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Glyphosate-Dependent Inhibition of Photosynthesis in Willow

Marcelo p. gomes.

1 Ecotoxicology of Aquatic Microorganisms Laboratory, GRIL, TOXEN, Department of Biological Sciences, Université du Québec à Montréal, Montréal, QC, Canada

2 Laboratório de Fisiologia Vegetal, Instituto de Ciências Biológicas, Departamento de Botânica, Universidade Federal de Minas Gerais, Belo Horizonte, Brazil

Sarah G. Le Manac’h

Louise hénault-ethier.

3 Institut des Sciences de l’Environnement, Université du Québec à Montréal, Montréal, QC, Canada

Michel Labrecque

4 Institut de Recherche en Biologie Végétale, Montreal Botanical Garden, Montréal, QC, Canada

Marc Lucotte

Philippe juneau, associated data.

We studied the physiological mechanisms involved in the deleterious effects of a glyphosate-based herbicide (Factor ® 540) on photosynthesis and related physiological processes of willow ( Salix miyabeana cultivar SX64) plants. Sixty-day-old plants grown under greenhouse conditions were sprayed with different rates (0, 1.4, 2.1, and 2.8 kg a.e ha -1 ) of the commercial glyphosate formulated salt Factor ® 540. Evaluations were performed at 0, 6, 24, 48, and 72 h after herbicide exposure. We established that the herbicide decreases chlorophyll, carotenoid and plastoquinone contents, and promotes changes in the photosynthetic apparatus leading to decreased photochemistry which results in hydrogen peroxide (H 2 O 2 ) accumulation. H 2 O 2 accumulation triggers proline production which can be associated with oxidative protection, NADP + recovery and shikimate pathway stimulation. Ascorbate peroxidase and glutathione peroxidase appeared to be the main peroxidases involved in the H 2 O 2 scavenging. In addition to promoting decreases of the activity of the antioxidant enzymes, the herbicide induced decreases in ascorbate pool. For the first time, a glyphosate-based herbicide mode of action interconnecting its effects on shikimate pathway, photosynthetic process and oxidative events in plants were presented.

Introduction

Glyphosate [ N -(phosphonomethyl)glycine)] is the most broadly used herbicide worldwide since the introduction of glyphosate-resistant (GR) plants ( Coupe et al., 2012 ). Although it has been suggested as one of the least toxic pesticides to animals and humans ( Williams et al., 2000 ; Cerdeira and Duke, 2006 ), the widespread use of glyphosate together with its great solubility trigger some concerns about its possible effects on the environment.

Glyphosate negative effects on non-target plants ( Bott et al., 2011 ) and aquatic organisms ( Vendrell et al., 2009 ; Inderjit and Kaushik, 2010 ) have been largely described. By inhibiting the EPSP synthase (EC 2.5.1.19), glyphosate-based herbicides prevent biosynthesis of aromatic amino acids ( Siehl, 1997 ) leading to shikimic acid accumulation ( Duke and Powles, 2008 ). The depletion of the aromatic amino acid pool leads to a reduction of protein synthesis necessary to growth maintenance ( Siehl, 1997 ). On the other hand, in some plants, aromatic amino acid deficiencies upon glyphosate application has not been found, although deleterious effects of the herbicide have been observed ( Lee, 1981 ; Wang, 2001 ; Serra et al., 2013 ). This indicates that glyphosate can affect other plant-physiological processes ( Gomes et al., 2014b ). Numerous studies demonstrated decreases in the photosynthetic rate of plants following treatment with glyphosate ( Mateos-Naranjo et al., 2009 ; Yanniccari et al., 2012 ; Zobiole et al., 2012 ). In glyphosate-sensitive plants, the herbicide causes inhibition of CO 2 assimilation ( Diaz Vivancos et al., 2011 ) and depletion of intermediates of the photosynthetic carbon reduction cycle ( Servaites et al., 1987 ) which could be linked to the unregulated flux of carbon into the shikimate pathway ( Siehl, 1997 ). Moreover, glyphosate can indirectly affect photosynthesis by inhibiting chlorophyll biosynthesis ( Fedtke and Duke, 2005 ) or inducing chlorophyll degradation ( Gomes et al., 2016a ), decreasing stomatal conductance ( Yanniccari et al., 2012 ), and provoking nutritional disturbances ( Cakmak et al., 2009 ; Su et al., 2009 ). Nowadays, a special attention has been giving to understand glyphosate-induced oxidative stress in plants ( Gomes et al., 2014b ).

Reactive oxygen species are essential in plant signaling; however, once accumulated, ROS become toxic, inducing irreversible changes in metabolism, cell cycle, and increase oxidative bursts ( Gomes et al., 2014a ). By interacting with biological molecules, ROS can induce destruction of DNA, lipids, and proteins ( Foyer and Noctor, 2011 ). To avoid oxidative damage due to ROS accumulation, plants have developed enzymatic (e.g., SOD, CAT, APX, GPX, and GR) and non-enzymatic (e.g., ascorbate and glutathione) systems ( Foyer and Noctor, 2011 ). The activity of antioxidant systems as well as the lipid peroxidation extent are oxidative stress markers which were shown to be modulated by glyphosate exposure ( Ahsan et al., 2008 ; Moldes et al., 2008 ; Miteva et al., 2010 ).

Glyphosate effects on photosynthesis of non-resistant plants were associated to the herbicide induced decreases in the abundance of photosynthetic pathway proteins together with the oxidation of the major redox pools ( Diaz Vivancos et al., 2011 ). However, it has been reported that glyphosate can also induce ROS accumulation ( Ahsan et al., 2008 ; Moldes et al., 2008 ; Miteva et al., 2010 ; Gomes et al., 2016a ) and glyphosate-resistance was related to the ability of plants to avoid oxidative bursts through activation of antioxidant systems ( Maroli et al., 2015 ). Photosynthesis-targeting herbicides, such as atrazine, are known to induce oxidative stress by inhibiting Hill’s reactions ( Fedtke and Duke, 2005 ). Plants exposed to these kinds of herbicides are not able to cope with the mass of triplet chlorophyll molecules produced due the blockage of the electron transport flow, resulting in cell oxidative bursts due to ROS accumulation. On the other hand, it is not clear how glyphosate can induce ROS accumulation in plants and if the oxidative stress induced by the herbicide could also be related to the observed decreases in photosynthesis. We hypothesized that the interference on shikimate pathway could induce ROS production and consequently affect photosynthesis of exposed plants. Therefore, in this study we accessed the physiological mechanisms involved in the deleterious effects of a glyphosate-based herbicide (Factor ® 540) on photosynthesis of willow ( Salix miyabeana cultivar SX64) plants. For the first time, a glyphosate-based herbicide mode of action interconnecting glyphosate effects on shikimate pathway, plant photosynthetic process and oxidative events were described.

Materials and Methods

Greenhouse experiments.

Salix miyabeana cultivar SX64 was chosen for this study due to its high tolerance to stress factors, fast growth and great biomass production ( Labrecque and Teodorescu, 2005 ). Moreover, this species has been indicated for phytoremediation programs, particularly in the context of riparian buffer strips, to reclaim agricultural contaminants ( Gomes et al., 2016b ). Cuttings of S. miyabeana approximately 20 cm long were grown in plastic boxes (35 l) filled with distilled water amended with King Max nutrient solutions A (7% P 2 O 5 , 11% K 2 O, 1.5% Mg, 1.27% S, 0.07% B, 0.002% Mo, 0.12% Zn) and B (4% N, 1% NH 4 + , 3% NO 3 -2 , 10% K 2 O, 2% Ca, 0.05% Fe, 0.05% Mn) (Montreal, QC, Canada), following the product’s instructions. The solutions were continuously aerated, and renewed every 15 days. The pH of the medium was checked and adjusted on a weekly basis to 6.5 ± 0.1. The greenhouse was maintained at 25/22°C (±3°C) day/night temperature with natural light supplemented by sodium vapor lamps to provide a 12 h photoperiod and an average photosynthetic active radiation of 825 μmol photons m -2 s -1 . After an initial growth period (45 days), rooted, healthy (without leaf chlorotic spots) and uniform (similar height) plants were used in all treatments. A randomized block design with seven containers (corresponding to the replicates) per treatment, in a 4 (herbicide concentrations) × 4 (times of evaluation) factorial scheme was used. One hundred microliters of a freshly prepared herbicide solutions were hand-sprayed uniformly on each of the first three fully expanded leaves (corresponding to seventh to ninth leaves counting down from the shoot apex). This spray volume did not result in any runoff from the leaves. The herbicide (0, 56.15, 84.21, and 112.30 mM of glyphosate) applied concentrations were equivalent to field applications of 0, 1.4, 2.1, and 2.8 kg glyphosate ha -1 , which represent scenarios of 50, 75, and 100% of the standard field herbicide concentration applied in agricultural areas in Quebec ( Gomes et al., 2016a ).

Photosynthetic (using chlorophyll fluorescence kinetic measurements) and biochemical evaluations were performed at 0, 6, 24, 48, and 72 h after the beginning of the treatments. The evaluations were stopped after 72 h of exposure as plants from the highest glyphosate treatment showed pronounced intoxication symptoms, including several necrotic spots and loss of leaves (data not shown). After photosynthetic and stomatal conductance evaluations, plants were harvested and thoroughly washed with distilled water. Samples of the seventh (first fully expanded leaf from the apex) to ninth leaves were immediately frozen in liquid nitrogen and stored in aluminum foil paper at -80°C until biochemical evaluations and oxidative damage evaluations.

Gas Exchange, Chlorophyll Fluorescence, and Pigment Concentrations

Gas exchange, chlorophyll fluorescence, and pigment contents were measured on samples from the first, second, and third fully expanded leaves (seventh–ninth leaves from the apex), which also received the herbicide, for a total of three measurements per plant. Measurements of stomatal conductance ( g s ) were performed using a leaf porometer (model SC-1, Decadon Devices Inc., Washington, DC, USA). Then, these leaves were dark-acclimated for 20 min and the chlorophyll fluorescence emission was assessed using a pulse-amplitude modulation (PAM) fluorometer (model PAM-2500, WALZ, Effeltrich, Germany). A RLC analysis was performed according to Juneau et al. (2015) . An 11 steps RLC was performed. Saturating pulses were triggered at 0.8 min intervals with varying actinic light intensity for each step (0, 31, 48, 76, 117, 179, 253, 405, 586, 874, and 1326 μmol photons m -2 s -1 ). Using the RLC, the evaluation of the following parameters was performed: the ETR ( Krall and Edwards, 1992 ), the qP ( van Kooten and Snel, 1990 ), the UQF rel ( Juneau et al., 2005 ), the NPQ ( Redondo-Gómez et al., 2008 ), and the F V / F M ( Kitajima and Butler, 1975 ). To compare treatments, fluorescence results from the 874 μmol photons m -2 s -1 (most similar irradiation in relation to light growth conditions) were used. Curves of ETR versus irradiance were also plotted and the ETR max and the I k were calculated according to Eilers and Peeters (1988) .

For pigments evaluations, three foliar disks of approximately 5 mm in diameter were taken from each leaf, and after determining the fresh weight of the samples, their chlorophyll and carotenoid pigments were extracted in 80% acetone after macerating the disks with a mortar and pestle. The spectral absorption of the extracts (from 300 to 800 nm) was measured using a Varian Cary ® 300 Bio UV-Vis spectrophotometer (Varian, USA). The concentrations (μg/g fresh leaf weight) of total chlorophylls and carotenoids were calculated using the equations described by Lichtenthaler and Wellburn (1983) .

Biochemical Evaluations

Shikimate and proline concentrations were evaluated following the methods described in Bates et al. (1973) and Bijay and Dale (1998) , respectively. To evaluate the pool of quinones in leaves, 0.1 g of fresh plant tissue was ground in liquid nitrogen, homogenized in 1000 μl of freeze-cold ethyl acetate and then centrifuged for 1 min at 6.590 × g ( Kruk and Karpinski, 2006 ). The supernatant was then transferred to a collecting tube and the procedure was repeated twice (by adding 1000 μl of freeze-cold ethyl acetate to the pellet) to assure high extraction efficiency. Ten microliters of cold 1 M sodium borohydride (NaBH 4 ) was added to the combined supernatant to convert quinone to its reduced form and then, samples were centrifuged for 2 min at 10.000 × g to remove impurities ( Yoshida et al., 2010 ). The standard of plastoquinone (PQ-9, 1 mM) was acquired from the laboratory of J. Kruk (Jagiellonian University, Poland). After dilution in ethanol, the amount of 20 μl of cold 1 M NaBH 4 was added to assure complete reduction of plastoquinone pool. The UHPLC (Agilent 1290 Infinity II LC, Wilmington, DE, USA) measurements were performed according to Yoshida et al. (2010) , using UV-VIS detector, fluorescence detector, column (50 mm × 2.1 mm) isocratic solvent system (methanol/hexane, 340/20 vol/vol), flow rate of 0.31 ml/min, absorption detection wavelength at 255 nm, fluorescence excitation/emission detection at 290/330 nm, and injection volume of 1 μl.

To assess oxidative responses, H 2 O 2 , MDA contents and the activity of antioxidant systems were studied following the methods described by Gomes et al. (2014c) . H 2 O 2 was extracted in 2 ml of 0.1% trichloroacetic acid (TCA) and after centrifugation at 12000 × g for 15 min, 300 μl of the centrifuged supernatant was reacted with 0.5 ml of 10 mM potassium phosphate buffer (pH 7.0) and 1 ml of 1 M KI. Samples were read at 390 nm and H 2 O 2 concentrations were determined using an extinction coefficient (𝜀) of 0.28 mM -1 cm -1 . The estimation of lipid peroxidation was based on the production of 2-thiobarbituric acid reactive metabolites, particularly MDA. Samples containing 200 mg of leaf and root tissue were macerated in 5 mL of 0.1% TCA. After complete homogenization, 1.4 mL of the homogenate was transferred to an eppendorf tube and centrifuged at 10,000 rpm for 5 min. An aliquot of 0.5 mL of the supernatant was added to 2 mL 0.5% (v/v) TBA (thiobarbituric acid) in 20% TCA. The mixture was heated in a water bath at 95°C for 30 min and then ice-cooled for 10 min. Readings were taken using a spectrophotometer at 535 and 600 nm.

To study the antioxidant enzymes, 0.1 g of leaves were macerated in 800 μl of an extraction buffer containing 100 mM potassium buffer (pH 7.8), 100 mM EDTA, 1 mM L -ascorbic acid and 2% PVP (m/v). The protein contents of samples were determined using the Bradford method. Activities of SOD (EC 1.15.1.1), CAT (EC1.11.1.6), APX (EC 1.11.1.11), GPX (E.C. 1.11.1.9), and GR (E.C. 1.6.4.2) were assessed. To evaluate the ascorbate pool [total ascorbate (AsA + DHA), AsA and DHA], 0.2 g of frozen tissue were ground in liquid nitrogen in a mortar and pestle and homogenized with 5 ml of 6.5% (w/v) m -phosphoric acid containing 1 mM NaEDTA.

Statistical Analyses

Results were expressed as the average of three replicates. Statistical analyses were performed using JMP software 10.0 (SAS Institute Inc). Results were submitted to normality (Shapiro–Wilk) and homogeneity (Bartlett) tests and then statistically evaluated. MANOVA univariate repeated measures, with Time as the within-subject factor and the herbicide concentrations as the main effects, were used to analyze differences in the variables studied during exposure to the treatments. Glyphosate, Time, and the interaction between glyphosate and time were included within the model. The sphericity of the data was tested by the Mauchly’s criteria to determine whether the univariate F tests for the within-subject effects were valid. In cases of invalid F , the Greenhouse–Geisser test was used to estimate epsilon (𝜀). Contrast analysis was used when there were significant differences in the variables between treatments (Supplementary Tables 1S and 2S ).

Pigment Content, Gas Exchange, and Chlorophyll Fluorescence

Total chlorophyll and plastoquinone concentrations were decreased in leaves of plants by herbicide exposure and by treatment time ( P > 0.001; Figure ​ Figure1 1 ). The carotenoid concentration was greater in herbicide-treated plants at 6 h for all applied doses ( Figure ​ Figure1 1 ); then, carotenoid concentration was decreased in plants exposed for at least 24 h to herbicide concentrations ( P < 0.0001). The stomatal conductance was decreased in herbicide-exposed plants for all the treatment times ( P < 0.05; Figure ​ Figure1 1 ). Similar effects were observed on the ETR max , the I k , and the qP, which were significantly reduced in treated plants ( P < 0.0001; Figure ​ Figure2 2 ). However, for the first evaluation (6 h), ETR max , I k , and qP were not decreased in plants treated with 1.4 kg a.e ha -1 ( P > 0.05; Figure ​ Figure2 2 ). The UQF rel increased in all treated plants ( Figure ​ Figure1 1 ). Concomitantly, the NPQ decreased in plants exposed for more than 24 h to the herbicide ( P < 0.05; Figure ​ Figure2 2 ). The maximal PSII photochemical efficiency ( F V / F M ) was decreased in herbicide-treated plants ( P < 0.0001). Decreased F V / F M was seen in plants treated with 1.4 kg a.e ha -1 only after 72 h of herbicide exposure ( P < 0.05; Figure ​ Figure2 2 ). Plants exposed to 2.1 kg a.e ha -1 showed decreases in F V / F M at 48 and 72 h of exposure ( P < 0.001). In contrast, in all the evaluations, plants exposed to 2.8 kg a.e ha -1 showed decreased F V / F M ( P < 0.01; Figure ​ Figure2 2 ).

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Time courses of pigment (total chlorophyl and carotenoids) concentrations, plastoquinone pool (PQ), and stomatal conductance ( g s ) in leaves of Salix miyabeana (cultivar SX64) plants spread with doses of increased (0, 1.4, 2.1, and 2.8 kg a.e ha -1 ) rates of the glyphosate based herbicide (Factor ® 540). Values are means ± SE of three replicates.

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Time courses of photosynthesis-related measurements [maximum electron transport rate (ETR max ), minimum saturating irradiance ( I k ), photochemical quenching (qP), non-photochemical quenching (NPQ), relative unquenched fluorescence (UQF rel ), and maximal photochemical efficiency of PSII ( F V / F M )] in leaves of Salix miyabeana (cultivar SX64) plants spread with doses of increased (0, 1.4, 2.1, and 2.8 kg a.e ha -1 ) rates of the glyphosate based herbicide (Factor ® 540). Values are means ± SE of three replicates.

Shikimate and Proline Contents

The shikimate and proline concentrations in leaves of herbicide-treated plants were always higher than the control ( P < 0.0001; Figure ​ Figure3 3 ). In plants exposed to 2.1 and 2.8 kg a.e ha -1 , an important shikimate accumulation was found after 72 h of herbicide-treatment ( P < 0.0001).

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Time courses of shikimate, proline and hydrogen peroxide concentrations, and lipid peroxidation (MDA concentrations) in leaves of Salix miyabeana (cultivar SX64) plants spread with doses of increased (0, 1.4, 2.1, and 2.8 kg a.e ha -1 ) rates of the glyphosate based herbicide (Factor ® 540). Values are means ± SE of three replicates.

H 2 O 2 Contents and Lipid Peroxidation

Compared to control, H 2 O 2 concentration was always higher in plants exposed to the herbicide ( P < 0.001; Figure ​ Figure3 3 ), and greatly increased in these plants after 72 h ( P < 0.01). Similarly, lipid peroxidation (MDA concentration) was always higher in plants exposed to the herbicide (Supplementary Table 2S ; Figure ​ Figure3 3 ). In all plants, MDA content slightly increased at 24 h ( P > 0.05). However, in plants treated with herbicide, a pronounced increase in MDA concentration was observed at 72 h ( P < 0.05).

Antioxidant Responses

Plants treated with herbicide showed higher activity of all evaluated antioxidant enzymes after 6 h in relation to control ( P < 0.05; Figure ​ Figure4 4 ). We found that: (1) SOD and APX activities were higher in herbicide-treated plants up to 24 h ( P < 0.0001), and then were reduced for the following exposure times ( P < 0.0001); (2) CAT activity was always higher in plants treated with herbicide ( P < 0.0001); (3) similar to SOD and APX, GPX activity was also reduced in herbicide treated plants at 48 and 72 h of exposure ( P < 0.0001); (4) GR activity was higher in herbicide treated plants up to 48 h of exposure ( P < 0.05). Regarding ascorbate pool ( Figure ​ Figure5 5 ) we found that, in relation to control: (1) total ascorbate concentrations (AsA + DHA) were higher in herbicide-treated plants up to 24 h of exposure, and then were reduced for the following exposure times ( P < 0.0001); (2) the concentrations of the ascorbate reduced form (AsA) were greater in control plants up to 24 h, did not differ between treatments at 48 h and was increased in herbicide treated plants for 72 h ( P < 0.0001); (3) the concentrations of oxidized form of ascorbate (DHA) were greater in herbicide treated plants up to 24 h and were reduced for the following exposure duration ( P < 0.0001); (4) the AsA/DHA ratio was lower in 6 and 24 h treated plants compared to control, but was higher for the following treatment times ( P < 0.0001).

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Time courses of superoxide dismutase (SOD), catalase (CAT), ascorbate peroxidase (APX), glutathione peroxidase (GPX), and glutathione reductase (GR) activities in leaves of Salix miyabeana (cultivar SX64) plants spread with doses of increased (0, 1.4, 2.1, and 2.8 kg a.e ha -1 ) rates of the glyphosate based herbicide (Factor ® 540). Values are means ± SE of three replicates.

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Time courses of total ascorbate (AsA + DHA), reduced ascorbate (AsA), oxidized ascorbate (DHA), and AsA/DHA ratio in leaves of Salix miyabeana (cultivar SX64) plants spread with doses of increased (0, 1.4, 2.1, and 2.8 kg a.e ha -1 ) rates of the glyphosate based herbicide (Factor ® 540). Values are means ± SE of three replicates.

In this study, for the first time, a wide investigation of the impacts of glyphosate-based herbicide on several physiological processes was done. We demonstrated that this type of herbicide affected not only the shikimate pathway, but several physiological processes in willow plants as previously reported by Gomes et al. (2016b) . Figure ​ Figure6 6 represents an integrative model interconnecting the studied physiological parameters (in particular, shikimate pathway, photosynthetic processes and oxidative events) affected by exposure to a glyphosate-based herbicide greater than 24 h (48 and 72 h). The various steps of this model are identified throughout the text as Figure ​ Figure6 6 , #1–19.

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Interconnected model of the effects of the glyphosate-based-herbicide (Factor ® 540) on shikimate pathway, photosynthesis and oxidative markers of willow plants. Numbers refer to the ones mentioned in the discussion. ABA, abscisic acid; ALA, δ-aminolevulinic acid; APX, ascorbate peroxidase; AsA, ascorbate; EPSPS, 5-enolpyruvylshikimate-3-phosphate synthase; ETR, electron transport rate; F V / F M , maximal PSII photochemical efficiency; GPX, glutathione peroxidase; g s , stomatal conductance; H 2 O 2 , peroxide; I k , minimum saturating irradiance; MDA, lipid peroxidation; NPQ, non-photochemical quenching; qP, photochemical quenching; SOD, superoxide dismutase; UQF rel , the relative unquenched fluorescence. Literature-based information in the models are expressed in italic words and in dotted arrows. While, observed data obtained in the present study are introduced in the model as bold words and non-dotted arrows.

The glyphosate-based herbicide clearly inhibited the shikimate pathway in willow plants, as demonstrated by the shikimate accumulation ( Figure ​ Figure3 3 ) and also reported by Huang et al. (2012) and Gomes et al. (2016b) . By inhibiting the shikimate pathway ( Figure ​ Figure6 6 , #1), the glyphosate-based herbicide may prevent the biosynthesis of several secondary plant compounds ( Siehl, 1997 ), including plastoquinones ( Figure ​ Figure1 1 ; Figure ​ Figure6 6 , #2). It is known that UQF rel is an indicator of closed PSII reaction centers (RCs) present under continuous illumination ( Juneau et al., 2005 ) and qP represents the proportion of open PSII RCs ( Maxwell and Johnson, 2000 ). Therefore, the higher UQF rel and lower qP in treated plants indicate that the plastoquinone pool, and thus the PSII RCs, were in a more reduced state than in control plants, a consequence of a lower PQ content ( Figure ​ Figure6 6 , #2 and 3) and/or less effective PSI. This may, together with the decrease in the I k (for doses > 1.4 kg a.e ha -1 ), have contributed to the observed lower ETR in treated plants ( Figure ​ Figure2 2 ; Figure ​ Figure6 6 , #4). Indeed, a lower ability for PSII to deliver electrons to the electron transport chain, leading to PSII saturation at low irradiance, may explain the observed decrease in photosynthesis observed here and in previous studies ( Huang et al., 2012 ; Yanniccari et al., 2012 ; Gomes et al., 2016b ). However, we demonstrated that other effects of the glyphosate-based herbicide have also caused the decrease in the ETR (see below).

The observed increase in the carotenoid concentration after 6 h in the herbicide-treated plants ( Figure ​ Figure1 1 ) could be related to the concomitant increase in H 2 O 2 concentration, since it is known that ROS presence can induce carotenogenic responses ( Fan et al., 1998 ). Indeed, by the activation of latent biosynthetic enzymes (such as glutathione transferase and glutathione reductase) or by the expression of genes coding for carotenogenic enzymes, ROS may regulate carotenoid concentration ( Aniya and Anders, 1992 ; Bouvier et al., 1998 ). Since the maximal PSII photochemical yield ( F V / F M ) is a proxy of the PSII integrity ( Walter et al., 2003 ), F V / F M up to 24 h in treated plants (with the exception of the highest dose; Figure ​ Figure2 2 ) indicates that the glyphosate-based herbicide had no effect on the PSII integrity up to 24 h of exposure. Similarly, negative effects of glyphosate in F V / F M of Lolium perenne plants were only observed after 3 days of exposure ( Yanniccari et al., 2012 ). This may be the consequence of the increased carotenoid concentration helping to prevent ROS-damages to PSII ( Gomes et al., 2013b ). Carotenoids are usually involved in the protection of the oxidative damage by the detoxification of oxygen singlets ( 1 O 2 ) produced by photosynthesis or by enzymatic conversion of other ROS to oxygen singlets ( Boussiba, 2000 ). Although plants exposed to the highest herbicide doses contain high carotenoid concentration, this was not sufficient to prevent oxidative damages to PSII (since we observed lower F V / F M value). These plants also showed higher lipid peroxidation ( Figure ​ Figure3 3 ) indicating oxidative damages ( Gunes et al., 2007 ), as it was also shown in maize ( Sergiev et al., 2006 ) and rice ( Ahsan et al., 2008 ). However, for exposure times longer than 24 h, plants treated with the glyphosate-based herbicide showed reduced carotenoid concentration ( Figure ​ Figure1 1 ). This can be a consequence of the inhibition of the shikimate pathway leading to decreased PQ concentration, since plastoquinone is a co-factor of the phytoene desaturase and ζ-carotene desaturase, enzymes involved in the carotenoid biosynthesis pathway ( Sandmann et al., 2006 ). Therefore, decreased plastoquinone concentration will affect directly carotenoid biosynthesis ( Figure ​ Figure6 6 , #5). In addition, the decrease in the non-photochemical energy dissipation (NPQ) [one of the mechanisms by which plants can dissipate excess light energy absorbed by PSII light-harvesting complexes in order to minimize the generation of the highly reactive 1 O 2 responsible for oxidative damages ( Demming-Adams and Adams, 2000 )], in plants treated with the glyphosate-based herbicide ( Figure ​ Figure2 2 ) can be related to the decreased biosynthesis of carotenoids. β-carotene is known to be the precursor of zeaxanthin, the first compound of xanthophyll cycle ( Bouvier et al., 1996 ), and therefore, reduced carotenoid concentration could lead to a lower efficiency of the xanthophyll cycle, reducing plant capacity for photoprotection and thus, leading to increased PSII damages (as shown by reduced F V / F M ) ( Figure ​ Figure6 6 , #6). These decreases in the photosynthetic activity (shown by the decrease in ETR) and in the NPQ may also have contributed to a higher production of ROS ( Sergiev et al., 2006 ; Ahsan et al., 2008 ; Gomes and Juneau, 2016 ; Gomes et al., 2016a ) due to over-excitation of chlorophylls ( Figure ​ Figure6 6 , #7).

As we found in the present study ( Figure ​ Figure3 3 ; Figure ​ Figure6 6 , #8), increased lipid peroxidation has been previously observed in glyphosate-exposed plants and was related to increased H 2 O 2 content in plants ( Moldes et al., 2008 ; Miteva et al., 2010 ; Gomes et al., 2016b ). Lipid peroxidation resulting from increased levels of ROS (such as H 2 O 2 ) has been shown to affect the integrity of the thylakoid membranes ( Richter, 1992 ), contributing to the noted decrease in ETR ( Figure ​ Figure6 6 , #9). Glyphosate was demonstrated to cause depletion of photosynthetic proteins leading to losses of photosynthetic capacity in plants ( Diaz Vivancos et al., 2011 ). However, it has long been recognized that H 2 O 2 is a potent inhibitor of photosynthesis, since, even at low concentrations, it can inhibit CO 2 fixation by oxidizing the thiol groups of some essential enzymes of the Calvin cycle ( Foyer and Noctor, 2011 ). We can therefore advance that the observed decrease in photosynthesis (ETR) in presence of glyphosate-based herbicide may also be directly linked to higher H 2 O 2 concentration ( Figure ​ Figure5 5 , #8). Carbon assimilation (and therefore photosynthesis) can also be negatively affected by the decreased stomatal conductance ( g s ) ( Figure ​ Figure1 1 ) in presence of herbicides ( Zobiole et al., 2010 ). Reduced g s , as also previously reported in Hordeum vulgare (barley) and Lolium perenne plants exposed to glyphosate ( Olesen and Cedergreen, 2010 ; Yanniccari et al., 2012 ), can limit photochemistry, resulting in decreased ETR ( Figure ​ Figure6 6 , #10). The observed ETR reduction could also be due to the alteration of the integrity of PSII (lower F V / F M ) ( Figure ​ Figure2 2 ; Figure ​ Figure6 6 , #11). In addition, the decrease in total chlorophyll concentration in presence of the glyphosate-based herbicide may be responsible for a lower light interception and thus, the noted lower electron transport rate ( Figure ​ Figure6 6 , #11). Decreased chlorophyll contents when plants are exposed to herbicide application have been demonstrated previously and have been attributed to an increase chlorophyll degradation or to a decrease in chlorophyll synthesis ( Cakmak et al., 2009 ; Mateos-Naranjo et al., 2009 ; Huang et al., 2012 ; Gomes et al., 2016a ).

In order to better understand the processes involved in H 2 O 2 accumulation (and herbicide-induced oxidative damage), we investigated the activity of antioxidant system in treated plants. Increases in proline synthesis is a common protective-response of plants to stress conditions ( Hayat et al., 2012 ). It is important to note, however, that proline can also act as a significant signaling molecule in plant physiological processes, mainly under stress conditions ( Hare and Cress, 1997 ). In the present study, we suggest that the observed proline accumulation in treated plants is associated to oxidative protection, NADP + recovery and shikimate pathway stimulation ( Figure ​ Figure6 6 , #12 and #13). As we also observed, proline biosynthesis is commonly stimulated by increased cellular-ROS concentration conditions ( Soshinkova et al., 2013 ). Although proline can be synthesized from ornithine, metabolic labeling studies indicate that, under stress conditions, proline is mainly produced from glutamate (as reviewed by Hare and Cress, 1997 ). Therefore, the proline accumulation found in our study indicates that this pathway is highly activated ( Figure ​ Figure6 6 , #12). A special function of proline in preventing oxidative damage and enhancing tolerance from abiotic oxidative stress has been proposed recently ( Soshinkova et al., 2013 ) and proline accumulation in plants in response to glyphosate exposure was documented ( Huang et al., 2012 ). Due to the loss of feedback control of the shikimate pathway by tyrosine (that regulates the activity of 3-deoxy- D -aravino-heptulosonate-7-phosphate synthase) ( Crowley, 2006 ), the herbicide (glyphosate) led to an unregulated flux of carbon into the shikimate pathway ( Siehl, 1997 ). As a result, there is an increased demand for erythrose-5-phosphate, the substrate of the first reaction of the shikimate pathway. Erythrose-5-phosphate is produced in the oxidative pentose phosphate pathway (OPPP), which is dependent on NAD(P) + availability and inhibited by NADPH ( Hare and Cress, 1997 ). During proline synthesis, NADPH is oxidized, therefore stimulating OPPP. Even a small change in the NAD(P) + /NADPH ratio may have a large effect on this redox-sensitive pathway ( Hare and Cress, 1997 ). The oxidation of NADPH during proline synthesis, coupled with the reduction of NADP + during the two oxidative steps of the OPPP, promotes a cycle of changes in NAD(P) + /NADPH ratio which stimulates proline biosynthesis, justifying its accumulation during stress ( Hare and Cress, 1997 ). Therefore, upon the glyphosate-based herbicide exposure, the proline accumulation in willow plants could also be linked to the OPPP stimulation for the production of the erythrose-5-phosphate which will be used in shikimate pathway ( Figure ​ Figure6 6 , #13). Supporting this hypothesis, Diaz Vivancos et al. (2011) observed decreased NADP/NADPH ratios in leaves of glyphosate-sensitive soybeans upon glyphosate treatment, as a result of the decreases in NADP + pool. As mentioned previously, under stress conditions, proline is mainly produced from glutamate ( Hare and Cress, 1997 ). Glutamate is also required during δ-aminolevulinic acid (ALA; a chlorophyll precursor) biosynthesis through ALA-synthetase and γ,δ-dioxivalerate cycles ( Beale, 1978 ). Therefore, if glutamate was preferentially used for proline biosynthesis (as suggested by proline greater accumulation in treated plants in relation to control; Figure ​ Figure3 3 ), a decrease in ALA biosynthesis may be obtained, therefore contributing to the decreased chlorophyll concentration observed in treated plants ( Figure ​ Figure6 6 , #14). As suggested by Gomes et al. (2016a) , the decrease in chlorophyll concentration may also be due to its degradation by increased ROS content ( Figure ​ Figure6 6 , #15).

Even though treated plants showed increased activities of antioxidant enzymes after 6 h exposure, they were not able to prevent both peroxide accumulation and lipid peroxidation, indicating a clear deleterious effect of the glyphosate-based herbicide through oxidative burst. Moreover, the strong inhibition of SOD, APX, and GPX activities observed in plants exposed to herbicide after 48 h ( Figure ​ Figure4 4 ) can be related to the increased H 2 O 2 and decreased ETR also observed in these plants. SOD is the first defense enzyme against oxidative stress ( Pompeu et al., 2008 ) and is closely related to stress resistance in plants ( Song et al., 2006 ). Indeed, this enzyme was involved in the PSII protection against the effects of prooxidant herbicides, limiting carbon dioxide and photoinhibitory conditions ( Foyer et al., 1994 ; Arisi et al., 1998 ). The observed decrease in SOD activity ( Figure ​ Figure4 4 ) can therefore contribute to the herbicide-deleterious effects on photosynthesis in willow plants. We also demonstrated the key role of APX and GPX to prevent H 2 O 2 accumulation in willow plants since: (1) decreased activities of both enzymes were related to increased H 2 O 2 concentration in leaves; (2) even if treated plants shown higher CAT activity, it was not able to prevent H 2 O 2 accumulation. The importance of APX and GR in avoiding oxidative stress has also been observed in metal(loid) treated plants ( Chaoui et al., 1997 ; Gomes et al., 2013a , b ) and the inactivation/degeneration of these enzymes has been related to increased H 2 O 2 concentrations and oxidative damages to plants ( Gomes et al., 2013b ). When H 2 O 2 accumulation exceeded the tolerance limit of plants, enzymatic systems are prone to protein carbonylation–an irreversible oxidative process in which the side chains of Lys, Arg, Pro, and Thr are converted to aldehyde or keto groups ( Sohal et al., 2002 ), which may have been occurring in willow plants exposed to the studied herbicide ( Figure ​ Figure6 6 , #16).

We also observed an interesting response of GR activity at 48 and 72 h, since its activity was not significantly decreased by the glyphosate-based herbicide exposure. GR is linked to APX and GPX activity by the glutathione-ascorbate cycle ( Foyer and Noctor, 2011 ). However, as mentioned, the GR activity did not follow APX and GPX patterns upon the herbicide exposure. The maintenance of GR activity in treated plants indicates that APX and GPX activities were not limited by substrate availability, reinforcing that the proposed oxidative damage (protein carbonylation) of the enzymes could be responsible for their degeneration. We may hypothesize that, similarly to the proline production, the higher NADP(H)-dependent-GR activity can favor OPPP and contribute as a source of NADP + for photochemistry.

In addition to being the substrate for APX, ascorbate is an important antioxidant component of the cellular redox potential and its activity is linked to ascorbate-glutathione metabolic cycle ( Foyer and Noctor, 2011 ). In the present study, we found a link between the reduced form of ascorbate (AsA) and the APX activity. Indeed, up to 24 h, treated plants showed higher APX activity concomitantly to the reduced AsA concentration in their leaves; similarly, decreased APX activity for the following treatment periods was related to the increased AsA content. On the other hand, the contrary was observed for the oxidized form of ascorbate (DHA). The accumulation of AsA, as noted by the increased AsA/DHA ratio in treated plants, shows that the DHA has been effectively recycled to AsA by ascorbate-glutathione cycle. We also observed that total ascorbate concentrations (AsA + DHA) were reduced in herbicide treated plants ( Figure ​ Figure5 5 ). It is known that ascorbate concentration and ETR are closely linked, as the light-dependent stimulation of ascorbate biosynthesis requires photosynthetic electron transport activity ( Yabuta et al., 2007 ). Thus, reduced ETR in treated plants could explain the observed reduction in ascorbate pool ( Figure ​ Figure6 6 , #17). Low ascorbate pool favors the increase in both ROS ( Figure ​ Figure6 6 , #18) and abscisic acid (ABA), leading to an increase in signal transduction through ROS-mediated and ABA-dependent signaling cascades ( Foyer and Noctor, 2011 ). Among others, the interactive effect of ROS and ABA in stomatal movement is well studied, with increased ROS and ABA content inducing stomatal closure ( Gomes et al., 2014a ). This mechanism can also be related to the observed herbicide-induced decreases in g s ( Figure ​ Figure6 6 , #19).

As expected, the primary target site of the studied glyphosate-based-herbicide (Factor ® 540) on willow plants is the shikimate pathway. We demonstrated, for the first time, that on top of the alteration of this primary target site, this herbicide induces a series of interconnected events that leads to decreased photosynthetic activity in willow plants. Furthermore, we showed that the herbicide-deleterious effects on photosynthesis are strongly related to herbicide-induced oxidative stress, and that reduction of photosynthesis may amplify the observed effect by inducing ROS production. Our results evidenced that as for photosynthesis-target herbicides, which trigger ROS production and oxidative stress, glyphosate herbicidal effect may be related to induction of ROS accumulation. The inhibition of shikimate pathway may induce changes in redox status with important consequences in leaf metabolism, mainly on photosynthesis. Glyphosate tolerance in plants, for instance, have been related to the ability of plants to deal with ROS accumulation through the activation of antioxidant systems ( Maroli et al., 2015 ). However, since photosynthetic processes of GR plants have been shown to be affected by glyphosate-based herbicides ( Zobiole et al., 2010 , 2011 , 2012 ), glyphosate may target other cellular sites, inducing ROS formation, for example, mitochondrial electron chain, as proposed by Gomes and Juneau (2016) . Although ROS formation may also be produced in the mitochondria, our model fits with several results presented in the literature about the effects of glyphosate in sensitive plants, highlighting the role of ROS induction in this herbicidal mechanism of action.

Author Contributions

MG, SL performed the experiments; MG, MLa, and PJ designed the experiments; MG and PJ wrote the paper; LH-E and MLu gave technical support and conceptual advice.

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

We thank Jerzy Kruk from the Department of Plant Physiology and Biochemistry, Jagiellonian University for providing us the plastoquinone standard.

Abbreviations

Funding. This research was supported by the Natural Science and Engineering Research Council of Canada (NSERC) through a Strategic grant awarded to MLa, PJ, and MLu. MG received a Ph.D. scholarship from Fonds de Recherche du Québec–Nature et Technologies (FRQNT) and LH-E received a Ph.D. scholarship from the Natural Science and Engineering Research Council of Canada (NSERC).

Supplementary Material

The Supplementary Material for this article can be found online at: http://journal.frontiersin.org/article/10.3389/fpls.2017.00207/full#supplementary-material

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Herbicide Effects on Photosynthesis

Herbicide Effects on Photosynthesis

How Are Cellular Respiration & Photosynthesis Almost Opposite ...

Weeds can reduce crop yield by competing for resources with crop plants. Reducing weeds on a large scale is best accomplished through the use of herbicides. Herbicides are a group of pesticides that control or eliminate the growth of weeds. Herbicides have many different modes of action, one of which is inhibiting photosynthesis -- a process vital to plant survival.

Types of Herbicides

Herbicides work to reduce weed pests through many mechanisms, but all serve to disrupt weed growth. Growth regulators diminish cell division and enlargement and are used largely to kill broadleaf weeds (i.e., primarily to protect grass crops such as corn), usually through the use of hormones. Pigment inhibitors break down chlorophyll (the pigment that gives plants their green color), which is necessary for photosynthesis to occur. Seedling growth inhibitors work by inhibiting plant growth just after germination, interfering with the growth of either roots or shoots (i.e., leaves). Other herbicides work by inhibiting the production of materials necessary for plant growth (e.g., amino acids or lipids).

What Is Photosynthesis?

Photosynthesis is the process by which plants use sunlight, carbon dioxide and water to make sugars (for enzymes and growth) and oxygen. The photosynthetic process is vital to life on earth, because it creates the oxygen animals, including humans, need for respiration. Biochemically speaking, photosynthesis is a fairly complicated process that takes place within plant cells and requires many enzymes and the transfer of electrons. If any of these photosynthetic systems is disrupted for any reason, the process will shut down and the plant will die. Photosynthesis, therefore, is the target of a group of herbicides known as photosynthetic inhibitors.

How Photosynthesis Inhibitors Work

Photosynthesis is driven largely by the transfer of electrons from chlorophyll molecules into the surrounding cytochromes. These electrons are passed along a series of cytochromes in what is known as an electron transport system. Sunlight activates these electrons where they are passed along another electron transport chain and are eventually used in a carbon-fixing reaction. Photosynthesis-inhibiting herbicides work by blocking the transfer of electrons. Without electron transfer, energy from the sun cannot be transformed into energy that is usable by plants to generate new tissue and sustain life.

Photosynthetic Inhibitor Uses

Inhibitors of photosynthesis are used mainly to control broad-leaved weed pests. That is, grass crops such as corn benefit the most from photosynthetic inhibitors.

Symptoms of Photosynthesis Inhibitors

Plants that have been exposed to photosynthesis-inhibiting herbicides will begin to appear yellow on the veins and around the edges of the oldest leaves, which will be followed by similar damage to younger leaves. Yellow spots may also appear on affected leaves.

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  • Arizona State University: Photosynthesis
  • Earthguide: Photosynthesis Diagram

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How Are Cellular Respiration & Photosynthesis Almost Opposite Processes?

When to List

Ways to measure, conceptual diagrams, literature reviews, other stressors that may influence herbicide effects.

Herbicides are chemicals used to manipulate or control undesirable vegetation. Herbicide application occurs most frequently in row-crop farming, where they are applied before or during planting to maximize crop productivity by minimizing other vegetation. They also may be applied to crops in the fall, to improve harvesting.

Agricultural use of herbicides in 2001 in millions of acres.

Herbicides are used in forest management to prepare logged areas for replanting. The total applied volume and area covered is greater but the frequency of application is much less than for farming (Shepard et al. 2004).

In suburban and urban areas, herbicides are applied to lawns, parks, golf courses and other areas. Herbicides are applied to water bodies to control aquatic weeds. These weeds can impede irrigation withdrawals or interfere with recreational and industrial uses of water (Folmar et al. 1979).

The potential effects of herbicides are strongly influenced by their toxic mode of action and their method of application. The molecular site of action is challenging to predict because structural associations have not been identified (Duke 1990), but modes of action are well-established.

Herbicides can act by inhibiting cell division, photosynthesis or amino acid production or by mimicking natural plant growth hormones, causing deformities (Ross and Childs 1996). Application methods include spraying onto foliage, applying to soils and applying directly to aquatic systems.

Figure 1 and Table 1 present the ten herbicides most used on agricultural land in the U.S. Glyphosate and atrazine were applied to more than double the crop field acreage than the third leading herbicide, 2,4-D, in 2001.

Herbicides may cause biological impairments of water bodies if they occur in water or sediment at sufficient concentrations. Most commonly, they enter surface water in runoff or leachate, but, because they have relatively low toxicity to fish and invertebrates (see Table 2). Acute toxicity is likely only when they are deliberately or accidentally applied directly to water bodies.

Direct applications may result in direct toxicity to non-target plants and animals or indirect effects due to the death and decomposition of plants. Impairments also are more likely when herbicides are applied together or with other pesticides (Streibig et. al. 1998), resulting in additive or synergistic effects.

Atrazine reacts synergistically with chlorpyrifos: the mixture was seven times more toxic to an earthworm species than the two individual pesticides (Lydy and Linck 2003). Atrazine also increased the effects of other pesticides in mosquito larvae and various flies (Belden and Lydy 2000, Lydy and Linck 2003). The surfactants used in herbicide solutions also can be toxic to biota and are not considered when testing active ingredients (Folmar et al. 1979).

Checklist of Sources, Site Evidence and Biological Effects

Herbicides are addressed in this module as proximate stressors. Herbicides should be a candidate cause when human sources and activities, site observations or observed effects support portions of the causal pathways (see Figure 2). The conceptual diagram and other information also may be useful in Step 3: Evaluate Data from the Case.

Rather than causing direct toxicity to organisms, herbicides may contribute to other stressors (e.g., instream habitat alteration via riparian devegetation). In such cases, herbicides can be considered as part of the pathway for the proximate cause of impairment.

The checklist below will help you identify key data and information useful for determining whether to include herbicides among your candidate causes. This list is intended to guide you in collecting evidence to support, weaken or eliminate herbicides as a candidate cause. For more information on specific entries, go to the When to List tab.

 simple conceptual diagram, depicting pathways from sources to impairments, related to herbicides.

Sources and Activities

  • Forest management
  • Agriculture/crop cultivation
  • Golf courses
  • Roads/rights of way
  • Aquatic weed control

Site Evidence

  • Dead or injured plants
  • Irrigation channels
  • Drainage from fields or lawns

Biological Effects

  • Inhibition of phytoplankton, periphyton or macrophytes
  • Reduced invertebrate species richness and abundance
  • Reduction of sensitive species and abundance of tolerant species
  • Temperature : Elevated temperature frequently tends to increase the toxicity of chemicals.
  • Moderate to high pH : pH will determine the ionic state and bioavailability of ionizable herbicides.
  • Dissolved oxygen (DO) : Signs of herbicide application are evidence of a causal pathway to low DO when DO is a candidate cause of a kill or other impairment.
  • Unspecified toxics : Surfactants in herbicide formulations can be more toxic to animals than the active ingredients (Folmar et al. 1979, Diamond and Durkin 1997). The toxicity of Roundup®, which increased with elevated pH, was attributed to the surfactant, rather than the active ingredient, glyphosate (Folmar et al. 1979). This result is reinforced by findings that more alkaline pH decreases glyphosate toxicity but increases surfactant toxicity (Diamond and Durkin 1997).

You also may wish to consider other causes with similar evidence:

  • Insecticides
  • Endocrine disruptors
  • Toxic metals

Sources and Activities that Suggest Listing Herbicides as a Candidate Cause

Site evidence that suggests listing herbicides as a candidate cause, biological effects that suggest listing herbicides as a candidate cause, site evidence that supports excluding herbicides as a candidate cause.

Forestry management practices, agricultural operations, and urban development and maintenance are all sources of herbicides that may enter surface waters and cause impairments. Herbicides are applied to forests after harvesting to suppress brush and noncommercial trees. For that use, the rate of application may be high and exposed streams are more likely to be of higher quality than agricultural or urban streams. Conversely, agricultural operations may contribute large quantities of herbicides because they may apply herbicides multiple times per year and they may be applied by planes, addition to irrigation water or spraying onto crops (see Figure 3).

Photo of farm equipment dispensing chemicals to protect crops.

Urban land uses can contribute as homeowners and managers of parks, golf courses and other lawns use herbicides for aesthetic enhancement. Herbicides also are used on rights of way for roads, pipelines, railroads and electrical transmission lines and for control of plants in cracks in pavements. Such urban and suburban uses are likely to contaminate storm waters.

Herbicides also are directly applied to waters to control vegetation in ponds, ditches, irrigation canals and recreational waters. Such applications are sources of exposure at the point of application and downstream.

Photo of a farm showing how runoff drains into ditches and streams.

Although herbicides in general have lower toxicity to animals than other pesticides, fish or invertebrate kills may be a sign of herbicide use. For example, acrolein has been applied to irrigation ditches at levels sufficient to be acutely lethal to fish and invertebrates (see acrolein in U.S. EPA 2009), and if not properly applied to fields it can cause kills in receiving waters. Kills also may be due to low dissolved oxygen (DO) concentrations resulting from plant materials decomposing in water.

Because herbicides tend to affect plants more quickly and severely than animals, the most useful biological sign of herbicides is effects on aquatic plants (Kreutzweiser et al. 1995, Van den Brink et al. 1997, Hall et al. 1997). This trait may help distinguish the biological effects of herbicides from those of insecticides and most other toxic chemicals. Secondary effects of herbicides are mediated by low DO concentrations from plant decomposition and changes in trophic structure due to plant community changes.

Herbicides may reduce taxa richness and abundance of fish and benthic macroinvertebrates due to reductions of sensitive species and increased abundance of tolerant species at high concentrations (Daam and Van den Brink 2007, Dewey 1986). It also has been contended that some herbicides, particularly atrazine, have specific mechanisms of action in aquatic frogs and fish, including developmental abnormalities (Hayes et al. 2006, Tillit et al. 2010). However, a review by the U.S. EPA found that evidence for such effects in amphibians was weak and inconsistent (U.S. EPA 2007).

Photo showing a dense collection of algae or underwater vegetation.

Herbicides and their metabolites can be measured in groundwater and surface water by gas chromatography (GC), mass spectrometry (MS), high performance liquid chromatography with diode-array detection (HPLC/DAD), liquid chromatography (LC), solid-phase extraction (SPE) or enzyme-linked immunosorbent assay (ELISA) (Scribner et al. 2000). Because there isn't a standard method for detecting all herbicides, measurements can be difficult, expensive and time-consuming.

Herbicide metabolites can have toxicity similar to that of the parent herbicide and are often found in higher concentrations (USGS 2010). Presently metabolites of triazines, chloroacetanilides, phenyl ureas and the phosphanoglycine glyphosate have been measured (Scribner et al. 2000, USGS 2010). Different herbicides and metabolites are measurable using different techniques, and the proper technique must be matched with the metabolite of interest. The USGS Toxic Substances Hydrology Program provides guidance, lab methods, field methods and literature related to detecting herbicides in ground and surface water.

  • USGS Toxic Substances Hydrology Program

About Conceptual Diagrams

Simple conceptual model diagram, detailed conceptual model diagram.

Conceptual diagrams are used to describe hypothesized relationships among sources, stressors and biotic responses within aquatic systems.

  • More about Conceptual Diagrams

Herbicides are used to control undesired plants on farms, in commercial forests, and on lawns and managed landscapes. Herbicides are sometimes applied directly to surface water for aquatic weed control. Typically herbicides are applied to soil or terrestrial vegetation, which can increase herbicides in groundwater discharge, atmospheric drift and runoff. The extent to which herbicides reach streams depends on factors such as precipitation, application timing and rates and environmental persistence of herbicides and their metabolites.

  • Simple and Detailed Conceptual Model Diagrams
  • Simple Conceptual Diagram (PPT) (ppt) (138.5 KB)
  • Detailed Conceptual Diagram (PPT) (ppt) (185 KB)

In streams, herbicides may be dissolved in the water column or bound to sediments, and their impact depends on the medium in which they occur. Exposures may be episodic (e.g., occurring during runoff events) or continuous (e.g., exposure to herbicide contaminated bed sediments). The bioavailability, uptake and toxicity of herbicides vary with environmental conditions (e.g., pH).

Increased herbicides in streams can adversely affect stream flora and fauna via several mechanisms, including reduced growth, condition, and reproduction; increased mortality; and changes in behavior. These effects can result in biologically impaired macrophyte, periphyton, phytoplankton, fish and invertebrate assemblages, which in turn can contribute to changes in community structure and ecosystem function.

High concentrations of herbicides and their metabolites in streams can have lethal and sub-lethal effects on aquatic biota, potentially changing community structure and ecosystem function. This conceptual diagram (Figure 7) illustrates linkages between human activities and sources (top of diagram), herbicide-related stressors (middle of diagram), and the biological responses that can result (bottom of diagram).

Snapshot of a detailed conceptual diagram related to herbicides.

In some cases, additional steps leading from sources to stressors, modes of action leading from stressors to responses, and other modifying factors are shown. This narrative generally follows the diagram top to bottom, left to right.

Linking Sources to Stressors

Anthropogenic activities and land uses, such as industry, urban development, forestry and agriculture can contribute herbicides to streams. Herbicide manufacturers, industrial facilities and wastewater treatment plants may discharge effluents containing herbicides. Accidental or unpermitted discharges also may occur.

Herbicides are sometimes applied directly to surface water for aquatic weed control (e.g., for water-based recreation). Herbicides may be applied to golf courses, lawns and other managed landscapes, forests, crop fields and orchards to control a variety of unwanted vegetation. In some cases, herbicides may be transported atmospherically in spray drift. These applied herbicides may enter streams via stormwater runoff, groundwater discharges or direct atmospheric deposition.

Stored herbicides, both at sites where they are used and at sites where they are manufactured, also may be transported to streams via runoff or groundwater transport. The extent to which these transport pathways occur depends upon several factors, including land cover, precipitation patterns, timing and rates of application and environmental persistence of the herbicides.

Linking Stressors to Biological Responses

In streams, herbicides can be dissolved in the water column or bound to sediments, and the effects they have will depend upon the medium in which they occur. Exposures may be episodic (e.g., pulsed deliveries with stormwater runoff) or continuous (e.g., long-term exposure to herbicide-contaminated sediments). The bioavailability, uptake, and toxicity of herbicides and their metabolites during these exposures depends on factors such as temperature, pH, and dissolved oxygen concentrations.

The most direct effects of herbicide pollution are decreased condition, growth, and reproduction, and increased mortality, of plants (i.e., macrophytes, periphyton and phytoplankton). For example, exposure to herbicides may lead to elevated internal herbicide concentrations and decreased photosynthesis, cell division, and amino acid production in plants. Effects on aquatic plants can indirectly affect fish and invertebrates by modifying habitat and food availability.

Exposure to herbicides also can directly increase mortality and change the behavior and reproduction of fish, amphibians and invertebrates. Possible changes in behavior include increased invertebrate drift and increased avoidance by fish.

Ultimately, these effects may result in changes in community structure (e.g., decreased richness, changes in functional feeding groups) and ecosystem function. For example, aquatic vegetation is especially susceptible to herbicides, so may decrease in abundance and richness. As a result, the relative abundances of invertebrate feeding groups may shift. However, herbicide-resistant and other non-target plants may increase in abundance with herbicide exposure, due to reduced competitive pressure from affected plants.

This section presents an annotated bibliography of references providing information on stressor-response relationships for herbicides, as well as general background on herbicide properties. This is not meant to be a comprehensive bibliography of references dealing with herbicides, but rather is meant to highlight a few references that may be especially useful.

  • Kegley SE, Hill BR, Orme S, Choi AH (2010) Pesticide Action Network Pesticide Database . Pesticide Action Network, North America, San Francisco CA.

This database has toxicity data for pesticides across many species. It provides a good starting point for finding pesticide use, occurrence, and effects data on the web.

  • Ross MA, Childs DJ (1996) Herbicide Mode-of-Action Summary . Purdue University, Department of Botany: Plant Pathology, West Lafayette IN. Report No. WS-23-W.

This publication provides a breakdown of seventy-eight common herbicides organized by translocation mechanism and then mode of action. It further subdivides the information into chemical type and then common and trade names. A brief paragraph describes each mode of action and types of vegetation that the herbicide is often used to control.

  • Stenersen J (2009) Chemical Pesticides: Mode of Action and Toxicology. CRC Press, Boca Raton FL.

This is a recent reference for mechanistic health and environmental toxicity information for pesticides, including herbicides and insecticides.

  • U.S. EPA (2009) Aquatic Life Benchmarks for Pesticide Registration . U.S. Environmental Protection Agency, Office of Pesticide Programs, Washington DC.

The aquatic life benchmarks (for freshwater species) provided in this module are based on toxicity values reviewed by U.S. EPA and used in the Agency's most recent risk assessments, developed as part of the decision-making process for pesticide (including herbicides) registration. Acute and chronic benchmarks are provided for fish, invertebrates and aquatic plants. The table of benchmarks provides links to supporting ecological risk assessments. Each aquatic life benchmark is based on the most sensitive, scientifically acceptable toxicity endpoint available to U.S. EPA for a given taxon. U.S. EPA's goal is to add to these benchmarks annually.

  • Belden J, Lydy MJ (2000) Impact of atrazine on organophosphate insecticide toxicity. Environmental Toxicology and Chemistry 19:2266-2274.
  • Daam MA, Van den Brink PJ (2007) Effects of Chlopyrifos, Carbendazim, and Linuron on the ecology of a small indoor aquatic microcosm. Archives of Environmental Contamination and Toxicology 53(1):22-35.
  • Dewey SL (1986) Effects of the herbicide atrazine on aquatic insect community structure and emergence. Ecology 67(1):148-162.
  • Diamond GL, Durkin PR (1997) Effects of Surfactants on the Toxicity of Glyphosate, with Specific Reference to RODEO. U.S. Department of Agriculture, Animal and Plant Health Inspection Service, Riverdale MD. SERA TR 97-206-1b.
  • Duke SO (1990) Overview of herbicide mechanisms of action. Environmental Health Perspectives 87:263-271.
  • Folmar LC, Sanders HO, Julin AM (1979) Toxicity of the herbicide glyphosate and several of its formulations to fish and aquatic invertebrates. Archives of Environmental Contamination and Toxicology 8:269-278.
  • Hall LW Jr, Anderson RD, Ailstock MS (1997) Chronic toxicity of atrazine to sago pondweed at a range of salinities: implications for criteria development and ecological risk. Archives of Environmental Contamination and Toxicology 33:261-267.
  • Hayes TB, Stuart AA, Mendoza M, Collins A, Noriega N, Vonk A, Johnston G, Liu R, Kpodzo D (2006) Characterization of atrazine-induced gonadal malformations in African clawed frogs (Xenopus laevis) and comparisons with effects of an androgen antagonist (cyproterone acetate) and exogenous estrogen (17B-estradiol): support for the demasculinization/feminization hypothesis. Environmental Health Perspectives 114(Supplement 1):134-141.
  • Kegley SE, Hill BR, Orme S, Choi AH (2010) Pesticide Action Network Pesticide Database .  Pesticide Action Network, North America, San Francisco CA.
  • Kreutzweiser DP, Capell SS, Sousa BC (1995) Hexazinone effects on stream periphyton and invertebrate communities. Environmental Toxicology and Chemistry 14(9):1521-1527.
  • Larson DL, McDonald S, Fivizzani AJ, Newton WE, Hamilton SJ (1998) Effects of the herbicide atrazine on Ambystoma tigrinum metamorphosis: duration, larval growth, and hormonal response. Physiological Zoology 71(6):671-679.
  • Lydy MJ, Linck SL (2003) Assessing the impact of triazine herbicides on organophosphate insecticide toxicity to the earthworm Eisenia fetida . Archives of Environmental Contamination and Toxicology 45:343-349.
  • Ross MA, Childs DJ (1996) Herbicide Mode-of-Action Summary .  Purdue University, Department of Botany: Plant Pathology, West Lafayette IN. Report No. WS-23-W.
  • Scribner EA, Thurman EM, Zimmerman LR (2000) Analysis of selected herbicide metabolites in surface and ground water of the United States. Science of the Total Environment 248(2-3):157-167.
  • Shepard JP, Creighton J, Duzan H (2004) Forestry herbicides in the United States: an overview. Wildlife Society Bulletin 32(4):1020-1027.
  • Streibig JC, Kudsk P, Jensen JE (1998) A general joint action model for herbicide mixtures. Pesticide Science 53(1):21-28.
  • Tate TM, Spurlock JO, Christian FA (1997) Effect of glyphosate on the development of Pseudosuccinea columella snails. Archives of Environmental Contamination and Toxicology 33:286-297.
  • Tillit DE, Papoulias DM, Whyte JJ, Richter CA (2010) Atrazine reduces reproduction in fathead minnow ( Pimephales promelas ). Aquatic Toxicology 99(2):149-159.
  • U.S. EPA (2003) Ambient Aquatic Life Water Quality Criteria for Atrazine: Revised Draft . Office of Water, Office of Science and Technology, Health and Ecological Criteria Division, Washington DC. EPA-822-R-03-023.
  • U.S. EPA (2007) White Paper on the Potential for Atrazine to Affect Amphibian Gonadal Development. U.S. Environmental Protection Agency, Office of Pesticide Programs, Washington DC.
  • U.S. EPA (2009) Aquatic Life Benchmarks for Pesticide Registration . U.S. Environmental Protection Agency, Office of Pesticide Programs, Washington DC.
  • U.S. EPA (2009) Ambient Aquatic Life Water Quality Criteria for Acrolein . Office of Water, Office of Science and Technology, Health and Ecological Criteria Division, Washington DC. CAS Registry No. 107-02-8. EPA/822/R-09/010
  • USGS (2010) Glyphosate herbicide found in many midwestern streams, antibiotics not common. U.S. Geological Survey.
  • Van den Brink PJ, Crum SJH, Glystra R, Bransen F, Cuppen JGM, Brock TCM (2009) Effects of a herbicide-insecticide mixture in freshwater microcosms: risk assessment and ecological effect chain. Environmental Pollution 157:237-249.
  • Van den Brink PJ, Hartgers EM, Fettweis U, Crum SJH, Van Donk E, Brock TCM (1997) Sensitivity of macrophyte-dominated freshwater microcosms to chronic levels of the herbicide Linuron. Ecotoxicology and Environmental Safety 38:13-24.

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  • Photosynthesis inhibitor herbicides
  • Crop production
  • Weed management
  • Herbicide mode of action and sugarbeet injury symptoms

The photosynthesis inhibitors include these herbicide families:

  • Benzothiadiazole
  • Phenyl-carbamate

Photosynthesis inhibitors disrupt the photosynthetic (food producing) process in susceptible plants by binding to specific sites within the photosystem II complex in plant chloroplasts. Inhibition of photosynthesis could result in slow starvation of the plant; however, in many situations, rapid death occurs perhaps from the production of secondary toxic substances.

Injury symptoms include interveinal yellowing (chlorosis) and death (necrosis) of leaf tissue beginning at the leaf margins and progressing toward the center of leaves.

The triazines, triazinones, phenylureas and uracils are taken up into the plant via the roots or foliage and move in the xylem to plant leaves. As a result, injury symptoms first will appear on the older leaves, along the leaf margin.

They also have relatively long persistence in soil.

The benzothiadiazoles and phenyl-carbamates are foliar-applied photosynthetic inhibitors and generally remain in the foliar portions of the treated plant. They are contact herbicides requiring thorough spray coverage of the foliage for good weed control. Movement from the foliage to roots is negligible.

Herbicide use

Atrazine for corn.

Injury symptoms

Residual of photosynthesis inhibitors in soil does not prevent seedlings from germinating or emerging. Injury symptoms occur after emergence, and the speed of appearance of symptoms will be more rapid with sunny days than with cloudy weather.

Symptoms will also be more severe and more rapid as the level of herbicide in the soil increases.

Sugarbeet plants may be in the two- to four-leaf stage before symptoms become noticeable, but plants can die in the early two-leaf stage. Initial symptoms include browning of the cotyledonary leaves and yellowing of the true leaf margins. Browning of leaves will increase with time and total desiccation may result. Older and larger leaves are affected before younger leaves (Photo 39).

Triazines residues are most likely to occur following years with low rainfall because chemical and microbial activity needed to degrade herbicides are limited in dry soil. There are other herbicide tank mixtures that can cause similar visual damage (Photo 40).

Postemergence triazine herbicides cause an initial yellowing followed by desiccation and leaf browning.

Site of action

D1 quinone-binding protein (photosystem II) of the photosynthetic electron transport chain, binding site A.

Atrazine injury on sugarbeet - chlorosis

Metribuzin (Sencor, Dimetric) for alfalfa, corn, field pea, lentil, soybean and potato.

Same as for the triazines; see previous section.

  • Linuron (Lorox) for corn, potato and soybean.
  • Diuron (Diuron) for alfalfa.
  • Tebuthiuron (Spike) for pastures and rangeland.

Same as for the triazines.

Terbacil (Sinbar) for alfalfa.

Bentazon (Basagran) for corn, dry bean, field pea and soybean.

Leaves become chlorotic and later turn brown and die. The older leaves die first. All older leaves can turn brown while the growing point remains green (Photo 41).

Sugarbeet can recover, produce new leaves and produce a nearly normal-size root at harvest if the growing point survives (Photo 42).

D1 quinone-binding protein (photosystem II) of the photosynthetic electron transport chain, binding site B.

Bentazon injury on sugarbeet-necrotic leaves

Bromoxynil (Buctril) for alfalfa, barley, corn, flax, oat and wheat.

Leaves become chlorotic and later turn brown and die. Contact with isolated spray droplets may cause a spotting or speckling of the leaves. The older sugarbeet leaves will be affected more than the young leaves (Photo 43). Sugarbeet can produce new leaves and a harvestable root if the growing point survives.

Desmediphan+phenmedipham (Betamix) for sugarbeet.

Betamix is registered for sugarbeet but injury sometimes occurs, most often in a hot and moist environment. Symptoms from Betamix are very similar to symptoms from bentazon and bromoxynil.

Injured leaves may turn brown and die (Photo 44). The older leaves die first, and the growing point may remain green and alive, even when most leaves are dead (Photo 45).

Sugarbeet plants with a surviving growing point will produce new leaves and a nearly normal-size root at harvest.

Betamix injury on sugarbeet - necrotic leaves

CAUTION: Mention of a pesticide or use of a pesticide label is for educational purposes only. Always follow the pesticide label directions attached to the pesticide container you are using. Be sure that the area you wish to treat is listed on the label of the pesticide you intend to use. Remember, the label is the law.

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IMAGES

  1. Photosynthesis Leaf Disk lab 2020-2021

    analysis of effect of herbicide on photosynthesis lab report

  2. Lab Report 1 Biology Photosynthesis

    analysis of effect of herbicide on photosynthesis lab report

  3. (PDF) Mechanism of Action of the Herbicide 4,6-Dinitro-o-cresol in

    analysis of effect of herbicide on photosynthesis lab report

  4. Lab Report on Photosynthesis

    analysis of effect of herbicide on photosynthesis lab report

  5. Schematic showing the relationship between herbicides with modes of

    analysis of effect of herbicide on photosynthesis lab report

  6. 💣 Photosynthesis lab report introduction. Photosynthesis lab webapi.bu

    analysis of effect of herbicide on photosynthesis lab report

VIDEO

  1. Photosynthesis Lab Procedure

  2. Biology Lab || Photosynthesis

  3. AP Bio Photosynthesis Lab- How to set up and analyze the floating leaf disc lab

  4. Lab 7- Photosynthesis: Floating Spinach Disks Lab

  5. Light is essential for Photosynthesis Practical Experiment

  6. Dehydrogenase Activity in Extracts of Chloroplasts (the light dependent reaction of photosynthesis)

COMMENTS

  1. BIOA01 Lab 3 Photosynthesis F2022

    LAB 3 Photosynthesis and Data Collection for the Formal Lab Report Overview. The following experiment will acquaint you with some of the principles of photosynthesis. You will set up an experiment to examine the effect of DCMU on the photosynthetic rate of an aquatic plant, Elodea densa.

  2. Herbicide Lab Report

    Lab report background and hypothesis in this experiment spinach leaves were used to look at the specific mechanism of photosynthesis, specifically regarding how ... To see if the herbicide stops photosynthesis before or after DCPIP takes proton. 0min: 0. 2min: 0. 4min: 0. 6min: 0. 8min: 0. 10min: 0. 12min: 0. 14min: 0. 16min: 0.

  3. Hill Reaction Lab Report

    The Effects of Herbicide and Access to Light on The Rate of Photosynthesis April 9, 2021 Hannah Berger Bio 153/ Introduction Plants are a vital source of energy for all living things. They are able to produce oxygen in the presence of sunlight and also produce a chemical compound that works as an electron acceptor (Energy efficiency of ...

  4. Department of Biochemistry & Biophysics Texas A&M University

    Visualization of Effect of Photosynthetic Herbicides on the Light Reactions of Photosynthesis: A Teaching Tool E A FUNKHOUSER and T D SHERMAN Department of Biochemistry & Biophysics Texas A&M University College Station, TX 77843-2128, USA Introduction The study of photosynthesis is central to beginning and advanced

  5. Using Herbicides to Understand the Light-Dependent Reactions of

    within the reading. This lab is a two-week lab. The first week you will be introduced to photosynthesis and methods we can use to observe photosynthesis. Your lab group will test the ability of spinach leaves to perform photosynthesis when exposed to light. You will also observe the effect of photosynthetic plants on the pH of an aqueous ...

  6. Glyphosate-Dependent Inhibition of Photosynthesis in Willow

    We studied the physiological mechanisms involved in the deleterious effects of a glyphosate-based herbicide (Factor ® 540) on photosynthesis and related physiological processes of willow (Salix miyabeana cultivar SX64) plants. Sixty-day-old plants grown under greenhouse conditions were sprayed with different rates (0, 1.4, 2.1, and 2.8 kg a.e ha-1) of the commercial glyphosate formulated salt ...

  7. Effects of the Herbicide Imazethapyr on Photosynthesis in PGR5- and NDH

    Photosynthesis is a very important metabolic pathway for plant growth and crop yield. This report investigated the effect of the herbicide imazethapyr on photosynthesis in the Arabidopsis thaliana pnsB3 mutant (a defect in the NDH pathway) and pgr5 mutant (a defect in the PGR5 pathway) to determine which cyclic electron transport chain (CET) of the NDH and PGR5 pathways is more important for ...

  8. Effects of Photosystem‐II‐Interfering Herbicides Atrazine ...

    H erbicides control weeds by inhibiting a variety of metabolic systems and are categorized based on their mode of action. A significant portion of commercial herbicides target the inhibition of essential plant-specific processes, such as photosynthesis, to minimize possible harmful effects on humans and the environment (Ashton and Crafts, 1973).For example, several herbicides specifically ...

  9. Herbicides in Photosynthesis Research

    Photosynthesis and herbicide research have a long common history. Soon after the introduction of a new group of highly effective herbicides in 1956, the substituted aryl ureas, their mode of action was recognized to be the inhibition of light‐driven electron flow and photosynthetic oxygen evolution. ... Different sensitivities of photosystem ...

  10. Herbicide Effects on Photosynthesis

    Herbicide Effects on Photosynthesis. Weeds can reduce crop yield by competing for resources with crop plants. Reducing weeds on a large scale is best accomplished through the use of herbicides. Herbicides are a group of pesticides that control or eliminate the growth of weeds. Herbicides have many different modes of action, one of which is ...

  11. PDF Binding Properties of Photosynthetic Herbicides with the QB Site of the

    In this work, we investigated the inhibitory effect on plant PSII of five commer-cial and widely used photosynthetic herbicides belonging to different chemical classes, namely diuron (i.e., DCMU (urea), the most commonly used inhibitor in photosynthesis research), metobromuron (urea), bentazon (benzothiadiazinone), terbuthylazine (triazine)

  12. Photosynthesis Alterations in Wheat Plants Induced by Herbicide, Soil

    The wheat plants were pretreated with the selective herbicide Serrate® (Syngenta) and subsequently subjected to drought or flooding stress for 7 days. The gas exchange parameters, chlorophyll a fluorescence and leaf pigment content were measured. The measurements were performed during the stress period and after 4 days of plants recovery. Herbicide pretreatment did not cause significant ...

  13. Photosynthesis Lab

    Photosynthesis Lab. Photosynthesis is one of the most important anabolic chemical reactions that allows life to exist on Earth. With water, light energy from the sun, and carbon dioxide from the air, photosynthetic organisms are able to build simple sugars. Organisms that can make their own food are called autotrophs, and are at the base of the ...

  14. BIOA01 Formal Lab Report

    The effect of herbicide 3-(3,4-dichlorophenyl)- 1,1-dimethylurea) on photosynthesis Tanisha Kunwar 1. 1 Dept. of Biological Sciences, University of Toronto Scarborough, Toronto, Canada. UTSC BIOA01 Lab PRA PRA0004 TA: Shegufta Abstract: This report examines how a herbicide affects the rate of photosynthesis in the aquatic plant, Elodea densa.

  15. Analytical methods to analyze pesticides and herbicides

    The effects of the type of adsorbent, the amount of adsorbent, the desorption solvent, the breakthrough volume, and the adsorption capacity on the extraction performance were also discussed. The results showed that the linear range was 6-600 ng/ml, and the coefficient of determination was 0.9943-0.9998 under the optimal conditions.

  16. BIOA01 Lab 3 Photosynthesis F2022.pdf

    LAB 3 Analysis of Effect of Herbicide on Photosynthesis / Collection of data for FLR Overview Today you will perform a controlled experiment that demonstrates important concepts relating to photosynthesis. You will examine the effect of the herbicide, 3-(3,4-dichlorophenyl)-1,1- dimethylurea (DCMU) on the photosynthetic activity of an aquatic plant, using amount of oxygen produced as an ...

  17. Use Floating Leaf Disks to Study Photosynthesis

    The unit of your photosynthesis rate will be 1/min or min-1. Changing Variables to Investigate Photosynthesis. Now that you are familiar with the leaf disk assay procedure and data analysis, you are ready to start your own investigations. Choose one variable that you want to investigate. You can find some suggestions in Table 2.

  18. Photosynthesis-based biosensors for environmental analysis of herbicides

    A scheme of photosynthesis-based biosensors entailing the bioreceptor (e.g., whole cells, PSII, reaction centers) intimately integrated into possible supports (e.g., paper, screen-printed electrodes, lab on a chip), associated with a dual transduction system (e.g., optical and electrochemical) is reported in Fig. 1.

  19. photo lab report

    Melissa Halverson Herbicides Effect of Photosynthesis Introduction Herbicides are chemical agents that are used to destroy plants or inhibit their growth. Plants cannot grow without the aid of photosynthesis which provides them with some of the nutrients that are needed to grow. The herbicide that was used in the experiment binds to the electron transport proteins associated with photosystem II.

  20. herbicide final .docx

    Photosynthesis is a process used by plants to convert sunlight into chemical energy. During this process, carbon dioxide, water and light energy react to form glucose, oxygen gas and water (MCB181 Lab Manual, 7-4, 9-2). CO2 + H2 O + Light Energy → C 6 H12 O6 + O2 + H2 O During a biochemical process known as the Hill Reaction, DCPIP (2,6-dichlorophenol indophenol) accepts (or steals ...

  21. Herbicides

    Herbicides are applied to water bodies to control aquatic weeds. These weeds can impede irrigation withdrawals or interfere with recreational and industrial uses of water (Folmar et al. 1979). The potential effects of herbicides are strongly influenced by their toxic mode of action and their method of application.

  22. Photosynthesis Lab Report Studoc

    20 to 40°C, photosynthesis decreased as it was affected by the concentration of oxygen and light. intensity (Canvin, Hew, Krotkov, 1969). Not only does carbon dioxide affect photosynthesis, but. it also affects cellular respiration. With the increased carbon dioxide concentration, it tends to.

  23. Photosynthesis inhibitor herbicides

    The photosynthesis inhibitors include these herbicide families: Triazine. Triazinone. Phenylurea. Benzothiadiazole. Nitrile. Phenyl-carbamate. Photosynthesis inhibitors disrupt the photosynthetic (food producing) process in susceptible plants by binding to specific sites within the photosystem II complex in plant chloroplasts. Inhibition of ...