerged Aquatic Plants and Recent Advances” for aquatic and acclimated amphibious plants. Thus, when compared with leaves of aquatic plants, those of terrestrial plants generally have larger overall apparent resistance to diffusion of CO 2 from the bulk medium to chloroplasts, so that slow CO 2 uptake restricts underwater photosynthesis. Underwater photosynthesis by leaves of terrestrial wetland species is lower than that achieved by aquatic species, when compared per unit of leaf dry mass ( Sand-Jensen et al., 1992 ; Colmer et al., 2011 ).

The few studies available show, however, that the low photosynthesis when under water enhances survival of submerged terrestrial plants ( Vervuren et al., 1999 ; Mommer et al., 2006b ; Vashisht et al., 2011 ). Both the sugars and O 2 produced would likely contribute to enhanced survival when submerged ( Mommer and Visser, 2005 ), and in the case of sugars especially when submergence lasts more than a few days and internal carbohydrates become depleted. Depletion of carbohydrates during submergence is considered a major factor influencing survival of submerged rice ( Setter and Laureles, 1996 ) and determining recovery following desubmergence and ultimately grain yield in flood-prone areas ( Bailey-Serres et al., 2010 ; Mackill et al., 2012 ). The O 2 produced in photosynthesis can travel from leaves to roots via aerenchyma, and so this endogenously produced O 2 improves the internal aeration of submerged plants (e.g., rice; Pedersen et al., 2009 ; Winkel et al., 2013 ).

A recent review ( Colmer et al., 2011 ) highlighted there are few studies of underwater photosynthesis by terrestrial wetland plants, and few of these compared rates underwater with those achieved by leaves in air. Similarly, a quantitative understanding of the potential role of underwater photosynthesis to whole plant carbon budgets during submergence seems to be lacking for terrestrial wetland species, whereas carbon budgets for several aquatic species (e.g., van der Bijl et al., 1989 ) and systems (e.g., Christensen et al., 2013 ) have been evaluated. For some terrestrial wetland species, only a few crude leaf level estimates of carbon budgets have been considered (e.g., in Colmer and Pedersen, 2008 ), but the potential contribution of underwater photosynthesis to carbon gain was demonstrated in growth studies of completely submerged rice, albeit under controlled conditions (e.g., Pedersen et al., 2009 ). Studies of whole plant carbon budgets in field conditions are generally lacking, even understanding of this aspect for the important wetland crop rice submerged in various field scenarios appears to be incomplete.

Detailed studies of underwater photosynthesis of terrestrial wetland species have focused on production and performance of submergence acclimated leaves. New leaves produced when under water by some terrestrial wetland species are better acclimated for underwater photosynthesis than the aerial leaves ( Mommer et al., 2007 ). The acclimated leaves have a thin cuticle and overall are also thinner and of less breath ( Mommer and Visser, 2005 ). These morphological and anatomical differences as compared with the usual leaves produced in air, reduce the resistance to CO 2 (and O 2 ) diffusion between the bulk medium and chloroplasts in submerged leaves, owing to narrower DBLs (suggested by Colmer et al., 2011 ), lower cuticle resistance ( Mommer et al., 2006b ), and shorter internal diffusion path lengths ( Mommer et al., 2006a ). However, although a reduced cuticle that enhanced underwater gas exchange occurs in several terrestrial wetland species (few species have been evaluated to date), the magnitude of the reduction in apparent resistance to gas exchange with the medium was not correlated with submergence tolerance for the species tested ( Mommer et al., 2007 ), highlighting the need for further experimental investigations.

Some recent work on underwater photosynthesis by submerged terrestrial wetland plants has evaluated the contribution of gas films on superhydrophobic leaf surfaces to gas exchange with floodwaters. Leaf surface hydrophobicity (i.e. water repellence) is a feature that sheds off water in wet aerial environments ( Smith and McClean, 1989 ; Brewer and Smith, 1997 ) and promotes “self cleansing,” enhancing leaf performance and reputably lowering susceptibility to pathogens ( Neinhuis and Barthlott, 1997 ). Some terrestrial wetland species have super hydrophobic leaves that when submerged retain a gas film, e.g., rice ( Raskin and Kende, 1983 ) and Phragmites australis and others ( Colmer and Pedersen, 2008 ). Gas films enhance CO 2 uptake for underwater photosynthesis ( Raskin and Kende, 1983 ; Colmer and Pedersen, 2008 ; Pedersen et al., 2009 ) and O 2 entry for respiration in darkness ( Colmer and Pedersen, 2008 ; Pedersen and Colmer, 2012 ). The enhancement by leaf gas films of CO 2 uptake (in light) and of O 2 (in darkness) was demonstrated by the marked declines in underwater photosynthesis and respiration when the films were experimentally removed ( Colmer and Pedersen, 2008 ; Pedersen et al., 2009 ; Pedersen and Colmer, 2012 ). In addition, leaves produced in air by terrestrial wetland species that did not form gas films when submerged (i.e. leaves of these species were not sufficiently hydrophobic), had lower rates of underwater photosynthesis than those that did form gas films ( Colmer and Pedersen, 2008 ; Colmer et al., 2011 ). As one example, leaf segments of rice with dissolved CO 2 set as in a field pond and with underwater photosynthesis measured as described in Section “The Rotating Wheel Incubator,” showed rates four- to fivefold higher for leaf segments with intact gas films compared to those with the films experimentally removed ( Winkel et al., 2013 ). Moreover, a field study using in situ monitoring of O 2 in rhizomes of Spartina anglica demonstrated the benefit of having leaf gas films to internal aeration during complete submergence, both during day and night tides ( Winkel et al., 2011 ). Summing up, leaf gas films enhance underwater photosynthesis and internal aeration of some terrestrial wetland plants when submerged, with benefits also demonstrated to growth when submerged in controlled experiments (e.g., rice; Pedersen et al., 2009 ).

Underwater Photosynthesis – Approaches and Methods

Conventional infrared gas analyzer (IRGA) systems following CO 2 exchange in air do not work under water, so dedicated measuring systems are required to quantify underwater net photosynthesis and dark respiration. DIC can be measured by injection of small aliquots of water into concentrated acid in a bubble chamber purged with gaseous N 2 carrying the released CO 2 into an IRGA ( Vermaat and Sand-Jensen, 1987 ). However, photosynthesis measurements based on DIC determinations are thus based on discrete measurements at selected times and can be complicated because of the large and variable combined pool of DIC in water (See Underwater Photosynthesis in Submerged Aquatic plants and Recent Advances and The CO 2 Equilibria in Water). Indirect methods to track DIC changes can be based on continuous measurements of pH in solution ( Maberly, 1996 ). The DIC technique to measure photosynthess has potential errors if: (i) DIC is removed by external carbonate precipitation, (ii) internal DIC accumulates in tissues or colony gels, (iii) DIC dissolution of solid carbonates occurs, or (iv) DIC is released from internal pools ( McConnaughey et al., 1994 ; Sand-Jensen et al., 2009 ). External measurements of pH to estimate DIC changes have the same potential errors as above and, moreover, also due to direct exchange of protons from tissues not always being closely coupled to DIC exchange. Therefore, most methods for studies of underwater net photosynthesis are based upon O 2 detection.

In contrast to gas exchange measurements of photosynthesis by leaves in air using open systems and CO 2 detection, underwater measurements commonly use closed systems and detection of O 2 . In addition to the rationale for O 2 detection described in the preceding paragraph, O 2 detection also enables measurements in waters of substantially different DIC concentrations (e.g., softwater lakes up to 100 μmol L −1 , ocean approximately 2000 μmol L −1 and hardwater lakes up to 10000 μmol L −1 ). The drawback of closed systems is that these are non-steady-state (i.e. DIC declining and O 2 increasing with time). Use of open systems with O 2 detection is constrained by reliable continuous detection of differences in O 2 concentrations between incoming and outgoing solutions from an appropriate chamber.

Changes in O 2 concentration over time are straightforward to measure with Clark type amperometric electrodes or more recently by use of O 2 sensitive optodes. Oxygen partial pressure (pO 2 ) or dissolved O 2 can be continuously monitored in water with an accuracy of 0.01 kPa or 0.2 μmol L −1 ( Strickland and Parsons, 1972 ). Photosynthesis determined from changes in O 2 and DIC pools dissolved in the surrounding water requires that those are much greater than changes in such pools within the plant tissue ( Sand-Jensen and Prahl, 1982 ). This is best achieved by having large incubation volumes relative to plant volumes. Alternatively, changes in tissue pools can be measured ( Sand-Jensen et al., 2005 ) or be deduced by establishment of true steady state where tissue concentrations remain constant or quasi steady state where tissue concentrations changes proportionally to external concentrations ( Sand-Jensen and Prahl, 1982 ). Measurements of underwater photosynthesis based upon O 2 evolution can include great error when plants with highly porous tissues (perhaps variable in volume and having much higher “solubility” of O 2 than water; See Medium and Tissue) are incubated in small chambers ( Hartman and Brown, 1967 ; Richardson et al., 1984 ). On the other hand, measurements of underwater photosynthesis based upon changes in DIC can include extreme error when plant tissues (or colony matrices in the case of algae and cyanobacteria) hold very large pools of DIC that do not change in concert with those in the surrounding water. For example, DIC in the colony gel of Nostoc zetterstedtii continues to support photosynthesis after water pools have been exhausted, and in darkness respiratory CO 2 replenishes this internal pool before being released to the water ( Sand-Jensen et al., 2009 ).

Measurements of radioactive labeling of the DIC pool with 14 C and the use of pulse amplitude modulated (PAM) fluorometry are also methods to measure photosynthetic performance under water; these technique are beyond the focus of the present paper so readers are referred to e.g., Adams et al. (1978) or Kemp et al. (1986) for methods on 14 C and to Silva et al. (2009) or Suggett et al. (2011) and chapters therein for PAM approaches.

The CO 2 Equilibria in Water

Understanding the chemistry of dissolved DIC and the proportional changes in its three constituents (CO 2 , HCO 3 - and CO 3 2 - ) depending on ionic strength, temperature, and primarily pH ( Mackereth et al., 1978 ) is essential because it determines the availability of the preferred CO 2 source and the supplementary HCO 3 - source for underwater net photosynthesis. When CO 2 dissolves in water, the following equilibrium is established:

CO 2 ’s reaction with water (H 2 O) forming carbonic acid (H 2 CO 3 ) is a time dependent process which in some organisms is catalyzed by the enzyme carbonic anhydrase. H 2 CO 3 can dissociate immediately into a proton (H + ) and bicarbonate ( HCO 3 - ) so the dissolution of CO 2 into water causes pH to drop. At high pH, HCO 3 - can further dissociate into a second H + and carbonate ( CO 3 2 - ). The relative distribution of the three main inorganic carbon species with pH is shown (Figure 5 ). The pKa 1 is 6.532 and is referred to as the apparent pKa 1 as only little CO 2 is converted into carbonic acid (hence the brackets in Eq. 4) while the majority remains in solution as CO 2 (aq) also referred to as free CO 2 ; pKa 2 is 10.329 ( Schwarzenbach and Meier, 1958 ; Stumm and Morgan, 1996 ; Gutz, 2012 ). Below pH 6, most of the DIC is present as CO 2 , which is usually more readily used for underwater photosynthesis than HCO 3 - . Between pH 7 and 10, HCO 3 - dominates, a carbon species that can be used as an additional carbon source among species in most taxonomic groups of aquatic plants except for pteridopytes and mosses ( Raven and Hurd, 2012 ). Only at pH higher than 10, a significant proportion of the DIC is in the form of CO 3 2 - which apparently is not taken up by any phototrophs in ionic form but can perhaps be made available in acid zones on plant surfaces by back titration with released protons (conversion toward the left in Eq. 4).

Figure 5. Relative speciation (%) of carbon dioxide (CO 2 ), bicarbonate ( HCO 3 - ), and carbonate ( CO 3 2 - ) in water as a function of pH . The example given is at 20°C and electrical conductivity of 250 μS cm −1 . Data were calculated using Gutz (2012) with the apparent p K 1 = 6.532 and p K 2 = 10.329 ( Schwarzenbach and Meier, 1958 ).

In freshwaters and seawater, the alkalinity (sum of alkaline ions buffering added H + ; units in mequiv. L −1 or mmol L −1 for monovalent HCO 3 - in water which is in air equilibrium of negligible OH − and CO 3 2 - ) is almost entirely controlled by the carbonate systems with insignificant contribution from silicate and phosphate, and with some contribution by borate in seawater. At pH above 9, OH − has a significant contribution to alkalinity being 0.074 mmol L −1 at pH 10 and 0.74 mmol L −1 at pH 11 at an alkalinity of 2 mmol equivalents L −1 (Table 3 ). It is thus convenient to distinguish between the total alkalinity (TA) and the CA ( Dickson, 1981 ; Stumm and Morgan, 1996 ). The chemical species contributions to the two alkalinities are:

Table 3 . Distribution of DIC, CO 2 , HCO 3 - , CO 3 2 - , and OH − as a function of pH at constant total alkalinity of 2 mmol H + equivalents L − 1 at 20°C .

Purging an aqueous solution with pure CO 2 alters the CA through the addition of ionic carbon species and also through pH related shifts in the partitioning of carbon species already present in the solution (Eqs 4 and 5). However, the TA is not affected by bubbling with CO 2 as every negatively charged ion is balanced by a proton (Eq. 6). For example, water fresh from the tap often contains CO 2 above air equilibrium and so bringing it to equilibrium by purging with atmospheric air would thus lower pCO 2 until a new equilibrium has been reached. According to Eq. 5, CA would decrease slightly as both CO 3 2 - and HCO 3 - decrease equivalent to the rise of OH − and pH.

For experimental purposes, an aqueous photosynthesis solution is usually prepared with a certain amount of DIC and then pH is adjusted with acid or base to that required to achieve the desired concentration of “free” (i.e. dissolved) CO 2 and HCO 3 - . Table 3 lists the relationship between pH and amounts of CO 2 , HCO 3 - , CO 3 2 - , and OH − at 20°C and a fixed TA, calculated from Gutz (2012) . The examples provided in the sections below demonstrate how to apply all the above information in practice.

In the next sections (See “The Rotating Wheel Incubator” to “The Open Natural System”) we describe methods in use for measurements of underwater photosynthesis. The methods scale from phytoelements to communities. The approaches involve laboratory and field techniques and so have different levels of control of key environmental variables influencing photosynthesis.

The Rotating Wheel Incubator

Principle: Leaf samples or algal thalli are incubated in glass vials of a known concentration of CO 2 in an aqueous medium, and the sealed vials of known volume are rotated on an incubator under well defined light and temperature conditions. O 2 produced during incubation is measured by an electrode/optode and underwater net photosynthesis can be calculated based on e.g., leaf area, fresh mass, dry mass, and/or chlorophyll. Alternatively, consumption of DIC can be used as a photosynthetic measure. Incubation in darkness provides data on dark respiration.

Medium and tissue

The choice of medium is basically between an artificial medium with a well defined ion and gas composition or ambient water with the ion and gas composition of natural habitats (essential chemical parameters such as pH, DIC, and alkalinity should be characterized). An example of an artificial medium is the Smart and Barko (1985) general purpose culture medium. This medium contains (mmol L −1 ) 0.62 Ca 2+ , 0.28 Mg 2+ , 0.28 SO 4 2−, and 1.24 Cl − and KHCO 3 (sometimes mixed with NaHCO 3 ) is used to generate the required DIC. HCl, NaOH (or KOH), atmospheric air or gas mixtures of known pCO 2 can be used to adjust pH to a required value based on the desired amount of free CO 2 . Since all incubations are short term, there are no micro nutrients or vitamins in this medium. Some studies have also used submergence solutions or “ambient” water from streams or lakes in order to establish a rate of photosynthesis under specific conditions ( Sand-Jensen et al., 1992 ; Nielsen, 1993 ; Sand-Jensen and Frost-Christensen, 1998 ) and these can also be adjusted to predefined pH, CO 2 and/or O 2 levels. Any production of O 2 by microalgae or consumption by microbial organisms in ambient water is accounted for in the blanks; micro-filtration of water is commonly used to remove background microflora.

Photorespiration, as previously demonstrated for rice ( Setter et al., 1989 ) and the aquatic pteridophyte, Isoetes australis , ( Pedersen et al., 2011 ), during incubation is a potential issue as the evolved O 2 is trapped in solution of the closed glass vial. The risk of photorespiration is increased during experiments at high temperature ( Long, 1991 ) and with very low DIC and CO 2 concentrations leading to low ratios of CO 2 to O 2 at the site of Rubisco ( Maberly and Spence, 1989 ; Sand-Jensen and Frost-Christensen, 1999 ). Therefore, the starting partial pressure of O 2 (pO 2 ) should be brought down to approximately 50% of air equilibrium, i.e., 10 kPa. This is sufficient to address the issue of photorespiration (provided that incubation do not last long periods so that O 2 produced increases above air equilibrium) and at the same time there is still enough O 2 in the medium to prevent tissue anoxia before photosynthesis starts producing O 2 ( Colmer and Pedersen, 2008 ). In practice, equal volumes of medium (including all ions) are bubbled with either air or N 2 . After mixing the two solutions, the pO 2 will be approximately 10 kPa and HCO 3 - can be added to the medium and pH adjusted accordingly to achieve the desired amount of free CO 2 (see example below).

In some situations, an organic buffer may be used to maintain a constant pH in the medium during incubation. In practice, however, HCO 3 - is a natural and often sufficient buffer in itself and we do not recommend using buffers if the CA is above 1 mmol L −1 as HCO 3 - would be sufficient to buffer against large pH fluctuations during incubation ( Sand-Jensen et al., 1992 ; Colmer and Pedersen, 2008 ). Moreover, organic buffers can also modify membrane porters and pH at plant surfaces modifying HCO 3 - use and influx/efflux of CO 2 ( Price and Badger, 1985 ; Larsson and Axelsson, 1999 ; Moulin et al., 2011 ). pH of the medium should be measured in a sample taken of the initial solution and then also in vials after incubations. With the ongoing advancement of optodes, pH may even be measured without opening the vials if applying pH sensitive microdots (See “O 2 Measurements” for description of O 2 sensitive microdots). If additional buffering is required, i.e. pH measurements after incubation show unacceptable drift in pH, then MES or TES buffers may be used, e.g., at a concentration of 5 mmol L −1 ( Pedersen et al., 2009 , 2010 ), though the possible influence of these buffers on HCO 3 - use must be kept in mind.

The vials (10–100 mL glass vials with ground glass stoppers) are filled with medium using a siphon. By siphoning the medium into the bottom of each vial, exchange of O 2 and particularly CO 2 with the atmosphere is minimized; prepare sufficient medium to flush the vials at least twice the volume, and fill the vials completely. An air bubble can hold 36-fold more O 2 as the same volume of deionized (DI) water at 25°C, so bubbles in the vials introduce significant error to the net photosynthesis measurements. A set of vials without tissue serves as blanks and is incubated along with the vials containing tissue samples in the rotating incubator. The blanks serve to provide the starting pO 2 in the vials and also to correct for any O 2 production or consumption (e.g., by algae, bacteria, or chemical processes) if ambient water is used as medium. Glass beads (Ø = 3–5 mm; two in each 25 mL vial) are added to each vial to provide mixing as the vials are rotating in the incubator.

The amount of tissue added to each vial depends on the activity of the tissue, the amount of DIC and free CO 2 , the light level (PAR), and the temperature. At saturating light and CO 2 levels and at 25°C, 0.5 mg fresh mass mL −1 medium is often a good choice as this will result in a rise of pO 2 by approximately 2–5 kPa within an hour of incubation providing reproducible and accurate determination of O 2 regardless of the technique employed (see below). However, both microelectrodes and optodes have a resolution of approximately 0.01 kPa so a change in 1 kPa could also be sufficient. At lower CO 2 and/or light levels, more tissue may be required or alternatively, longer incubation times are needed. However, small tissue samples are preferred to prevent self shading and to promote good mixing in the vials so that tissues are well exposed to light and chemicals during incubation.

Example 1: Preparation of artificial floodwater with CA of 2.0 mmol L −1 and 200 μmol free CO 2 L −1 . Prepare a solution of DI water containing Ca 2+ , Mg 2+ , SO 4 2− , and Cl − at the concentrations described above. Divide the solution into two containers and bubble one half of the solution with air and the other half with N 2 for 20 min and then mix the two solutions. Add the required amount of DIC (Table 3 , highlighted in yellow for this example) which is 2.2 mmol L −1 . Add the DIC in the form of KHCO 3 , NaHCO 3 or a mixture, and acidify the solution to pH 7.35 using HCl. This results in a solution with a CA of 2 mmol L −1 (in mmol L −1 : 1.995 HCO 3 - + 0.002 CO 3 2 - ) and 200 μmol L −1 CO 2 (Table 3 ).

Incubator with light and temperature control

The incubator provides constant temperature and mixing throughout the incubation. It consists of a vertically rotating wheel where glass bottles or vials can be clipped on facing the light source. The wheel rotates at about 10 rpm in a tank with temperature controlled water and a transparent glass or Perspex wall for illumination at various irradiances (Figure 1 C).

The rotating wheel incubator was originally invented for photosynthesis measurements in phytoplankton ( Steemann Nielsen, 1952 ) and the typical light source in commercially available models consists of a rack of fluorescent tubes. However, it is hard to achieve PAR levels much above 500 μmol photons m −2 s −1 with fluorescent light so high pressure metal halide lamps (mercury or sodium) or light emitting plasma lamps are required to provide the levels of light needed to light saturate net photosynthesis by leaves of many terrestrial species and some macroalgae with thick thalli.

Photosynthesis versus light curves (i.e. light response curves) are obtained by: (i) regulating light intensities by varying the distance of the light source to the incubator, (ii) placing neutral shading filters in front of the light source, (iii) placing a box with neutral shading filters of variable transmission in front of individual vials, or (iv) by wrapping the vials in layers of neutral shading mesh, or by a combination of these various approaches.

O 2 measurements

The O 2 produced or consumed during incubation can be measured directly in the glass vials using O 2 electrodes or optodes. In the absence of good electrodes or optodes, the Winkler titration can also be applied; see Strickland and Parsons (1972) for details.

Contemporary methods for O 2 measurements in water involve either Clark type electrodes or optodes. A Clark type O 2 electrode measures pO 2 as molecular O 2 transverses a membrane before the electrochemical reaction on the cathode results in a current which is linearly proportional to the pO 2 of the medium. Since the electrode consumes O 2 , a conventional large O 2 electrode is quite stirring sensitive and it is thus much more convenient to use an O 2 microelectrode which consumes little O 2 to address the stirring issue during measurements; O 2 microelectrodes can have a stirring sensitivity of less than 1% ( Revsbech and Jørgensen, 1986 ; Revsbech, 1987 ). Oxygen microelectrodes typically have a temperature coefficient of approximately 1–3%°C −1 ( Revsbech, 1987 ; Gundersen et al., 1998 ) so temperature control during measurements is essential. The temperature effect on electrodes (and optodes, see below) is primarily caused by changes in diffusion and electrochemistry. In addition, temperature also influences solubility of gases, and metabolic rate of the tissues.

The measuring principle of optodes is quite different from that of a Clark type electrode. In the optode, light excites a fluorophore coated onto the tip of fiber optics and the excited light is subsequently transmitted back and measured by a spectroradiometer ( Klimant et al., 1997 ). Alternatively, the fluorophore can be coated onto tiny plastic patches which (microdots) can be mounted directly in the medium where O 2 is to be measured; the microdot with the fluorophore is then illuminated from the outside through the transparent wall of the container. Molecular O 2 quenches the florescence so that the transmitted signal can be calibrated toward O 2 in the medium; the relationships between quenching and pO 2 is non-linear. Optodes do not consume O 2 and are thus completely insensitive to stirring. However, O 2 optodes can have higher temperature coefficients than Clark type microelectrodes and require even better temperature control during measurements ( Kragh et al., 2008 ). On the other hand, optodes can be built into the individual glass vials (microdots glued onto the glass wall inside the vial) and the O 2 concentration can be measured in a non-destructive manner ( Kragh et al., 2008 ). The great advance of this approach is that vials can remain sealed and be returned to the rotating wheel if a preliminary reading shows that longer incubation is required in order to obtain the necessary accuracy, or O 2 evolution can be followed over time to ensure quasi steady state measurements or to elucidate possible temporal patterns.

Supporting measurements and calculations

After measuring O 2 of each vial, the tissue must be processed according to standard procedures to establish the area, the fresh mass or dry mass, the chlorophyll concentration, or all of the above. The underwater net photosynthesis is calculated as the net O 2 evolution rate per unit tissue per unit time. In practice, the change in O 2 content in each vial (change in O 2 concentration multiplied by the volume of the vial; individual volumes of vials (i.e. minus the volume of the glass beads, etc.) must be established) divided by the incubation time and divided by the amount of tissue (i.e., mass, area or any other of the above mentioned parameters used to scale photosynthesis per sample unit). An example of a CO 2 response curve established with the technique described here in Section “The Rotating Wheel Incubator” is shown in Figure 6 .

Figure 6. Underwater net photosynthesis versus CO 2 concentration in the medium for excised leaf segments of Hordeum marinum . Leaf segments (30 mm) were incubated in 35 mL glass vials with various well defined CO 2 concentrations on a rotating wheel with PAR of 350 μmol photons m −2 s −1 at 20°C (see Figure 1 C). O 2 evolution was measured with a Clark type O 2 microelectrode and underwater net photosynthesis was calculated as O 2 evolution per projected area per unit time (See “The Rotating Wheel Incubator”). Data (mean ± SE, n =5) from ( Pedersen et al., 2010 ). Note: leaves of H. marinum are superhydrophobic and so possess a gas film when underwater.

The Closed Chamber with Injection Ports

Principle: a leaf or algal thalli sample is incubated in a closed chamber with internal mixing and possessing injection ports and fitted with an electrode/optode that follows O 2 concentration. The amount of free CO 2 can be manipulated by injection of acid or base while a fitted pH electrode allows calculation of the exact CO 2 level. The approach enables production of a complete light or CO 2 response curve based on the same sample, and underwater net photosynthesis can be calculated based on e.g., leaf area, fresh mass, dry mass, and/or chlorophyll concentration. Incubation in darkness can provide data on dark respiration.

Chamber with light and temperature control

The chamber for measurements of underwater net photosynthesis enables measurements with light, temperature, and CO 2 manipulations in water, with monitoring of O 2 with time. Chambers are commercially available for underwater photosynthesis measurements on macro algae, phytoplankton, or isolated chloroplasts and these are made from glass, acrylic glass, or acetal. These chambers can also be custom built to match specific electrodes, light sources, and fitted with extra ports for temperature and PAR sensors and injection of acid/bases or inhibitors. The chamber must be made from a material the can be sterilized and also have a least one transparent side to enable illumination of the sample. The light source can be diode based (650 nm red diode) or “full spectrum” halogen light to simulate white sunlight. Pay attention to the fact that some lighting devices are unable to produce sufficient light to saturate the photosynthesis of some terrestrial leaves or thick macroalgae thalli. Illumination (even by means of fiber optics) produces heat, so cooling of the chamber by a water jacket is crucial.

A light sensor small enough to measure inside the chamber is also essential. The spherical PAR sensor US-SQS/L (Walz, Effeltrich, Germany) is of a size (Ø = 3.7 mm) that enables permanent installation in most chambers.

Finally, the issue of mixing must be addressed. The simplest solution is to use a glass coated stir bar (avoid Teflon coated stir bars as these can hold O 2 ) which is isolated from the sample with a coarse mesh to prevent contact with the tissue. It may be necessary to fix the tissue in the swirling current; if the tissue rotates with the water current in the chamber, the DBL will be larger than if the tissue is fixed. The thicker DBL increases the apparent resistance to CO 2 uptake or O 2 escape.

O 2 and pH measurements

O 2 measurements in the closed chamber are similar to O 2 measurements in the vials described in Section “O 2 Measurements.” An O 2 sensor (Clark type electrode or optode) is fixed in the chamber in one of the ports, or if an optode is used, a patch with fluorophore can be glued onto the interior wall. A pH electrode is fitted in a second port and the signals from both sensors are logged onto a computer with data acquisition software. Calibration of both O 2 and pH sensors should be performed in the chamber to avoid stirring related artifacts to the calibrations. Remember to pay extra attention to temperature if using O 2 optodes. It may take a while for the temperature of the solution inside the chamber to equilibrate with that of the cooling jacket, and working in a constant temperature room or keeping solutions in a thermostated water bath will significantly reduce the time it takes before a temperature steady state is obtained; always measure temperature directly in the chambers. Temperature influences electrode or optode performance, solubility of gases, and metabolic rate of the tissues (see Section “O 2 Measurements”). After insertion of tissue and filling of the chamber with medium (see below), pH can be manipulated by injection of small amounts of acid or base through one of the injection ports. Fit a 27G needle in one of the injection ports and let it function as “over pressure valve” to prevent pressurization during injection of acid or base (or inhibitors); the needle may be left in the stopper during the experiment as diffusion of gases in water is too slow to result in experimental artifacts.

As described in Section “The Rotating Wheel Incubator” for incubations of tissues in closed vials on the wheel, substantial photorespiration can occur if O 2 is allowed to build up in the medium. Therefore, the susceptibility to photorespiration should initially be established for each tissue type. The linearity of O 2 production with increasing external pO 2 is easily tested the following way: a medium with total DIC of 5.0 mmol L −1 is prepared from KHCO 3 in a 5.0 mmol L −1 TES buffer adjusted to pH 8.00 and with a pO 2 of 10 kPa. The tissue is then allowed to photosynthesize up to a pO 2 of 30 kPa. Here, approximately 500 μmol O 2 has been produced from 500 μmol CO 2 and because of the TES buffer the pH has remained at 8.0. Although the DIC pool has declined to 4.5 mmol L −1 , free CO 2 has changed by only 10% from 110 to 100 μmol L −1 . If the O 2 evolution occurs linearly in this range, it means that the approximately threefold lower CO 2 :O 2 in the medium, with likely even greater changes in internal CO 2 :O 2 , has not increased photorespiration. If the curve exhibits a saturation tendency (i.e. declining rate of net O 2 production with increasing pO 2 ), photorespiration has probably increased with increasing pO 2 in the chamber.

Medium and tissue may be prepared as described in Section “Medium and Tissue.” However, as a CO 2 response curve in the closed photosynthesis chamber often involves conversion of HCO 3 - into free CO 2 (dissolved), e.g., by manipulation of pH, enough HCO 3 - must initially be present in the medium to produce the required levels of free CO 2 . Following injection of small amounts of acid or base to manipulate free CO 2 , the rate of net photosynthesis changes accordingly so that a new rate of net O 2 production (slope of dissolved O 2 with time) is established at each dissolved CO 2 . However, pH may also change slightly in the time interval because CO 2 is extracted from the system as it is fixed via photosynthesis (Eqs 1 and 4). Hence, for every rate of underwater net photosynthesis determined in a time interval, the mean CO 2 concentration must be calculated in order to present the CO 2 response curve of the tissue.

Example 2: average free CO 2 concentration in the pH range from 7.25 to 7.30 in a medium with total DIC of 2.0 mmol L −1 . According to Gutz (2012) , CA of such a solution at pH 7.25 would be 1.77 mmol L −1 having 223 μmol CO 2 L −1 ; at pH 7.30 CA would be 1.80 mmol L −1 and have 203 μmol CO 2 L −1 . Consequently, the average free CO 2 concentration in the pH range was 213 μmol CO 2 L −1 .

After each experiment, the incubated tissue must be characterized to enable calculation of underwater net photosynthesis rates; the supporting measurements are as those described in section “Supporting Measurements and Calculations.”

pH Drift Approach to Establish CO 2 Compensation Points

Principle: Leaf or algal thalli samples are incubated in glass vials for 16–18 h where after pH and CA or DIC are measured. CO 2 compensation points and carbon extraction capacity of tissues can be calculated. The method is also used as a diagnostic test for bicarbonate ( HCO 3 - ) use in underwater photosynthesis.

These long term incubations are used to test how far net photosynthesis of a given plant sample at saturating light can extract DIC, i.e. to deplete CO 2 and HCO 3 - and drive up pH. Because the objective is to determine the ultimate DIC extraction capacity and maximum upper pH in a standardized way, all incubation vials are prepared to have an equal standard DIC concentration (usually 1–2 mmol L −1 for alkaline waters and 0.1–0.3 mmol L −1 for softwaters) and a pH, CO 2 , and O 2 concentration corresponding to air equilibrium ( Sand-Jensen et al., 1992 , 2009 ). Artificial media and natural waters can be applied. However, to minimize O 2 build up and the risk of photorespiration during extended incubation the initial O 2 can be reduced to 20–50% of air equilibrium. To ensure the maximum possible DIC depletion, the amount of plant material is typically three times larger than in the incubations described in Sections “The rotating Wheel Incubator” and “The Closed Chamber with Injection Ports” though it must still be able to move freely in the vials to ensure adequate mixing.

The initial and final DIC and pH must be determined in order to calculate the DIC extraction capacity during incubation and the CO 2 compensation point after incubation. Provided no internal pools of DIC and protons interfere with conditions in the water/medium and no precipitation or dissolution of carbonates takes place, DIC can be determined in the medium from CA, pH, temperature, and ionic strength; CA in turn can be determined by acidimetric titration ( Stumm and Morgan, 1996 ). The risk of carbonate precipitation is small in artificial media of KHCO 3 and much larger in natural waters and artificial media where Ca(HCO 3 ) 2 dominates, the reason being that K 2 CO 3 is highly soluble and CaCO 3 is poorly soluble. Calcium carbonate precipitation is likely to take place in pH drift experiments where final pH exceeds 10. Therefore, it is always recommended to directly measure DIC. This can be done by injecting of small water samples into concentrated acid in a bubble chamber purged with N 2 gas carrying the released CO 2 into an IRGA ( Vermaat and Sand-Jensen, 1987 ). Water samples may need to be filtered (with no atmospheric contact) if minute CaCO 3 crystals have been formed in the external water of high pH. It is generally recommendable to determine (or check) CO 2 compensation points by depletion experiments in media of low initial DIC (<50 μmol L −1 ) and low pH (<6.5) where the interference by HCO 3 - is low and CaCO 3 is not formed.

The pH drift technique has also been used to determine DIC consumption at intervals during the ongoing drift of pH upwards ( Maberly and Spence, 1983 ; Spence and Maberly, 1985 ). DIC, pH, the proportion of carbon species and O 2 change during the time of incubation. Because all parameters may influence photosynthesis, and exchange with internal DIC and proton pools in the incubated tissue may interfere with calculations, we cannot recommend the procedure for determining rates of net photosynthesis considering the much more accurate and straightforward methods being available today (as described in this review).

Community Photosynthesis in Large Chambers

Principle: Community photosynthesis is measured in large closed chambers with linear dimensions of 0.5–0.6 m, or larger, to minimize edge effects and make certain that natural changes of plant density, tissue capacity and irradiance through the canopy are maintained. Photosynthetic rates are measured by O 2 and DIC, as for phytoelements in small chambers (See The Closed Chamber with Injection Ports), but photosynthetic parameters and their dependence on DIC and temperature are markedly different for communities than phytoelements.

Submerged aquatic plants grow in communities of variable density where the spatial structure and self shading are prominent features ( Sand-Jensen, 1989 ). Light limitation is substantial and the efficiency of photosynthesis at low light is therefore important ( Binzer and Sand-Jensen, 2002a , b ). The photosynthetic chamber needs to be large enough to include tall communities ( Binzer et al., 2006 ; Middelboe et al., 2006 ). It is made of glass or transparent acrylic glass and viewed from above, the shape of photosynthetic chambers can be cylindrical, rectangular, or quadratic. The cylindrical shape can be advantageous because the surface area of side walls relative to chamber volume is smaller than in rectangular or quadratic chambers, and these two latter types may also have “dead corners” with stagnant waters. The light sources are high pressure metal halide lamps (mercury or sodium) or light emitting plasma lamps because only those provide a sufficiently high irradiance (>1000 μmol photon m −2 s −1 ). The light sources must be placed more than 0.5 m above the photosynthetic chamber and the light path both above the chamber and around the chamber walls are surrounded by totally reflecting material to reduce the influence of distance with depth in the chamber both when plants are absent or present. Irradiance is measured with depth in the water and through canopies of different densities using a small spherical PAR sensor. To ensure statistically reliable determinations of vertical attenuation a series (e.g., 10) of measurements are performed at different positions ( Middelboe et al., 2006 ). Temperature, O 2 , DIC, and pH are set and measured as described in Section “The Closed Chamber with Injection Ports,” while mixing is provided by large submersible pumps ensuring current velocities above 2 cm s −1 . Temperature control is attained by direct cooling and warming of the water in the incubation chamber or by placing it in a larger temperature controlled holding tank. In the latter case some temperature variations (1–3°C) is difficult to avoid between light and darkness.

Algal communities for measurements can be collected attached to stones or established over a period of one or several years on artificial tiles of desired size set out in the field and later brought to the laboratory for measurements in the photosynthetic chamber ( Binzer et al., 2006 ; Middelboe et al., 2006 ). Rooted submerged plants can be harvested from natural stands with the 3D structure kept intact when roots and rhizomes are interwoven. In other cases, individual plants are placed in a homogeneous pattern on the chamber bottom in small plastic bags surrounding the root system. Alternatively the individuals are tied to a frame on the chamber bottom. Plant density is determined as fresh mass, dry mass, or plant surface area normalized to bottom area. Leaf area indices (LAI) ranging from 1 to 12 are useful for comparisons among species. Vertical distribution of plant biomass and surface can be determined by cutting the plants sequentially in well defined strata starting at the top of the canopy.

The setup is suited to evaluate the influence on community photosynthesis by variable irradiance, temperature, DIC (including variable CO 2 and HCO 3 - ), canopy density, and spatial structure ( Sand-Jensen et al., 2007 ).

Community photosynthesis can also be determined over longer periods of time by employing the large chambers in an open mode. This allows for exchange of O 2 and CO 2 with the atmosphere to prevent that the chambers undergo too extensive accumulation and depletion in the water during several days of alternating light or dark periods. For calculation of photosynthesis and respiration, exchange rates between air and water must be determined. The flux (F exch , mol m −2 s −1 ) between water and air for O 2 is given by the equation:

where K is the exchange coefficient (piston velocity, ms −1 ), C act is the actual and C equ is the equilibrium concentration of O 2 (mol m −3 ) in water at the actual temperature ( Staehr et al., 2012b ). Piston velocity is controlled by surface turbulence and can, therefore, be considered a constant for a given mixing regime determined by the strength and location of the pumps and the dampening influence of the plant community. Thus, K must be directly measured for a given plant density and mixing regime. This is best done in the dark, where only dark respiration (mol m −2 s −1 ) takes place, by modifying C act to for example 10 or 30 kPa and measuring the total O 2 flux (F) over time as a result of respiration and exchange with the atmosphere above from time changes in O 2 concentrations in the water:

From 30 kPa the actual pO 2 will first rapidly decline as a combined result of respiration and loss to the atmosphere and gradually decline less rapidly as pO 2 approaches equilibrium with the atmosphere and respiration alone drives pO 2 further downwards. Calculations of pO 2 changes over time in relation to differences in the pO 2 gradient between water and air produces a straight line (Eq. 8) permitting calculation of R and K assuming that they remain constant for a given mixing intensity and plant density.

Community measurements operated in an open mode have the main advantage for future application that fluxes of O 2 , DIC, Ca 2+ , H + , and nutrient ions (NH 4 + , NO 3 −, and PO 4 3− ) can be determined during repeated diel light dark cycles for several weeks while the submerged plants may also grow. Combined field measurements have been operated in open chambers and mesocosms under a strict mixing regime under natural temperature and light conditions both for phytoplankton (e.g., Markager and Sand-Jensen, 1989 ), submerged aquatic plants (e.g., Liboriussen et al., 2005 ), and flooded terrestrial plants (e.g., Setter et al., 1988 ).

The Open Natural System

Principle: Natural ecosystems dominated by submerged aquatic plants have free undisturbed gas exchange with the atmosphere and input/output of water. Determination of ecosystem metabolism by open water measurements requires accurate calculations of atmospheric exchange of O 2 and CO 2 . The main advantages of the ecosystem approach is that environmental conditions and processes are natural and temporal patterns can be followed over months or years, while allowing plant density and acclimation to gradients in light, DIC, and other environmental variables to develop.

Photosynthesis of submerged aquatic plants derived from analysis of ecosystems can only be determined when rooted plants or macroalgae are the main phototrophs responsible of more than 90% of ecosystem photosynthesis. Only then can the patterns obtained be referred to the metabolism of macrophytes accepting that a minor error (<10%) due to photosynthesis of microalgae may be present. The dominance of submerged aquatic plants can be realized in shallow plant rich ponds, lakes, streams, and coastal lagoons. Open water measurements are used to follow changes in O 2, DIC, pH, temperature, and irradiance, and enable calculation of ecosystem net production, plant gross production, and community respiration assuming fully mixed conditions ( Odum, 1956 ; Staehr et al., 2012a ). Meteorological observations of wind direction, wind velocity, atmospheric pressure, etc., in standing waters and current velocity, water depth, slope, and bed roughness in flowing waters, can be used to estimate physical exchange coefficients of gases (i.e. piston velocities) and thus calculate fluxes between water and atmosphere using empirical models ( Sand-Jensen and Staehr, 2011 ). Flow chambers, floating chambers, inert gases, and coverage of water surfaces by impermeable floating plastic can be used for direct determination of exchange coefficients which are critical in all determinations of ecosystem metabolism ( Staehr et al., 2012a , b ). Rooted plants with gas filled lacunae formation and release of gas bubbles can introduce error. Oxygen storage may delay establishment of steady state exchange of O 2 following dark light switches by some 10–20 min for most rooted plants ( Westlake, 1978 ) and loss of bubbles is negligible in swift flowing waters, while bubble release may account for 10% of net O 2 release in slow flowing waters (Kragh et al., unpublished data).

The strength of these measurements is that they provide natural rates under fully realistic and undisturbed environmental conditions. They can reveal the coupling between O 2 and carbon metabolism, the natural precipitation and dissolution of carbonates and the direct involvement of accumulation and release of acids in the photosynthetic process. Measurements have shown fast exchange rates of protons between macrophytes and water following diurnal light dark switches partly uncoupled from exchanges of DIC during photosynthesis and respiration; a phenomenon that is not unraveled in short term laboratory measurements with detached phytoelements (Kragh et al., unpublished data). Ecosystem measurements can also reveal how early summer growth in biomass and late summer senescence influence plant metabolism and how ongoing desiccation of ponds may suddenly stop photosynthesis and accelerate decomposition, while refilling may restart photosynthesis and growth ( Christensen et al., 2013 ). Modeling approaches, as successfully used for canopy level understanding of terrestrial systems systems ( Ainsworth and Long, 2005 ), should also be applied more widely in studies of aquatic systems (e.g., Binzer and Sand-Jensen, 2002a , b ). All techniques for measuring and calculating ecosystem process are basically available ( Staehr et al., 2012a ) and awaits broad scale application.

Studies of photosynthesis by aquatic and submerged wetland plants are few compared with research on photosynthesis in air, but underwater systems are attracting more attention. Light and CO 2 availability under water are often low to submerged plants. Low CO 2 together with impeded escape of O 2 can result in high photorespiration as a component determining net photosynthesis. Focus studies of contrasting species and systems are required to develop our understanding of “models” since the environment under water is more complex than in air and there is a diversity of photosynthetic mechanisms (i.e. C 3 , C 4 , CAM, and bicarbonate use) in aquatic species.

The physical and chemical environments of overland floods are only poorly known and few data exist on light extinction and CO 2 and O 2 concentrations in floodwaters. Such data are crucial to design relevant laboratory experiments on submergence tolerance of terrestrial plants and to establish, for example, carbon budgets during submergence on leaf lamina as well as for whole plants. Also, studies on leaf acclimation of terrestrial plants to facilitate gas exchange and light utilization under water are also only in their infancy; these acclimations influence underwater photosynthesis as well as internal aeration of plant tissues during submergence.

Finally, a challenge also exists to assess the influence of light, inorganic carbon, and temperature on natural aquatic communities of variable density instead of only studying detached leaves in the scenarios of rising CO 2 and temperature. Use of mathematical modeling, both at the leaf and community levels, will provide valuable additional understanding of underwater photosynthesis. Improved knowledge of plant and environmental factors determining rate of underwater net photosynthesis at various scales (leaf-to-community) is essential for understanding aquatic plant ecophysiology, submergence tolerance of terrestrial plants, and productivity of the many aquatic and flood-prone ecosystems worldwide.

Conflict of Interest Statement

The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Acknowledgments

The present work was carried out with support of the Danish Council for Independent Research grant no. 09-072482 and under the Lake Restoration Centre, a Villum Kann Rasmussen Centre of Excellence. Timothy D. Colmer acknowledges support from the Australian Research Council (Discovery Grant).

Adams, M. S., Guilizzoni, P., and Adams, S. (1978). Relationship of dissolved inorganic carbon to macrophyte photosynthesis in some Italian lakes. Limnol. Oceanogr. 23, 912–919.

CrossRef Full Text

Ainsworth, E. A., and Long, S. P. (2005). What have we learned from 15 years of free-air CO 2 enrichment (FACE)? A meta-analytic review of the responses of photosynthesis, canopy properties and plant production to rising CO 2 . New Phytol. 165, 351–372.

Pubmed Abstract | Pubmed Full Text | CrossRef Full Text

Arens, K. (1933). Physiologisch polarisierter Massenaustausch und Photosynthese bei submersen Wasserpflanzen. I. Planta 20, 621–658.

Armstrong, W. (1979). Aeration in higher plants. Adv. Bot. Res. 7, 225–332.

Bailey-Serres, J., Fukao, T., Ismail, A. M., Heuer, S., and Mackill, D. J. (2010). Submergence tolerant rice: SUB1’s journey from landrace to modern cultivar. Rice 3, 138–147.

Bailey-Serres, J., and Voesenek, L. A. C. J. (2008). Flooding stress: acclimations and genetic diversity. Annu. Rev. Plant Biol. 59, 313–339.

Binzer, T., and Sand-Jensen, K. (2002a). Importance of structure and density of macroalgae communities ( Fucus serratus ) for photosynthetic production and light utilisation. Mar. Ecol. Prog. Ser. 235, 53–62.

Binzer, T., and Sand-Jensen, K. (2002b). Production in aquatic macrophyte communities: a theoretical and empirical study of the influence of spatial light distribution. Limnol. Oceanogr. 47, 1742–1750.

Binzer, T., Sand-Jensen, K., and Middelboe, A. L. (2006). Community photosynthesis of aquatic macrophytes. Limnol. Oceanogr. 51, 2722–2733.

Black, M. A., Maberly, S. C., and Spence, D. H. N. (1981). Resistances to carbon dioxide fixation in four submerged freshwater macrophytes. New Phytol. 89, 557–568.

Borum, J., Pedersen, O., Greve, T. M., Frankovich, T. A., Zieman, J. C., Fourqurean, J. W., et al. (2005). The potential role of plant oxygen and sulphide dynamics in die-off events of the tropical seagrass, Thalassia testudinum . J. Ecol. 93, 148–158.

Brewer, C. A., and Smith, W. K. (1997). Patterns of leaf surface wetness for montane and subalpine plants. Plant Cell Environ. 20, 1–11.

Christensen, J. P. A., Sand-Jensen, K., and Staehr, P. A. (2013). Fluctuating water levels control water chemistry and metabolism of a charophyte pond. Freshw. Biol . doi:10.1111./fwb.12132

Cole, J. J., Prairie, Y. T., Caraco, N. F., McDowell, W. H., Tranvik, L. J., Striegl, R. G., et al. (2007). Plumbing the global carbon cycle: integrating inland waters into the terrestrial carbon budget. Ecosystems 10, 172–185.

Colmer, T. D., and Pedersen, O. (2008). Underwater photosynthesis and respiration in leaves of submerged wetland plants: gas films improve CO 2 and O 2 exchange. New Phytol. 177, 918–926.

Colmer, T. D., and Voesenek, L. A. C. J. (2009). Flooding tolerance: suites of plant traits in variable environments. Funct. Plant Biol. 36, 665–681.

Colmer, T. D., Winkel, A., and Pedersen, O. (2011). A perspective on underwater photosynthesis in submerged terrestrial wetland plants. AoB Plants 2011, lr030.

Dickson, A. G. (1981). An exact definition of total alkalinity and a procedure for the estimation of alkalinity and total inorganic carbon from titration data. Deep Sea Res. A 28, 609–623.

Duarte, C. M. (1991). Seagrass depth limits. Aquat. Bot. 40, 363–377.

Frost-Christensen, H., and Sand-Jensen, K. (1992). The quantum efficiency of photosynthesis in macroalgae and sumberged angiosperms. Oecologia 91, 377–384.

Gundersen, J. K., Ramsing, N. B., and Glud, R. N. (1998). Predicting the signal of O 2 microsensors from physical dimensions, temperature, salinity, and O 2 concentration. Limnol. Oceanogr. 43, 1932–1937.

Gutz, I. G. R. (2012). CurTiPot – pH and Acid-base Titration Curves: Analysis and Simulation Software, Version 3.6.1 [Online]. Available at: http://www2.iq.usp.br/docente/gutz/Curtipot.html [accessed 16 Dec 2012 2013].

Hartman, R. T., and Brown, D. L. (1967). Changes in internal atmosphere of submersed vascular hydrophytes in relation to photosynthesis. Ecology 48, 252–258.

Helder, R. J. (1985). Diffusion of inorganic carbon across an unstirred layer: a simplified quantitative approach. Plant Cell Environ. 8, 399–408.

Holmer, M., Pedersen, O., Krause-Jensen, D., Olesen, B., Petersen, M. H., Schopmeyer, S., et al. (2009). Sulfide intrusion in the tropical seagrasses Thalassia testudinum and Syringodium filiforme . Estuar. Coast. Shelf Sci. 85, 319–326.

Johnson, F. H., Eyring, H., and Stover, B. J. (1974). The Theory of Rate Processes in Biology and Medicine . New York: Wiley.

Kelly, M. G., Thyssen, N., and Moeslund, B. (1983). Light and the annual variation of oxygen- and carbon-based measurements of productivity in a macrophyte-dominated river. Limnol. Oceanogr. 28, 503–515.

Kemp, W. M., Marlon, R. L., and Jones, T. W. (1986). Comparison of methods for measuring production by the submersed macrophyte, Potamogeton perfoliatus L. Limnol. Oceanogr. 31, 1322–1334.

Kirk, J. T. O. (1994). Light and Photosynthesis in Aquatic Ecosystems . New York: Cambridge Univ Press.

Klimant, I., Kühl, M., Glud, R. N., and Holst, G. (1997). Optical measurement of oxygen and temperature in microscale: strategies and biological applications. Sens. Actuators B Chem. 38, 29–37.

Kragh, T., Søndergaard, M., and Tranvik, L. (2008). Exposure to sunlight and phosphorus-limitation on bacterial degradation of coloured dissolved organicmatter (CDOM) in freshwater. FEMS Microbiol. Ecol. 64, 230–239.

Lambers, H., Chapin, F. S. III, and Pons, T. L. (2008). Plant Physiological Ecology . Heidelberg: Springer Verlag.

Larsson, C., and Axelsson, L. (1999). Bicarbonate uptake and utilization in marine macroalgae. Eur. J. Phycol. 34, 79–86.

Liboriussen, L., Landkildehus, F., Meerhoff, M., Bramm, M. E., Søndergaard, M., Christensen, C., et al. (2005). Global warming: design of a flow-through shallow lake mesocosm climate experiment. Limnol. Oceanogr. Methods 3, 1–9.

Long, S. P. (1991). Modification of the response of photosynthetic productivity to rising temperature by atmospheric CO 2 concentrations: has its importance been underestimated? Plant Cell Environ. 14, 729–739.

Lucas, W. J., and Smith, F. A. (1973). The formation of alkaline and acid regions at the surface of Chara corallina cells. J. Exp. Bot. 24, 1–14.

Maberly, S. C. (1990). Exgenous sources of inorganic carbon for photosynthesis by marine macroalgae. J. Phycol. 26, 439–449.

Maberly, S. C. (1996). Diel, episodic and seasonal changes in pH and concentrations of inorganic carbon in a productive lake. Freshw. Biol. 35, 579–598.

Maberly, S. C., and Madsen, T. V. (2002). Freshwater angiosperm carbon concentrating mechanisms: processes and patterns. Funct. Plant Biol. 29, 393–405.

Maberly, S. C., and Spence, D. H. N. (1983). Photosynthetic inorganic carbon use by freshwater plants. J. Ecol. 71, 705–724.

Maberly, S. C., and Spence, D. H. N. (1989). Photosynthesis and photorespiration in freshwater organisms: amphibious plants. Aquat. Bot. 34, 267–286.

Mackereth, F. J. H., Heron, J., and Talling, J. F. (1978). Water Analysis: Some Revised Methods for Limnologists . Cumbria: Freshwater Biological Association.

Mackill, D. J., Ismail, A. M., Singh, U. S., Labios, A. V., and Paris, T. R. (2012). Development and Rapid Adoption of Submergence-Tolerant (Sub1) Rice Varieties . San Diego: Academic Press, 299–352.

Madsen, T. V., and Sand-Jensen, K. (1991). Photosynthetic carbon assimilation in aquatic macrophytes. Aquat. Bot. 41, 5–40.

Madsen, T. V., Sand-Jensen, K., and Beer, S. (1993). Comparison of photosynthetic performance and carboxylation capacity in a range of aquatic macrophytes of different growth forms. Aquat. Bot. 44, 373–384.

Markager, S., and Sand-Jensen, K. (1989). Patterns of night-time respiration in a dense phytoplankton community under a natural light regime. J. Ecol. 77, 49–61.

McConnaughey, T. A. (1991). Calcification in Chara corallina : CO 2 hydroxylation generates protons for bicarbonate assimilation. Limnol. Oceanogr. 36, 619–628.

McConnaughey, T. A., La Baugh, J. W., Rosenberry, D. O., Striegl, R. G., Reddy, M. M., Schuster, P. F., et al. (1994). Carbon budget for a groundwater-fed lake: calcification supports summer photosynthesis. Limnol. Oceanogr. 39, 1319–1332.

Middelboe, A. L., and Markager, S. (1997). Depth limits and minimum light requirements of freshwater macrophytes. Freshw. Biol. 37, 553–568.

Middelboe, A. L., Sand-Jensen, K., and Binzer, T. (2006). Highly predictable photosynthetic production in natural macroalgal communities from incoming and absorbed light. Oecologia 150, 464–476.

Mommer, L., Lenssen, J. P. M., Huber, H., Visser, E. J. W., and De Kroon, H. (2006a). Ecophysiological determinants of plant performance under flooding: a comparative study of seven plant families. J. Ecol. 94, 1117–1129.

Mommer, L., Pons, T. L., and Visser, E. J. W. (2006b). Photosynthetic consequences of phenotypic plasticity in response to submergence: Rumex palustris as a case study. J. Exp. Bot. 57, 283–290.

Mommer, L., and Visser, E. J. W. (2005). Underwater photosynthesis in flooded terrestrial plants: a matter of leaf plasticity. Ann. Bot. 96, 581–589.

Mommer, L., Wolters-Arts, M., Andersen, C., Visser, E. J. W., and Pedersen, O. (2007). Submergence-induced leaf acclimation in terrestrial species varying in flooding tolerance. New Phytol. 176, 337–345.

Moulin, P., Andría, J. R., Axelsson, L., and Mercado, J. M. (2011). Different mechanisms of inorganic carbon acquisition in red macroalgae (Rhodophyta) revealed by the use of TRIS buffer. Aquat. Bot. 95, 31–38.

Neinhuis, C., and Barthlott, W. (1997). Characterization and distribution of water-repelent, self-cleaning plant surfaces. Ann. Bot. 79, 667–677.

Nielsen, S. L. (1993). A comparison of aerial and submerged photosynthesis in some Danish amphibious plants. Aquat. Bot. 45, 27–40.

Nielsen, S. L., and Sand-Jensen, K. (1989). Regulation of photosynthetic rates of submerged rooted macrophytes. Oecologia 81, 364–368.

Odum, H. T. (1956). Primary production in flowing waters. Limnol. Oceanogr. 1, 102–117.

Olesen, B., and Madsen, T. V. (2000). Growth and physiological acclimation to temperature and inorganic carbon availability by two submerged aquatic macrophyte species, Callitriche cophocarpa and Elodea canadensis . Funct. Ecol. 14, 252–260.

Opdyke, B. N., and Walker, J. C. G. (1992). Return of the coral reef hypothesis: basin to shelf partitioning of CaCO 3 and its effect on atmospheric CO 2 . Geology 20, 733–736.

Parolin, P. (2009). Submerged in darkness: adaptations to prolonged submergence by woody species of the Amazonian floodplains. Ann. Bot. 103, 359–376.

Parry, M. L., Canziani, O. F., Palutikof, J. P., van der Linden, P. J., and Hanson, C. E. (eds) (2007). Climate change 2007: impacts, adaptation and vulnerability. Contribution of Working Group II to the Fourth Assessment Report of the Intergovernmental Panel on Climate Change (Cambridge: Cambridge University Press), 982.

Pedersen, O., and Colmer, T. D. (2012). Physical gills prevent drowning of many wetland insects, spiders and plants. J. Exp. Biol. 215, 705–709.

Pedersen, O., Malik, A. I., and Colmer, T. D. (2010). Submergence tolerance in Hordeum marinum : dissolved CO 2 determines underwater photosynthesis and growth. Funct. Plant Biol. 37, 524–531.

Pedersen, O., Pulido, C., Rich, S. M., and Colmer, T. D. (2011). In situ O 2 dynamics in submerged Isoetes australis : varied leaf gas permeability influences underwater photosynthesis and internal O 2 . J. Exp. Bot. 62, 4691–4700.

Pedersen, O., Rich, S. M., and Colmer, T. D. (2009). Surviving floods: leaf gas films improve O 2 and CO 2 exchange, root aeration, and growth of completely submerged rice. Plant J. 58, 147–156.

Pedersen, O., Vos, H., and Colmer, T. D. (2006). Oxygen dynamics during submergence in the halophytic stem succulent Halosarcia pergranulata . Plant Cell Environ. 29, 1388–1399.

Pilon, J., and Santamaría, L. (2001). Seasonal acclimation in the photosynthetic and respiratory temperature responses of three submerged freshwater macrophyte species. New Phytol. 151, 659–670.

Price, G. D., and Badger, M. R. (1985). Inhibition by proton buffers of photosynthetic utilization of bicarbonate in Chara corallina . Aust. J. Plant Physiol. 12, 257–267.

Prins, H. B. A., Snel, J. F. H., Helder, R. J., and Zanstra, P. E. (1980). Photosynthetic HCO 3 - utilization and OH - excretion in aquatic angiosperms: light-induced pH changes at the leaf surface. Plant Physiol. 66, 818–822.

Raskin, I., and Kende, H. (1983). How does deep water rice solve its aeration problem? Plant Physiol. 72, 447–454.

Raven, J. A., and Hurd, C. L. (2012). Ecophysiology of photosynthesis in macroalgae. Photosyn. Res. 113, 105–125.

Revsbech, N. P. (1987). An oxygen microelectrode with a guard cathode. Limnol. Oceanogr. 34, 474–478.

Revsbech, N. P., and Jørgensen, B. B. (1986). “Microelectrodes: their use in microbial ecology,” in Advances in Microbial Ecology , ed. K. C. Marshall (New York: Plenum Press), 293–352.

Rich, S. M., Pedersen, O., Ludwig, M., and Colmer, T. D. (2013). Shoot atmospheric contact is of little importance to aeration of deeper portions of the wetland plant Meionectes brownii ; submerged organs mainly acquire O 2 from the water column or produce it endogenously in underwater photosynthesis. Plant Cell Environ. 36, 213–223.

Richardson, K., Griffiths, H., Reed, M. L., Raven, J. A., and Griffiths, N. M. (1984). Inorganic carbon assimilation in the Isoetids, Isoetes lacustris L. and Lobelia dortmanna L. Oecologia 61, 115–121.

Sand-Jensen, K. (1983). Photosynthetic carbon-sources of stream macrophytes. J. Exp. Bot. 34, 198–210.

Sand-Jensen, K. (1989). Environmental variables and their effect on photosynthesis of aquatic plant-communities. Aquat. Bot. 34, 5–25.

Sand-Jensen, K., Baastrup-Spohr, L., Winkel, A., Moller, C. L., Borum, J., Brodersen, K. P., et al. (2010). Plant distribution patterns and adaptations in a limestone quarry on Oland. Svensk Botanisk Tidskrift 104, 23–31.

Sand-Jensen, K., Binzer, T., and Middelboe, A. L. (2007). Scaling of photosynthetic production of aquatic macrophytes - a review. Oikos 116, 280–294.

Sand-Jensen, K., and Frost-Christensen, H. (1998). Photosynthesis of amphibious and obligately submerged plants in CO 2 -rich lowland streams. Oecologia 117, 31–39.

Sand-Jensen, K., and Frost-Christensen, H. (1999). Plant growth and photosynthesis in the transition zone between land and stream. Aquat. Bot. 63, 23–35.

Sand-Jensen, K., and Gordon, D. M. (1984). Differential ability of marine and fresh-water macrophytes to utilize HCO 3 - and CO 2 . Mar. Biol. 80, 247–253.

Sand-Jensen, K., and Krause-Jensen, D. (1997). Broad-scale comparison of photosynthesis in terrestrial and aquatic plant communities. Oikos 80, 203–208.

Sand-Jensen, K., Pedersen, M. F., and Nielsen, S. L. (1992). Photosynthetic use of inorganic carbon among primary and secondary water plants in streams. Freshw. Biol. 27, 283–293.

Sand-Jensen, K., Pedersen, O., Binzer, T., and Borum, J. (2005). Contrasting oxygen dynamics in the freshwater isoetid Lobelia dortmanna and the marine seagrass Zostera marina . Ann. Bot. 96, 613–623.

Sand-Jensen, K., and Prahl, C. (1982). Oxygen-exchange with the lacunae and across leaves and roots of the submerged vascular macrophyte, Lobelia dortmanna L. New Phytol. 91, 103–120.

Sand-Jensen, K., Raun, A. L., and Borum, J. (2009). Metabolism and resources of spherical colonies of Nostoc zetterstedtii . Limnol. Oceanogr. 54, 1282–1291.

Sand-Jensen, K., and Staehr, P. (2011). CO 2 dynamics along Danish lowland streams: water–air gradients, piston velocities and evasion rates. Biogeochemistry 111, 615–628.

Santamaría, L., and van Vierssen, W. (1997). Photosynthetic temperature responses of fresh- and brackish-water macrophytes: a review. Aquat. Bot. 58, 135–150.

Schwarzenbach, G., and Meier, J. (1958). Formation and investigation of unstable protonation and deprotonation products of complexes in aqueous solution. J. Inorg. Nuclear Chem. 8, 302–312.

Sculthorpe, C. D. (1967). The Biology of Aquatic Vascular Plants . London: Edward Arnold Ltd.

Setter, T. L., Kupkanchanakul, T., Waters, I., and Greenway, H. (1988). Evaluation of factors contributing to diurnal changes in O 2 concentrations in floodwater of deepwater rice fields. New Phytol. 110, 151–162.

Setter, T. L., and Laureles, E. V. (1996). The beneficial effect of reduced elongation growth on submergence tolerance of rice. J. Exp. Bot. 47, 1551–1559.

Setter, T. L., Waters, I., Wallace, I., Bekhasut, P., and Greenway, H. (1989). Submergence of rice. I. Growth and photosynthetic response to CO 2 enrichment of floodwater. Aust. J. Plant Physiol. 16, 251–263.

Silva, J., Sharon, Y., Santos, R., and Beer, S. (2009). Measuring seagrass photosynthesis: methods and applications. Aquatic Biol. 7, 127–141.

Smart, R., and Barko, J. (1985). Laboratory culture of submersed freshwater macrophytes on natural sediments. Aquat. Bot. 21, 251–263.

Smith, W. K., and McClean, T. M. (1989). Adaptive relationship between leaf water repellency, stomatal distribution, and gas exchange. Am. J. Bot. 76, 465–469.

Spence, D. H. N., and Maberly, S. C. (1985). “Occurence and ecological importance of HCO 3 - use among aquatic higher plants,” in Carbon Uptake by Aquatic Photosynthetic Organisms , eds W. J. Lucas and J. L. Berry (Rockville: American Society of Plant Physiologists), 125–143.

Staehr, P., Testa, J., Kemp, W. M., Cole, J., Sand-Jensen, K., and Smith, S. (2012a). The metabolism of aquatic ecosystems: history, applications, and future challenges. Aquatic Sci. 74, 15–29.

Staehr, P. A., Baastrup-Spohr, L., Sand-Jensen, K., and Stedmon, C. (2012b). Lake metabolism scales with lake morphometry and catchment conditions. Aquat Sci 74, 1–15.

Steemann Nielsen, E. (1946). Carbon sources in the photosynthesis of aquatic plants. Nature 158, 594–596.

Steemann Nielsen, E. (1952). The use of radioactive carbon (C 14 ) for measuring organic production in the sea. J. Conseil 18, 117–140.

Stirling, C. M., Davey, P. A., Williams, T. G., and Long, S. P. (1997). Acclimation of photosynthesis to elevated CO 2 and temperature in five British native species of contrasting functional type. Glob. Chang. Biol. 3, 237–246.

Stirling, C. M., Heddell-Cowie, M., Jones, M. L., Ashenden, T. W., and Sparks, T. H. (1998). Effects of elevated CO 2 and temperature on growth and allometry of five native fast-growing annual species. New Phytol. 140, 343–354.

Strickland, J. D. H., and Parsons, T. R. (1972). A Practical Handbook of Seawater Analysis . Ottawa: Bulletin of Fisheries Reseach Board of Canada.

Stumm, W., and Morgan, J. J. (1996). Aquatic Chemistry . New York: John Wiley & Sons.

Suggett, D. J., Prásil, O., and Borowitzka, M. A. (2011). Chlorophyll a Fluorescence in Aquatic Sciences: Methods and Applications . Dordrecht: Springer.

van der Bijl, L., Sand-Jensen, K., and Hjermind, A. L. (1989). Photosynthesis and canopy structure of a submerged plant Potamogeton pectinatus in a Danish lowland stream. J. Ecol. 77, 947–962.

Vashisht, D., Hesselink, A., Pierik, R., Ammerlaan, J. M. H., Bailey-Serres, J., Visser, E. J. W., et al. (2011). Natural variation of submergence tolerance among Arabidopsis thaliana accessions. New Phytol. 190, 299–310.

Vermaat, J. E., and Sand-Jensen, K. (1987). Survival, metabolism and growth of Ulva lactuca under winter conditions - a laboratory study of bottlenecks in the life-cycle. Mar. Biol. 95, 55–61.

Vervuren, P. J. A., Beurskens, S. M. J. H., and Blom, C. W. P. M. (1999). Light acclimation, CO 2 response and long-term capacity of underwater photosynthesis in three terrestrial plant species. Plant Cell Environ. 22, 959–968.

Vervuren, P. J. A., Blom, C. W. P. M., and De Kroon, H. (2003). Extreme flooding events on the Rhine and the survival and distribution of riparian plant species. J. Ecol. 91, 135–146.

Vogel, S. (1994). Life in Moving Fluids: The Physical Biology of Flow . Princeton: Princeton University Press.

Westlake, D. F. (1978). Rapid exchange of oxygen between plant and water. Verh. Int. Verein. Limnol. 20, 2363–2367.

Winkel, A., Colmer, T. D., Ismail, A. M., and Pedersen, O. (2013). Internal aeration of paddy field rice ( Oryza sativa L.) during complete submergence – importance of light and floodwater O 2 . New Phytol. 197, 1193–1203.

Winkel, A., Colmer, T. D., and Pedersen, O. (2011). Leaf gas films of Spartina anglica enhance rhizome and root oxygen during tidal submergence. Plant Cell Environ. 34, 2083–2092.

Keywords: flooding tolerance, light extinction, carbon dioxide, wetland plants, photorespiration

Citation: Pedersen O, Colmer TD and Sand-Jensen K (2013) Underwater photosynthesis of submerged plants – recent advances and methods. Front. Plant Sci. 4 :140. doi: 10.3389/fpls.2013.00140

Received: 17 February 2013; Accepted: 24 April 2013; Published online: 21 May 2013.

Reviewed by:

Copyright: © 2013 Pedersen, Colmer and Sand-Jensen. This is an open-access article distributed under the terms of the Creative Commons Attribution License , which permits use, distribution and reproduction in other forums, provided the original authors and source are credited and subject to any copyright notices concerning any third-party graphics etc.

*Correspondence: Ole Pedersen, The Freshwater Biological Laboratory, Department of Biology, University of Copenhagen, Helsingørsgade 51, 3400 Hillerød, Denmark. e-mail: opedersen@bio.ku.dk

Disclaimer: All claims expressed in this article are solely those of the authors and do not necessarily represent those of their affiliated organizations, or those of the publisher, the editors and the reviewers. Any product that may be evaluated in this article or claim that may be made by its manufacturer is not guaranteed or endorsed by the publisher.

Practical Biology

A collection of experiments that demonstrate biological concepts and processes.

Observing earthworm locomotion

Practical Work for Learning

Published experiments

Investigating factors affecting the rate of photosynthesis, class practical.

In this experiment the rate of photosynthesis is measured by counting the number of bubbles rising from the cut end of a piece of Elodea or Cabomba .

Lesson organisation

The work could be carried out individually or in groups of up to 3 students (counter, timekeeper and scribe).

Apparatus and Chemicals

Students may choose to use:.

Thermometer, –10 °C –110°C

Coloured filters or light bulbs

Push-button counter

Potassium hydrogencarbonate powder or solution (Hazcard 95C describes this as low hazard)

For each group of students:

Student sheets, 1 per student

Beaker, 600 cm 3 , 1

Metre ruler, 1

Elodea ( Note 1 ) or other oxygenating pond plant ( Note 2 )

Electric lamp

Clamp stand with boss and clamp

Health & Safety and Technical notes

Normal laboratory safety procedures should be followed. There is a slight risk of infection from pond water, so take sensible hygiene precautions, cover cuts and wash hands thoroughly after the work is complete.

Read our standard health & safety guidance

1 Elodea can be stored in a fish tank on a windowsill, in the laboratory or prep room. However it is probably a good idea to replace it every so often with a fresh supply from an aquarist centre or a pond. (It’s worth finding out if any colleague has a pond.) On the day of the experiment, cut 10 cm lengths of Elodea , put a paper-clip on one end to weigh them down and place in a boiling tube of water in a boiling tube rack, near a high intensity lamp, such as a halogen lamp or a fluorescent striplight. Check the Elodea to see if it is bubbling. Sometimes cutting 2–3 mm off the end of the Elodea will induce bubbling from the cut end or change the size of the bubbles being produced.

2 Cabomba (available from pet shops or suppliers of aquaria – used as an oxygenator in tropical fish tanks) can be used as an alternative to Elodea , and some people find it produces more bubbles. It does, though tend to break apart very easily, and fish may eat it very quickly.

3 If possible, provide cardboard to allow students to shield their experiment from other lights in the room.

Ethical issues

Look out for small aquatic invertebrates attached to the pond weed used, and remove them to a pond or aquarium.

Effect of N-forms (at 5 mM) and dissolved O 2 levels (1.8 ± 0.2; 2.6 ± 0.2; 3.8 ± 0.2; 5.3 ± 0.2 mg. L − 1 ) of nutrient solution on shoot fresh mass of bell pepper. Different letters show significant different, Duncan’s test ( p  ≤ 0.01)

Effect of N-forms (at 5 mM) and dissolved O 2 levels (1.8 ± 0.2; 2.6 ± 0.2; 3.8 ± 0.2; 5.3 ± 0.2 mg. L − 1 ) of nutrient solution on shoot dry mass of bell pepper. Different letters show significant different, Duncan’s test ( p  ≤ 0.01)

Effect of N-forms (at 5 mM) and dissolved O 2 levels (1.8 ± 0.2; 2.6 ± 0.2; 3.8 ± 0.2; 5.3 ± 0.2 mg. L − 1 ) of nutrient solution on root fresh mass of bell pepper. Different letters show significant different, Duncan’s test ( p  ≤ 0.01)

Effect of N-forms (at 5 mM) and dissolved O 2 levels (1.8 ± 0.2; 2.6 ± 0.2; 3.8 ± 0.2; 5.3 ± 0.2 mg. L − 1 ) of nutrient solution on root dry mass of bell pepper. Different letters show significant different, Duncan’s test ( p  ≤ 0.01)

Many studies have reported a decrease in biomass in plants that are fed with sole NH 4 + , including tomato [ 18 ], cucumber [ 19 ], lettuce [ 20 ], and onion [ 21 ]. This reduction in plant growth can be attributed to various factors, such as a decrease in nutrient uptake, hormonal imbalance, ethylene evolution, futile transmembrane NH 4 + cycling, and carbon skeleton depletion in the root [ 2 ]. The reduction in photosynthesis [ 22 ] and leaf area [ 23 ] is also related to the reduction in plant growth caused by NH 4 + . On the other hand, an increase in biomass has been observed in cucumber [ 24 ], tomato [ 25 ], eggplant [ 8 ], and watermelon [ 26 ] when the O 2 concentration is increased. It has been shown that tomato growth is inhibited at 33% of the ambient oxygen concentration (2.5-3 mg.L -1 ) in a hydroponic system [ 27 ]. The lack of oxygen in the root environment not only directly reduces the activity of the root, but also indirectly reduces the amount of photosynthesis, which in turn reduces the transfer of photosynthetic materials to the root, ultimately leading to a sharp reduction in root growth and destruction [ 28 ]. Biczak et al. [ 29 ] found a significant reduction in the maximum quantum yield of photosystem II under hypoxia conditions, especially in leaves at lower positions on the pepper plant. In the current experiment, consistent outcomes were noted in plants grown with NO 3 − , showing similar results. However, there was no discernible variance in the maximum quantum yield of photosystem II among pepper plants fed with NH 4 + regardless of the varying levels of oxygen concentration. Oxygen deficiency in the nutrient solution can have a detrimental effect on photosynthesis in pepper plants. Without sufficient oxygen available in the root zone, the plant’s ability to uptake nutrients [ 30 ] and water is hindered, leading to decreased nutrient and water availability for the plant. This can disrupt the plant’s metabolic processes, including photosynthesis, as oxygen is necessary for energy production through the electron transport chain. As a result, photosynthetic activity is reduced [ 28 ], leading to decreased growth, lower yields, and overall poor plant health in pepper plants. Ensuring proper aeration and oxygen levels in the nutrient solution is crucial for optimizing photosynthesis and promoting healthy growth in pepper plants.

Waterlogging and reduced oxygen levels can negatively impact plant growth by decrease in chlorophyll production or its decomposition can lead to less intense photosynthesis and a lack of carbohydrates in the plant [ 30 ].

SPAD index, photosynthesis, and chlorophyll fluorescence

The results showed that chlorophyll content (SPAD index) was higher in leaves of plants fed with NH 4 + than those fed with NO 3 − at all O 2 concentration in nutrient solution (Fig.  5 ). The highest O 2 level caused a significant increase in chlorophyll content in NH 4 + fed plants, although different O 2 levels in nutrient solution do not affect chlorophyll content of NO 3 − grown plants. Increased chlorophyll content in NH 4 + -fed plants has been reported in cucumber [ 4 , 19 ]. Ammonium has been found to have varying effects on chlorophyll concentration in plants. Studies have shown that mild (5 mM) ammonium concentration can lead to an increase in chlorophyll content in certain plant species, independent of their tolerance capacity [ 31 ]. However, exposure to higher concentrations of ammonium has been associated with a decrease in chlorophyll content in plants, leading to oxidative stress and changes in the activity of antioxidative enzymes [ 29 , 32 ]. Therefore, the effect of ammonium on chlorophyll concentration in plants is dependent on the concentration of ammonium, and the specific plant species. The opposite results observed, where higher chlorophyll levels were found in NH 4 + -grown plants but with lower overall growth compared to nitrate-fed plants, can be explained as follows: The relationship between chlorophyll concentration and plant growth is intricate and can be influenced by various factors. For instance, studies have shown that NH 4 + can enhance foliar color and increase chlorophyll concentration, but at the same time, it can negatively impact plant growth, as observed in cucumber [ 4 ] and tomato [ 19 ]. Similar results were observed with shading on Kalmia latifolia cultivars [ 33 ], while salt stress decreased chlorophyll concentration and inhibited the growth of maize plants [ 34 ]. In our previous experiment, we also found that NH 4 + -grown plants exhibited higher nitrogen (N) uptake compared to nitrate-fed plants [ 4 ]. Additionally, higher concentrations of total amino acids were observed in the NH 4 + -fed plants [ 4 ]. Therefore, the higher chlorophyll concentration in NH 4 + -grown plants could be attributed to the increased N absorption by these plants. However, the elevated N concentration resulting from NH 4 + uptake can lead to nutrient imbalances within the plants [ 4 ]. Consequently, the relationship between chlorophyll concentration and plant growth is context-dependent, influenced by environmental conditions, and may vary across different plant species. In an experiment, it was observed that waterlogging (2 mg L -1 O 2 concentration) had a significant impact on the amount of chlorophyll in leaves of tropical tolerant trees. While some species showed little change, others exhibited no difference in chlorophyll levels [ 35 ]. The hypoxia stress treatment significantly inhibited Phyllostachys praecox plant growth. Leaf chlorophyll contents was initially improved and then reduced with plant growth time [ 36 ]. The decrease in chlorophyll content can be attributed to several factors. One possible reason for the decrease in chlorophyll is the reduction in enzymes responsible for synthesizing photosynthetic pigments [ 37 ]. Under stress conditions, the activity of the chlorophyllase enzyme tends to increase [ 38 ], leading to a breakdown of chlorophyll. Additionally, the biosynthesis of new chlorophyll is hindered as stress conditions promote the synthesis of other compounds like proline. This shift in synthesis pathways reduces the availability of glutamate, a precursor needed for both chlorophyll and proline production. The variation in chlorophyll levels among plants under oxygen stress conditions can also be influenced by factors such as root structure and defense mechanisms. Some plants may develop misplaced roots [ 35 ] or employ other defense mechanisms to cope with waterlogging, which could affect chlorophyll content. Previous studies have reported significant decreases in fresh and dry mass in roots and stems, as well as chlorophyll content in corn plants exposed to waterlogged conditions [ 39 ]. These findings support the results of the current research. Waterlogging conditions can cause visible changes in leaf appearance and physiological characteristics include closing of stomata, reductions in the rate of photosynthesis and uptake of essential mineral nutrients, as well as alterations in plant growth hormones, often resulting in leaf discoloration and eventual yellowing [ 40 ]. This can be attributed to alterations in leaf structure and function caused by waterlogging. Furthermore, studies have shown that the concentration of chlorophyll decreases during oxygen deprivation [ 41 ]. Increased respiration has been observed in plants supplied with NH 4 + [ 42 ], which requires higher oxygen levels. The assimilation of NH 4 + necessitates an adequate oxygen supply for root cell respiration and the provision of carbon skeletons from the Krebs cycle. Higher oxygen levels in the nutrient solution may mitigate NH 4 + toxicity by facilitating the detoxification of this ion through the provision of carbon skeletons for its incorporation into amino acids [ 43 ].

Effect of N-forms (at 5 mM) and dissolved O 2 levels (1.8 ± 0.2; 2.6 ± 0.2; 3.8 ± 0.2; 5.3 ± 0.2 mg. L − 1 ) of nutrient solution on leaf SPAD index (leaf chlorophyll content) of bell pepper. Different letters show significant different, Duncan’s test ( p  ≤ 0.01)

Nitrate has been found to improve plant tolerance to oxygen deficiency, with foliar NO 3 − assimilation being relevant to plant tolerance to oxygen deficiency [ 44 ]. Additionally, it has been observed that nitrogen application increases the plant’s tolerance to oxygen deficiency, with NO 3 − treated plants showing higher CO 2 assimilation and sucrose production compared to NH 4 + treated plants under flooding conditions [ 45 ]. The current experiment demonstrated that the rate of photosynthesis was significantly higher in leaves of plants that were fed with NO 3 − compared to those fed with NH 4 + in all O 2 concentration in nutrient solution (Fig.  6 ). Specifically, the plants fed with NO 3 − exhibited the highest leaf photosynthetic rate at 3.8 and 5.3 mg. L -1 O 2 levels, while the lowest rate was observed at O 2 levels of 1.8 and 2.6 mg. L -1 . These findings suggest that the detrimental impact of NH 4 + supply on pepper growth may primarily be attributed to the inhibition of net photosynthesis activity. Previous studies by Board [ 46 ] and Lizaso et al. [ 41 ] have reported that a deficiency in O 2 can reduce net photosynthesis. Furthermore, it was observed that leaves of plants fed with NO 3 − had a higher stomatal conductance compared to those fed with NH 4 + at O 2 levels of 2.6 ± 0.2 and 3.8 ± 0.2 mg. L -1 (Fig.  7 ). Pepper plants exhibited the highest stomatal conductance with NO 3 − nutrition at an O 2 level of 2.6 mg. L -1 and the lowest with NH 4 + nutrition at a level of 3.8 mg. L -1 O 2 . The nitrogen form did not affect stomatal conductance at the lowest and highest O 2 concentrations in the nutrient solution. Additionally, the sub-stomatal CO 2 concentration was found to be higher in leaves of NH 4 + -fed plants compared to those of NO 3 − -fed plants at 3.8 and 5.3 mg. L -1 O 2 levels. As shown in the current experiment (Fig.  6 ), NH 4 + nutrition can influence photosynthetic rates in plants. High levels of ammonium can inhibit photosynthesis, leading to a decrease in CO 2 fixation. This reduction in CO 2 assimilation can indirectly affect sub-stomatal CO 2 concentrations. In contrast to NH 4 + -fed plants, NO 3 − -supplied plants generally exhibited a trend of decreasing sub-stomatal CO 2 concentrations with increasing O 2 levels in the nutrient solution (Fig.  8 ). These results can be related to the higher rate of photosynthesis in these treatments (Fig.  6 ). Stomatal resistance was higher in leaves of NH 4 + -fed plants compared to those of NO 3 − -fed plants, with the highest stomatal resistance observed at 3.8 mg. L -1 O 2 in the nutrient solution for NH 4 + -grown plants (Fig.  9 ). This response is thought to be a mechanism by which plants regulate water loss, because, NH 4 + restricts the water uptake in plants [ 47 ]. Unlike NH 4 + -fed plants, O 2 levels had no effect on stomatal resistance in NO 3 − -supplied plants. Water use efficiency was higher in leaves of NO 3 − -fed plants compared to those of NH 4 + -fed plants at all O 2 concentration in nutrient solution (Fig.  10 ). NO 3 − -fed plants exhibited the highest water use efficiency at 3.8 and 5.3 mg. L -1 O 2 levels, while the lowest efficiency was observed at a level of 1.8 and 2.6 mg. L -1 O 2 . In NH 4 + -grown plants, different O 2 levels in the nutrient solution had no effect on water use efficiency. In contrast of the current experiment, research has shown that NH 4 + -fed Casuarina equisetifolia plants exhibited higher water use efficiency and lower water consumption compared to plants supplied with NO 3 – , regardless of the water supply conditions [ 48 ]. This higher water use efficiency in NO 3 – grown plants, as shown in the current experiment, was due to the higher rate of photosynthesis in these plants. The effect of oxygen deficiency in the nutrient solution on water use efficiency is a topic of interest in agricultural research. Studies have shown that oxygen deficiency in the nutrient solution can have immediate effects on the water uptake of plants [ 10 ]. Root asphyxia caused a decrease in water uptake [ 10 ]. Additionally, under root asphyxia conditions, plants may adapt to the new condition by relying on a metabolism of “nitrate respiration,” which could impact water and nitrate uptake processes, important factors for plant nutrition [ 10 ]. These findings highlight the importance of considering oxygen levels in the nutrient solution when aiming to optimize water use efficiency in plant cultivation. Moreover, the instantaneous carboxylation efficiency was higher in leaves of NO 3 − -fed plants compared to NH 4 + -fed plants (Fig.  11 ). Nitrate-fed plants demonstrated the highest instantaneous carboxylation efficiency at 3.8 and 5.3 mg. L -1 O 2 levels, while the lowest efficiency was observed at a level of 1.8 and 2.6 mg. L -1 O 2 . Similar to water use efficiency, different O 2 levels in the nutrient solution had no effect on the instantaneous carboxylation efficiency in NH 4 + -grown plants. The effect of ammonium nutrition on instantaneous carboxylation efficiency varies among plant species. Some studies have shown that ammonium nutrition can lead to a higher CO 2 assimilation rate per leaf area compared to nitrate nutrition, indicating a potentially higher instantaneous carboxylation efficiency [ 49 ]. However, other research suggests that the growth of plants under high ammonium nutrition may be impaired, which could potentially affect carboxylation efficiency [ 50 ]. Additionally, the redox metabolism and mitochondrial electron transport chain play a role in the response to ammonium nutrition, which may also impact carboxylation efficiency [ 51 ]. Overall, the effect of ammonium nutrition on instantaneous carboxylation efficiency is complex and may depend on various factors such as plant species, growth conditions, and the specific mechanisms involved in ammonium assimilation and tolerance. Lastly, leaf transpiration was found to be the highest in NO 3 − -fed plants at the two initial concentrations of O 2 (1.8 and 2.6 mg. L -1 ). The leaf transpiration of the other treatments remained at the same level (Fig.  12 ). The lowest leaf transpiration was observed in NH 4 + -fed plants at the 3.8 mg. L -1 O 2 . The effect of ammonium nutrition on transpiration is influenced by various factors, including the water lodging conditions. Research has shown that plants grown on ammonium nutrition exhibit lower water use efficiency compared to those receiving nitrate [ 52 ]. Additionally, the relationship between transpiration and soil water content suggests that once plants wilt, the transpiration rate should be roughly proportional to the available water content of the soil [ 53 ]. However, the specific interaction between ammonium nutrition and water lodging on transpiration requires further investigation.

Effect of N-forms (at 5 mM) and dissolved O 2 levels (1.8 ± 0.2; 2.6 ± 0.2; 3.8 ± 0.2; 5.3 ± 0.2 mg. L − 1 ) of nutrient solution on photosynthetic rate of bell pepper plants. Different letters show significant different, Duncan’s test ( p  ≤ 0.01)

Effect of N-forms (at 5 mM) and dissolved O 2 levels (1.8 ± 0.2; 2.6 ± 0.2; 3.8 ± 0.2; 5.3 ± 0.2 mg. L − 1 ) of nutrient solution on stomatal conductance of bell pepper plants. Different letters show significant different, Duncan’s test ( p  ≤ 0.01)

Effect of N-forms (at 5 mM) and dissolved O 2 levels (1.8 ± 0.2; 2.6 ± 0.2; 3.8 ± 0.2; 5.3 ± 0.2 mg. L − 1 ) of nutrient solution on sub-stomatal CO 2 concentration of bell pepper plants. Different letters show significant different, Duncan’s test ( p  ≤ 0.01)

Effect of N-forms (at 5 mM) and dissolved O 2 levels (1.8 ± 0.2; 2.6 ± 0.2; 3.8 ± 0.2; 5.3 ± 0.2 mg. L − 1 ) of nutrient solution on stomatal resistance of bell pepper plants. Different letters show significant different, Duncan’s test ( p  ≤ 0.01)

Effect of N-forms (at 5 mM) and dissolved O 2 levels (1.8 ± 0.2; 2.6 ± 0.2; 3.8 ± 0.2; 5.3 ± 0.2 mg. L − 1 ) of nutrient solution on water use efficiency of bell pepper plants. Different letters show significant different, Duncan’s test ( p  ≤ 0.01)

Effect of N-forms (at 5 mM) and dissolved O 2 levels (1.8 ± 0.2; 2.6 ± 0.2; 3.8 ± 0.2; 5.3 ± 0.2 mg. L − 1 ) of nutrient solution on mesophyll efficiency of bell pepper plants. Different letters show significant different, Duncan’s test ( p  ≤ 0.01)

Effect of N-forms (at 5 mM) and dissolved O 2 levels (1.8 ± 0.2; 2.6 ± 0.2; 3.8 ± 0.2; 5.3 ± 0.2 mg. L − 1 ) of nutrient solution on transpiration of bell pepper plants. Different letters show significant different, Duncan’s test ( p  ≤ 0.01)

The maximal quantum yield of PSII photochemistry, variable fluorescence, and maximum fluorescence were higher in NH 4 + grown plants compared to NO 3 − grown plants, but they increased in NO 3 − -fed plants with the elevation of O 2 concentration in the nutrient solution to 5.3 mg. L -1 , without significant difference with NH 4 + grown plants in the same O 2 level (Figs.  13 , 14 and 15 ). Minimum fluorescence also was higher in NH 4 + grown plants compared to NO 3 − grown plants, but it decreased with the elevation of O 2 concentration in the nutrient solution to 5.3 mg. L -1 to the level of NO 3 − grown plants, in the same O 2 level (Fig.  16 ). In NO 3 − grown plants, different O 2 levels in nutrient solution had no effect on the minimum fluorescence of leaves. Ammonium nutrition has been shown to enhance chlorophyll fluorescence, specifically the F v /F m parameter, in plants. A study on kohlrabi plants revealed that those given nitrogen as NH 4 + showed a 21% increase in chlorophyll content, along with a reduction in the chlorophyll a: b ratio and decreased ground state fluorescence compared to plants supplied with nitrate [ 54 ]. In agree with the results of the current experiment, it was observed that the NH 4 + -grown Salvinia natans plants exhibited a higher maximum quantum yield of PSII photochemistry (F v /F m ) in hypoxic and anoxic conditions compared to NO 3 − -fed plants [ 55 ]. However, Roosta et al. [ 19 ] reported that the form of nitrogen source does not affect the chlorophyll fluorescence of cucumber, which did not agree with the results of this research. The high nonphotochemical quenching shown in tomatoes fed with NH 4 + or urea indicated that PS II was the inhibitory site of NH 4 + -N which was directly uptaken by roots, or liberated via the urea hydrolysis cycle. However, in the current experiment opposite results were observed, which may be due to the lower concentration of NH 4 + in nutrient solution, and higher chlorophyll content in the NH 4 + -fed plants compared to the NO 3 − -fed plants [ 56 ]. Similar to the present study, Roosta and Schjoerring [ 4 ] found that NH 4 + nutrition at the medium concentration (5mM) compared to NO 3 − nutrition increased the chlorophyll concentration in cucumber leaves. Therefore, NH 4 + does not affect the reaction center of photosystem II at 5 mM because of several protective mechanisms. If the carbon dioxide supply becomes limiting due to decreasing stomatal conductance as it was shown in current experiment, photorespiration acts as an alternative electron sink for the light reaction [ 57 ]. The latter protects PSII from damage during NH 4 + stress, and therefore, F v /F m is not a good indicator for detecting plant NH 4 + response at mild NH 4 + stress [ 58 ].

Hypoxic stress has been found to significantly impact the chlorophyll fluorescence parameter F v /F m , which serves as a sensitive indicator of environmental stress in plants [ 59 ]. A study conducted on cucumber plants subjected to hypoxia treatment demonstrated a decrease in F v /F m , indicating the occurrence of photoinhibition in photosynthesis [ 60 ]. Interestingly, this outcome aligns with the results of the current experiment involving plants supplied with nitrate. These findings collectively underscore the detrimental effects of hypoxic stress on chlorophyll fluorescence and photosynthetic activity in plants.

Effect of N-forms (at 5 mM) and dissolved O 2 levels (1.8 ± 0.2; 2.6 ± 0.2; 3.8 ± 0.2; 5.3 ± 0.2 mg. L − 1 ) of nutrient solution on maximal quantum yield of PSII photochemistry (F v /F m ) of bell pepper plants. Different letters show significant different, Duncan’s test ( p  ≤ 0.01)

Effect of N-forms (at 5 mM) and dissolved O 2 levels (1.8 ± 0.2; 2.6 ± 0.2; 3.8 ± 0.2; 5.3 ± 0.2 mg. L − 1 ) of nutrient solution on variable fluorescence (F v ) of bell pepper plants. Different letters show significant different, Duncan’s test ( p  ≤ 0.01)

Effect of N-forms (at 5 mM) and dissolved O 2 levels (1.8 ± 0.2; 2.6 ± 0.2; 3.8 ± 0.2; 5.3 ± 0.2 mg. L − 1 ) of nutrient solution on maximum fluorescence (F m ) of bell pepper plants. Different letters show significant different, Duncan’s test ( p  ≤ 0.01)

Effect of N-forms (at 5 mM) and dissolved O 2 levels (1.8 ± 0.2; 2.6 ± 0.2; 3.8 ± 0.2; 5.3 ± 0.2 mg. L − 1 ) of nutrient solution on minimum fluorescence (F o ) of bell pepper plants. Different letters show significant different, Duncan’s test ( p  ≤ 0.01)

Conclusions

This study has found that the overall growth of pepper plants was significantly reduced by NH 4 + at low O 2 concentrations in nutrient solution. However, the highest levels of oxygen increased vegetative growth, particularly root growth in NH 4 + fed plants. The results also showed that chlorophyll content was higher in leaves of plants fed with NH 4 + than in those fed with NO 3 − at low O 2 concentrations in nutrient solution. The highest O 2 level caused a significant increase in chlorophyll content in NH 4 + fed plants. Photosynthetic rate, water use efficiency, and instantaneous carboxylation efficiency were all higher in the leaves of NO 3 − -fed plants compared to those of NH 4 + -fed plants at all O 2 levels. Nitrate-fed plants had the highest instantaneous carboxylation efficiency at 3.8 and 5.3 mg. L -1 O 2 levels and the lowest at levels of 1.8 and 2.6 mg. L -1 O 2 . Maximal quantum yield of PSII photochemistry, variable fluorescence, and maximum fluorescence were higher in NH 4 + grown plants compared to NO 3 − grown plants, but they increased in NO 3 − -fed plants with the elevation of O 2 concentration in the nutrient solution to 5.3 mg. L -1 , without significant difference with NH 4 + grown plants in the same O 2 level. Therefore, to grow healthy pepper plants in a floating hydroponic system, it is important to control the O 2 level and ensure it is not lower than 5.3 mg. L -1 .

Data availability

All the data generated or analyzed during the current study were included in the manuscript. The raw data is available from the corresponding author on reasonable request.

Zhonghua T, Yanju L, Xiaorui G, Yuangang Z. The combined effects of salinity and nitrogen forms on Catharanthus roseus: the role of internal ammonium and free amino acids during salt stress. J Plant Nutr Soil Sci. 2011;174:135–44.

Article   Google Scholar  

Britto DT, Kronzucker HJ. NH4 + toxicity in higher plants: a critical review. J Plant Physiol. 2002;159:567–84.

Article   CAS   Google Scholar  

Aluko OO, Li C, Yuan G, Nong T, Xiang H, Wang Q et al. Differential effects of ammonium (NH4+) and potassium (K+) Nutrition on Photoassimilate partitioning and growth of Tobacco Seedlings. Plants. 2022;11.

Roosta HR, Schjoerring JK. Effects of ammonium toxicity on nitrogen metabolism and elemental profile of cucumber plants. J Plant Nutr. 2007;30:1933–51.

Shilpha J, Song J, Jeong BR. Ammonium phytotoxicity and tolerance: an insight into ammonium Nutrition to Improve Crop Productivity. Agronomy. 2023;13.

Gerendás J, Zhu Z, Bendixen R, Ratcliffe RG, Sattelmacher B. Physiological and biochemical processes related to ammonium toxicity in higher plants. J Plant Nutr Soil Sci. 1997;160:239–51.

van Dongen JT, Licausi F, Nick P. Low-oxygen stress in plants: Oxygen Sensing and adaptive responses to Hypoxia. Plant Cell Monogr. 2014;21.

Roosta HR, Bagheri MH, Hamidpour M, Roozban MR. Interactive effects of nitrogen form and oxygen concentration on growth and nutritional status of eggplant in hydroponics. J Agric Sci Technol. 2016;18:731–9.

Google Scholar  

Chun C, Takakura T. Rate of Root respiration of lettuce under various dissolved oxygen concentrations in Hydroponics. Environ Control Biol. 1994;32:125–35.

Morard P, Lacoste L, Silvestre J. Effect of oxygen deficiency on uptake of water and mineral nutrients by tomato plants in soilless culture. J Plant Nutr. 2000;23:1063–78.

Morard P, Lacoste L, Silvestre J. Effect of oxygen deficiency on mineral nutrition of excised tomato roots. J Plant Nutr. 2004;27:613–26.

Brix H, Lorenzen B, Morris JT, Schierup H-H, Sorrell BK. Effects of oxygen and nitrate on ammonium uptake kinetics and adenylate pools in Phalaris arundinacea L. and Glyceria maxima (Hartm.) Holmb. Proc R Soc Edinb Sect B Biol Sci. 1994;102:333–42.

Greenway H, Gibbs J. Mechanisms of anoxia tolerance in plants. II. Energy requirements for maintenance and energy distribution to essential processes. Funct Plant Biol. 2003;30:999–1036.

Article   CAS   PubMed   Google Scholar  

Men S, Chen H, Chen S, Zheng S, Shen X, Wang C et al. Effects of supplemental nitrogen application on physiological characteristics, dry matter and nitrogen accumulation of winter rapeseed (Brassica napus L.) under waterlogging stress. Sci Rep. 2020;10.

Yetisir H, Çaliskan ME, Soylu S, Sakar M. Some physiological and growth responses of watermelon [Citrullus lanatus (Thunb.) Matsum. And Nakai] grafted onto Lagenaria siceraria to flooding. Environ Exp Bot. 2006;58:1–8.

Boru G, Vantoai T, Alves J, Hua D, Knee M. Responses of soybean to oxygen deficiency and elevated root-zone carbon dioxide concentration. Ann Bot. 2003;91:447–53.

Article   CAS   PubMed   PubMed Central   Google Scholar  

Strasser RJ, Srivastava A, Tsimilli-Michael M. The fluorescence transient as a tool to characterize and screen photosynthetic samples. Probing Photosynth Mech Regul Adapt. 2000;:443–80.

Roosta HR, Schjoerring JK. Response of tomato plant to ammonium and nitrate nutrition using the relative addition rate technique. Acta Hortic. 2021;1315:495–501.

Roosta HR, Sajjadinia A, Rahimi A, Schjoerring JK. Responses of cucumber plant to NH4 + and NO3- nutrition: the relative addition rate technique vs. cultivation at constant nitrogen concentration. Sci Hortic (Amsterdam). 2009;121:397–403.

Wheeler T. The Physiology of Crop Yield. Second edition., By RKM, Hay, Porter JR. Oxford: Blackwell Publishing (2006), pp. 314, £34.99(paperback). ISBN 1-4051-0859-2. Exp Agric. 2007;43:530–530.

Gamiely S, Randle WM, Mills HA, Smittle DA, Banna GI. Onion plant growth, Bulb Quality, and Water Uptake following ammonium and Nitrate Nutrition. HortScience. 2019;26:1061–3.

Takács E, Técsi L. Effects of NO3-/NH4 + ratio on photosynthetic rate, Nitrate Reductase Activity and Chloroplast Ultrastructure in three cultivars of Red Pepper (Capsicum annuum L). J Plant Physiol. 1992;140:298–305.

Guo S, Chen G, Zhou Y, Shen Q. Ammonium nutrition increases photosynthesis rate under water stress at early development stage of rice (Oryza sativa L). Plant Soil. 2007;296:115–24.

Yoshida S, Kitano M, Eguchi H. Growth of lettuce plants (Lactuca sativa L.) under control of dissolved O2 concentration in hydroponics. Biotronics. 1997;26:39–45.

Kläring HP, Zude M. Sensing of tomato plant response to hypoxia in the root environment. Sci Hortic (Amsterdam). 2009;122:17–25.

Roosta HR, Akbari A, Raghami M, Bikdeloo M. Response of growth, physiological characteristics and concentration of some mineral nutrients of local grafted watermelon to oxygen deficiency stress in hydroponic system. Iran J Hortic Sci. 2022;53:647–65.

Chérif M, Tirilly Y, Bélanger RR. Effect of oxygen concentration on plant growth, lipidperoxidation, and receptivity of tomato roots to Pythium F under hydroponic conditions. Eur J Plant Pathol. 1997;103:255–64.

Cakmak I. Possible roles of zinc in protecting plant cells from damage by reactive oxygen species. Rev New Phytol. 2000;146:185–205.

Biczak R. Quaternary ammonium salts with tetrafluoroborate anion: phytotoxicity and oxidative stress in terrestrial plants. J Hazard Mater. 2016;304:173–85.

Steffens D, Hütsch BW, Eschholz T, Lošák T, Schubert S. Water logging may inhibit plant growth primarily by nutrient deficiency rather than nutrient toxicity. Plant Soil Environ. 2005;51:545–52.

Sanchez-Zabala J, González-Murua C, Marino D. Mild ammonium stress increases chlorophyll content in Arabidopsis thaliana. Plant Signal Behav. 2015;10.

Wang J, Lu W, Tong Y, Yang Q. Leaf morphology, photosynthetic performance, chlorophyll fluorescence, stomatal development of lettuce (Lactuca sativa L.) exposed to different ratios of red light to blue light. Front Plant Sci. 2016;7.

Brand MH. Shade influences plant growth, leaf color, and chlorophyll content of Kalmia latifolia L. Cultivars. HortScience. 1997;32:206–8.

Turan MA, Elkarim AHA, Taban N, Taban S. Effect of salt stress on growth, stomatal resistance, proline and chlorophyll concentrations on maize plant. Afr J Agric Res. 2009;4:893–7.

Herrera A. Responses to flooding of plant water relations and leaf gas exchange in tropical tolerant trees of a black-water wetland. Front Plant Sci. 2013;4:MAY.

Ma J, Rukh G, Ruan Z, Xie X, Ye Z, Liu D. Effects of Hypoxia stress on growth, Root respiration, and metabolism of Phyllostachys praecox. Life. 2022;12.

Murkute AA, Sharma S, Singh SK. Studies on salt stress tolerance of citrus rootstock genotypes with arbuscular mycorrhizal fungi. Hortic Sci. 2006;33:70–6.

Reddy M, Vora A. Salinity induced changes in pigment composition and chlorophyllase activity of wheat. Indian J Plant Physiol. 1986;29:331–4.

CAS   Google Scholar  

Mahmood U, Hussain S, Hussain S, Ali B, Ashraf U, Zamir S et al. Morpho-Physio-biochemical and molecular responses of maize hybrids to salinity and waterlogging during stress and recovery phase. Plants. 2021;10.

Kozlowski TT. Soil aeration, flooding, and Tree Growth. Arboric Urban for. 1985;11:85–96.

Lizaso JI, Melendez LM, Ramirez R. Early flooding of two cultivars of tropical maize. II. Nutritional responses. J Plant Nutr. 2001;24:997–1011.

Kuzyakov Y, Gavrichkova O. REVIEW: Time lag between photosynthesis and carbon dioxide efflux from soil: a review of mechanisms and controls. Glob Chang Biol. 2010;16:3386–406.

Roosta HR, Schjoerring JK. Root carbon enrichment alleviates ammonium toxicity in cucumber plants. J Plant Nutr. 2008;31:941–58.

Oliveira HC, Freschi L, Sodek L. Nitrogen metabolism and translocation in soybean plants subjected to root oxygen deficiency. Plant Physiol Biochem. 2013;66:141–9.

de Carvalho PA, de Oliveira LEM, Domiciano D, de Carvalho JN, Prudente D, de O, Guimarães RJ. Effect of nitrogen source and oxygen deficiency on carbon metabolism and antioxidant system of rubber tree plants (Hevea spp). Aust J Crop Sci. 2018;12:116–25.

Board JE. Waterlogging effects on Plant nutrient concentrations in soybean. J Plant Nutr. 2008;31:828–38.

Cao X, Zhong C, Zhu C, Zhu L, Zhang J, Wu L, et al. Ammonium uptake and metabolism alleviate PEG-induced water stress in rice seedlings. Plant Physiol Biochem. 2018;132:128–37.

Martínez-Carrasco R, Pérez P, Handley LL, Scrimgeour CM, Igual M, Martín D, Molino I, et al. Regulation of growth, water use efficiency and δ13C by the nitrogen source in Casuarina equisetifolia Forst. And Forst. Plant. Cell Environ. 1998;21:531–4.

Guo S, Zhou Y, Shen Q, Zhang F. Effect of ammonium and nitrate nutrition on some physiological processes in higher plants - growth, photosynthesis, photorespiration, and water relations. Plant Biol. 2007;9:21–9.

Hoque MS, Masle J, Udvardi, MK, Ryan PR, Upadhyaya NM. Over-expression of the rice OsAMT1-1 gene increases ammonium uptake and content, but impairs growth and development of plants under high ammonium nutrition. Funct Plant Biol. 2006;:153–63.

Podgórska A, Ostaszewska M, Gardeström P, Rasmusson AG, Szal B. In comparison with nitrate nutrition, ammonium nutrition increases growth of the frostbite1 Arabidopsis mutant. Plant Cell Environ. 2015;38:224–37.

Article   PubMed   Google Scholar  

Ups SH, Leidi EO, Silberbush M, Soares MIM, Lewis OEM. Physiological aspects of ammonium and nitrate fertilization. J Plant Nutr. 1990;13:1271–89.

Gardner WR, Ehlig CF. The influence of soil water on transpiration by plants. J Geophys Res. 1963;68:5719–24.

Blanke MM, Bacher W, Pring RJ, Baker EA. Ammonium nutrition enhances chlorophyll and glaucousness in Kohlrabi. Ann Bot. 1996;78:599–604.

Jampeetong A, Brix H. Oxygen stress in Salvinia natans: interactive effects of oxygen availability and nitrogen source. Environ Exp Bot. 2009;66:153–9.

Nasraoui-Hajaji A, Gouia H. Photosynthesis sensitivity to NH4–N change with nitrogen fertilizer type. Plant Soil Environ. 2014;60:274–9.

Wilhelm C, Selmar D. Energy dissipation is an essential mechanism to sustain the viability of plants: the physiological limits of improved photosynthesis. J Plant Physiol. 2011;168:79–87.

Baker NR, Rosenqvist E. Applications of chlorophyll fluorescence can improve crop production strategies: an examination of future possibilities. J Exp Bot. 2004;55:1607–21.

Ball MC, Butterworth JA, Roden JS, Christian R, Egerton JJG. Applications of chlorophyll fluorescence to forest ecology. Aust J Plant Physiol. 1995;22:311–9.

Torzillo G, Bernardini P, Masojídek J. On-line monitoring of chlorophyll fluorescence to assess the extent of photoinhibition of photosynthesis induced by high oxygen concentration and low temperature and its effect on the productivity of outdoor cultures of Spirulina platensis (Cyanobacteria). J Phycol. 1998;34:504–10.

Download references

Acknowledgements

Not applicable.

The authors are grateful to the Arak University for funding this study.

Author information

Authors and affiliations.

Department of Horticultural Sciences, Faculty of Agriculture and Natural Resources, Arak University, Arak, Iran

Hamid Reza Roosta

You can also search for this author in PubMed   Google Scholar

Contributions

Hamid Reza Roosta: Conceptualization, Methodology, Software, Validation, Supervision, Visualization, Writing- Original Draft, Formal Analysis, Investigation, Resources, Data curation, Project administration, Writing - Review and preparation of final version. All authors have read and agreed to the published version of the manuscript.

Corresponding author

Correspondence to Hamid Reza Roosta .

Ethics declarations

Ethics approval and consent to participate.

No plants were collected in this experiment. Pepper seeds were purchased from Azar Abkesht Sahand Company (Tehran, Iran).

Consent for publication

Competing interests.

The authors declare no competing interests.

Additional information

Publisher’s note.

Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.

Rights and permissions

Open Access This article is licensed under a Creative Commons Attribution 4.0 International License, which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons licence, and indicate if changes were made. The images or other third party material in this article are included in the article’s Creative Commons licence, unless indicated otherwise in a credit line to the material. If material is not included in the article’s Creative Commons licence and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder. To view a copy of this licence, visit http://creativecommons.org/licenses/by/4.0/ . The Creative Commons Public Domain Dedication waiver ( http://creativecommons.org/publicdomain/zero/1.0/ ) applies to the data made available in this article, unless otherwise stated in a credit line to the data.

Reprints and permissions

About this article

Cite this article.

Roosta, H.R. The responses of pepper plants to nitrogen form and dissolved oxygen concentration of nutrient solution in hydroponics. BMC Plant Biol 24 , 281 (2024). https://doi.org/10.1186/s12870-024-04943-7

Download citation

Received : 14 December 2023

Accepted : 25 March 2024

Published : 13 April 2024

DOI : https://doi.org/10.1186/s12870-024-04943-7

Share this article

Anyone you share the following link with will be able to read this content:

Sorry, a shareable link is not currently available for this article.

Provided by the Springer Nature SharedIt content-sharing initiative

• Capsicum annuum
• Soilless culture

BMC Plant Biology

ISSN: 1471-2229

• General enquiries: [email protected]

WVU researcher studying worst western US megadrought in 1,200 years

Wednesday, April 17, 2024

A West Virginia University researcher is working to gain a better understanding of the 23-year megadrought that is affecting drylands in the western United States. The megadrought is an ongoing climate crisis for natural ecosystems, agricultural systems and human water resources, but researchers have a limited understanding of the phenomenon. (Submitted Photo)

Drylands in the western United States are currently in the grips of a 23-year “megadrought,” and one West Virginia University researcher is working to gain a better understanding of this extreme climate event.

Steve Kannenberg , assistant professor of biology at the WVU Eberly College of Arts and Sciences , is using observations from existing networks of scientific instrument stations across the region to inch toward that goal.  

The megadrought is an ongoing climate crisis for natural ecosystems, agricultural systems and human water resources, but researchers have a limited understanding of the phenomenon.

With joint National Science Foundation funding from Ecosystem Science Cluster and the Established Program to Stimulate Competitive Research, commonly known as EPSCoR, Kannenberg is seeking to identify where this drought has been most severe.

Data should reveal where the conditions have depleted groundwater and soil moisture and identify which dryland plants have been most affected.

The term “drylands” refers to areas where water availability limits the health of ecosystems.

“In West Virginia, we have plenty of water,” he said. “But, if you go out to Utah, for example, it’s very hot, very dry. And the health of the vegetation is determined by how much water is in the soil and how much water is in the air.”

Data on the west’s climatological history can be obtained by studying tree growth rings in drylands. Using tree rings, researchers have found the current 23-year drought period is the most severe over the last 1,200 years. Kannenberg will pair tree ring data with measurements of soil moisture, groundwater and ecosystem fluxes via eddy covariance flux towers.

“These are, essentially, fancy weather stations that can sense the ecosystem breathing,” he said. “It can quantify how much carbon is going into the vegetation from the atmosphere as plants photosynthesize during the day, and likewise, how much carbon is breathed out back into the atmosphere at night, because ecosystems respire like we do.”

The towers can also measure how much water is coming in via rain, how much goes out through plants to the atmosphere and how much evaporates from the soil surface.

Globally, megadroughts are projected to increase in frequency and severity in the coming decades, and Kannenberg’s synthesized data may help inform researchers about other dryland and non-dryland biomes.

He’s also focused on carbon capture. The photosynthetic rate of the vegetation across drylands affects their ability to store carbon, but trees can only photosynthesize when there’s sufficient water available. This process is fairly consistent in eastern forests, but difficult to predict in drylands.

“If you think of a forest here in West Virginia, there’s obviously a lot of carbon stored in the vegetation,” he said. “This makes it a very important carbon sink, globally. It’s easy for scientists to predict how much carbon gets taken up by these trees every year because we know that the environment during the spring, summer and fall is pretty conducive to photosynthesis.”

However, with far less vegetation in western landscapes, less carbon is stored in drylands. Water availability is inconsistent and unpredictable, and the amount of carbon western vegetation can take up each year varies significantly. In drought years, little carbon may be absorbed at all.

“Studies show dryland ecosystems in particular are really important for determining how much carbon gets taken up by the whole Earth’s surface globally,” Kannenberg said. “Not because they take up a ton of carbon, but because they’re so inconsistent over time. Understanding photosynthesis and carbon storage in these dryland ecosystems is important, even though it might not look like there’s a ton of carbon stored in the vegetation on the landscape.”

Kannenberg said there are various management actions available to help mitigate some of the current impacts and prepare for those to come, because as the planet gets hotter, the atmosphere gets drier. In many regions, like the southwestern U.S., which are already very dry, feedback loops warm the air and dry the atmosphere, which in turn will accelerate future drought events.

“Historically, megadroughts are a rare, rare thing,” he said. “But there have been a number of them throughout time, and they’re going to get more frequent and more severe in the future.”

MEDIA CONTACT: Laura Jackson Research Writer WVU Research Communications 304-215-1019; [email protected]

Call 1-855-WVU-NEWS for the latest West Virginia University news and information from WVUToday .