experiment on photosynthesis light intensity

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Photosynthesis Virtual Lab

This lab was created to replace the popular waterweed simulator which no longer functions because it is flash-based. In this virtual photosynthesis lab , students can manipulate the light intensity, light color, and distance from the light source.

A plant is shown in a beaker and test tube which bubbles to indicate the rate of photosynthesis. Students can measure the rate over time. There is an included data table for students to type into the simulator, but I prefer to give them their own handout ,

The handout is a paper version for students to write on as the work with the simulator. The document is made with google docs so that it can be shared with remote students.

There are several experiments that can be done in the lab that would complement this virtual experiment. For example, students can use elodea and measure the number of bubbles released when the plant is under a bright light. Algae beads can also be used to measure changes in pH as the plants consume carbon dioxide.

In experiment 2, students specifically look at light color to determine which wavelength of light increases the rate of photosynthesis. Students should discover that green light has a very slow rate. Their collected data is then compared to a graph of the absorption spectrum of light.

Shannan Muskopf

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Effect of light intensities on the photosynthesis, growth and physiological performances of two maple species

Jinfeng zhang.

1 Beijing Key Laboratory for Forest Resources and Ecosystem Processes, Beijing Forestry University, Beijing, China

2 Optoelectronic College, Beijing Institute of Technology, Beijing, China

Buddhi Dayananda

3 School of Agriculture and Food Sciences, The University of Queensland, Brisbane, QLD, Australia

Associated Data

The raw data supporting the conclusions of this article will be made available by the authors, without undue reservation.

Photoinhibition decreases photosynthetic capacity and can therefore affect the plant survival, growth, and distribution, but little is known about how it affects on kindred tree species. We conducted field experiments to measure the photosynthetic, growth and physiological performances of two maple species ( Acer mono and A. pseudosieboldianum ) seedlings at four light intensities (100%, 75%, 55%, and 20% of full light) and evaluated the adaptability of seedlings. We found that: (1) A. mono seedlings have larger light saturated photosynthetic rates ( A max ), the light saturation point (LSP), and lower light compensation point (LCP) than A. pseudosieboldianum seedlings, thus indicating that the former has a stronger light utilization ability. (2) A. mono seedlings under 75% light intensity and had higher seedling height (SH), basal stem diameter (BSD), leaf number (LN), leaf area per plant (LAPP) and total dry weight (TDW), while A. pseudosieboldianum seedling at 55% light intensity displayed greater growth advantages, which agreed with their response of light saturated photosynthetic rate. Morphological plasticity adjustments such as decreased root shoot ratio (RSR) and increased specific leaf area (SLA) showed how seedlings adapt to weak light environments. (3) 100% and 20% light intensities increased the malondialdehyde (MDA) content of two maple seedlings, indicating that very strong or very weak light could lead to the imbalance of reactive oxygen species (ROS) metabolism. The regulation of antioxidant enzyme activities such as superoxide dismutase (SOD), peroxidase (POD) and catalase (CAT), as well as the content of osmoregulation substances such as free proline and soluble protein, are the main mechanisms of plant adaptation to light stress. Although both A. mono and A. pseudosieboldianum are highly shade tolerant, subtle differences in the photosynthetic, morphological and physiological traits underpinning their shade tolerance suggest A. pseudosieboldianum has the advantage to deal with the light threat. Future studies should focus on the expression level of photosynthesis-related genes and cell, to better understand the adaptation mechanism of plants to light variation which facilitates forest development, either natural or via silvicultural practices. This information expands our understanding of the light-regulating mechanism of trees, which contributes to develop management practices to support natural forest regeneration.

Introduction

Photoinhibition often occurs when light energy is excessive, which reduces photochemical efficiency and even causes photooxidative system damage ( Ma et al., 2015 ; Dias et al., 2018 ). Furthermore, low light intensity influences photosynthesis, which is central to plant productivity, and can therefore severely restrict plant growth ( Zhu et al., 2014 ), and even death ( Wang et al., 2021 ). During the evolutionary process, plants had various adaptive strategies to decrease the potential damage caused by light stress ( Walters et al., 1993 ). Many studies have shown that plants can reduce the direct absorption of light energy by modifying morphological and photosynthetic plasticity, such as decreasing specific leaf weight (SLW), increasing specific leaf area (SLA) or enhancing light utilization capacity through the reduction in the light saturation point (LSP) and lower light compensation point (LCP) ( Kaelke et al., 2001 ; Zhu et al., 2014 ; Sugiura et al., 2016 ). Moreover, plant species can adjust their physiological characteristics in response to the variation in light intensity. For example, high levels of antioxidant enzyme activity which enable the rapid clearance of reactive oxygen species (ROS) ( Ma et al., 2015 ; Ozturk et al., 2021 ). Similarly, osmoregulation substances also play a key role in protecting plants from injury ( Kishor et al., 2005 ; Kučerová et al., 2019 ).

The early growth and survival of seedlings are very important for their successful supplement into the young tree stage, and light intensity plays a determinant role in this stage ( Loik and Holl, 1999 ; Razzak et al., 2017 ). However, in forest development and succession, the light environment varies greatly at both temporal and spatial scales ( Avalos and Mulkey, 2014 ). For example, the destruction and fragmentation of forests are bound to cause sharp changes in light intensity, which may not be beneficial for the regeneration of many trees ( Paquette et al., 2012 ; Yao et al., 2014 ). Even in the forest, the distribution of light is uneven due to the gap and stratification ( Popma and Bongers, 1988 ; Tripathi et al., 2020 ). The adaptability of seedlings to different light environments may determine the status of the tree species in the forest community ( Valladares et al., 2002 ; Rabara et al., 2017 ). In addition, previous studies on seedlings in canopy gaps or forest edges suggest that native tree seedlings may be inhibited by high light ( Yu and Hao, 1998 ; Wu et al., 2006 ).

Maple trees, Acer mono and Acer pseudosieboldianum , belong to the Aceraceae family, which are late succession and shade-tolerant species widely distributed in the natural mixed-broadleaved Korean pine forests in Changbai Mountains, Northeast, China ( Ye et al., 2014 ). These two maple trees are also widely used in landscape architecture construction due to their bright colors ( Xie et al., 2021 ). Previous field investigations found that numerous A. mono has developed into the dominant species in the main story, while A. pseudosieboldianum is the most important constructor in a forest sub-story ( Zhu et al., 2007 ; Ye et al., 2014 ; Zhang et al., 2015 ). Both maple trees are shade tolerant and kindred species, but they have different distribution patterns and abundances in the forest, which may be caused the differentiation in light requirements for the establishment and growth of seedlings ( Paquette et al., 2012 ). Hence, the identification of light requirements is necessary to understanding the regeneration of tree species and facilitating forest development, either natural or via silvicultural practices.

Here, we investigated the light acclimation capacity of A. mono and A. pseudosieboldianum seedlings in response to light conditions, and we hypothesized that: 1) A. mono seedlings may exhibit high photosynthetic efficiency under high light, while A. pseudosieboldianum seedlings may be limited. 2) The photosynthetic, morphological and physiological traits underpinning seedlings’ shade tolerance may give A. pseudosieboldianum an advantage in coping with light threats.

Materials and methods

Seed collection and seedling propagation.

We collected, A. mono and A. pseudosieboldianum seeds from mixed-broadleaved Korean pine forests (127°40’~128°16’ E, 41°35’~42°25’ N) in Changbai Mountains Northeast, China, from late September to early October 2020. Twenty independent individual maple trees were selected. The wings of the seeds were removed during seed collection, and the seeds were soaked in warm water at 45°C (initial temperature) in the laboratory to break the dormancy. The soaked time lasts for 7 days, and the water was renewed every 12 hours. The seeds were mixed with the appropriate amount of sand and put into a pot (30 cm inner diameter, 35 cm height, with good air permeability). Then, the pot with seeds was buried in the ground at 60 cm depth.

We dug out the pots with seeds on April 10, 2021, and then separated the seeds from the pots. The seeds were soaked in 0.5% KMnO 4 solution to disinfect for 3 h, and sterilized seeds were thoroughly rinsed with purified water. The seedbed was built at the Northeast Asia botanical garden in Changbai Mountains. For the seedbed soil disinfection, 1:1500 phoxim was used for insecticidal treatment, then 1:500 carbendazim was used for sterilization, and sowed seeds on 15 April.

Experimental design

To obtain light transmittance, photosynthetically active radiation (PAR) sensors (S-LIA-M003) with HOBO Micro Station Loggers (H21-002) (Onset Computer Corporation, USA) were installed in the forest gap, forest edge and understory of mixed-broadleaved Korean pine forests. The time step for data recording was set at 30 minutes. The light transmittance was calculated according to the following formula:

Four light intensity gradients were set up with different layers of black shade nets in the Northeast Asia botanical garden of Changbai Mountains, Northeast, China. The setup with 100% full light (L 100 ) served as a control, three weak light intensities were set up according to the light transmittance to simulate the forest gap, forest edge and understory. Three weak light intensity treatments were 75% (L 75 ), 55% (L 55 ), and 20% (L 20 ) of full light, which were set up with one layer, two layers, and three layers of nets, and each layer of shading net had three holes. In addition, branches from neighboring trees overtopping the experimental area were removed to secure homogeneous illumination.

On June 5, 2021, the healthy and homogenous seedlings (mean height of A. mono and A. pseudosieboldianum were 19.42 ± 5.32 cm and 21.32 ± 7.57 cm respectively, mean ± SD) were transferred to twenty plastic pots (20 cm inner diameter, 25 cm height, with holes in the bottom, six seedlings per pot) filled with a mixture of black soil, sand, branny, and pearlite (2:2:1:1, v/v/v, 40 kg m -3 ). During the first 15 days, all pots were placed in the built layers shading net for seedling retarding. On July 20, 2021, twenty plastic pots were randomly divided into four groups with five repetitions in each group and moved into the shade nets. In the early stages of the trial, the seedlings were watered every two days.

Photosynthetic measurements

Fully developed leaves (the second, third and the fourth from the top) of three robust seedlings of each tree species were randomly selected under each light environment from August 18 to 25, 2021. Photosynthesis ( P n ) was measured using a portable photosynthesis system (LI-6400, LiCor, Lincoln, NE, USA) at 10 levels of the photosynthetic photon flux density (PPFD), starting from 0, then 40, 60, 80, 100, 150, 200, 400, 600, 800, and 1200 μmol·m -2 ·s -1 . During the measurements, the ambient CO 2 concentrations, the temperature of the leaf chamber and air relative humidity were fixed for 380 μmol·mol –1 , 30°C and 50% respectively. The data were recorded between 8:30 and 11:30 a.m. To fit the photosynthetic light-response curve, we used the non-rectangular hyperbolic photosynthetic model proposed by Ye et al. (2013) . The light-saturated photosynthetic rates ( A max ), LCP, LSP and dark respiration rate ( R d ) were derived from the photosynthetic light-response curve.

Morphological measurements

We harvested seedlings on October 15, 2021. Each seedling, together with its taproots, was carefully removed from the soil, placed into sealed bags, and then transported to the laboratory. The seedlings were carefully washed with tap water and dried using filter papers. The number of leaves in the seedlings was counted, and the seedling height (SH) and basal stem diameter (BSD) were measured using a vernier caliper. The leaf area per plant (LAPP) was measured by a scanner (Canon scan lide120) and analysed using image analyzer (Image J). The seedlings were sorted into leaves, stems, and roots and subsequently dried in a dry oven at 85°C for 48h until constant mass, and then weighed with an electronic balance (EX224ZH 1/10000g; Ohaus Instruments, Changzhou, China). The total dry weight (TDW), root shoot ratio (RSR) and SLA were calculated based on Kelly et al. (2015) :

Physiological measurements

Leaves of two maple seedlings were randomly selected from one pot per treatment on September 1, 2021. The leaves were cut and mixed, they were randomly divided into three groups as three repetitions. The activities of superoxide dismutase (SOD), peroxidase (POD) and catalase (CAT) were determined by the guaiacol method ( Beauchamp and Fridovich, 1971 ), UV absorption method ( Thomas et al., 1982 ), and azoblue tetrazole photoreduction method ( Díaz-Vivancos et al., 2008 ). The content of malondialdehyde (MDA), soluble protein and free proline were determined by the thiobarbituric acid technique ( Deng et al., 2012 ), Coomassie Brilliant Blue G-250 method and ninhydrin staining ( Bates et al., 1973 ; Kučerová et al., 2019 ).

Data analysis

We used the One-way ANOVA to analyze the differences in photosynthetic, morphological and physiological parameters of the two species under different light intensities and the differences between different species under the same light intensity, and Duncan’s multiple range test was used to detect differences between means. All analyses were performed within SPSS (Version 21.0) and Origin 2019.

Light response curves

The light response curves of maple seedlings varied with species. When PPDF< 200 μmol·m -2 ·s -1 , the light response curves of the two species under different light intensities were similar, and the P n increased sharply with the increase of PPDF ( Figures 1A, B ). When PPDF>400 μmol·m -2 ·s -1 , the P n of A. pseudosieboldianum seedlings tended to be stable and reached the LSP ( Figure 1A ). The P n of A. mono tended to be stable when the PPDF>600 μmol·m -2 ·s -1 ( Figure 1B ).

Light-photosynthetic response curves of two maple seedlings under different light intensities. (A) , A. pseudosieboldianum ; (B) , A. mono .

Photosynthetic parameters

The two maple seedlings exposed to 100% intensity showed the lowest A max ( Table 1 ). The A max of A. pseudosieboldianum seedlings was the highest under 55% intensity, while that of A. mono was the highest under 75% intensity. With the decrease in light intensity, the LSP of A. pseudosieboldianum seedlings decreased gradually, and the LSP under 100% intensity was significantly greater than 75%, 55% and 20% intensity ( P< 0.05); A similar response was observed for A. mono seedlings, but it was not significant under the different light intensities. Compared with the 100% light intensity, 75%, 55% and 20% intensity decreased LCP for two species, and the LCP of A. mono seedlings under 100% intensity was significantly greater than 75%, 55% and 20% intensity ( P< 0.05). The R d of two maple seedlings decreased gradually with the increase of light intensity, but a significant difference was not observed.

Table 1

Photosynthetic characteristics of two maple seedlings under different light intensity treatments.

A max , the light-saturated photosynthetic rate; LSP, light saturation point; LCP, light compensation point; R d , dark respiration rate. Small letters indicate significant differences under different light intensities (P< 0.05).

Morphological characters

The shading was beneficial to the growth of two maple seedlings. For example, 55% light intensity resulted in the highest SH, BSD, LN, LAPP, and TDW of A. pseudosieboldianum seedlings, and the seedlings under 75%, 55%, and 20% light intensity were significantly higher than those under 100% light intensity ( P< 0.05) ( Table 2 ). The SH, BSD, LN, and TDW of A. mono seedlings under 75% and 55% light intensity were significantly higher than those under 100% and 55% light intensity ( P< 0.05), and the LAPP was significantly different under different light intensity ( P< 0.05). Two maple seedlings showed decreased RSR in response to dropped light intensity while the SLA increased ( Table 2 ).

Table 2

The growth parameters of two maple seedlings under different light intensity treatments (mean ± SD).

SH, seedling height; BSD, basal stem diameter; LN, leaf number; LAPP, leaf area per plant; TDW, total dry weight; RAS, root shoot ratio; SLA, specific leaf area. Small letters indicate significant differences under different light intensities (P< 0.05).

Antioxidant enzymes activity and MDA content

The SOD activity of A. mono seedling seedlings under 20% and 100% light intensity was higher than that under 55 and 20% light intensity(significance was not observed); and the SOD activity of A. pseudosieboldianum seedlings under 100% light intensity was significantly higher than 55% light intensity ( P< 0.05) ( Figure 2A ). Compared with the 100% light intensity, 75% light intensity decreased POD and CAT activity, while 55% and 20% light intensity increased POD and CAT activity of A. mono seedlings, especially 20% light was significantly higher than 75% light ( P< 0.05). Compared with 100% light intensity, 75%, 55% and 20% light intensity reduced the POD and CAT activity of A. pseudosieboldianum seedlings, and the CAT activity under 100% light was significantly higher than that of 55% and 20% light ( P< 0.05) ( Figures 2B, C ). The 20% light intensity resulted in the lowest MDA content of A. mono seedlings, while the MDA content of A. pseudosieboldianum seedlings was the lowest when the light intensity was 100% ( Figure 2D ).

Effect of light intensity on leaf antioxidant enzymes ( A , SOD; B , POD; C , CAT) and MDA (D) . Small letters indicate significant differences under different light intensities ( P < 0.05).

Content of soluble protein and free proline contents

The soluble protein content of two maple seedlings was significantly different under different light intensities ( Figure 3A , P< 0.05). Among them, 75% light intensity resulted in the lowest soluble protein content of A. mono seedlings, while the soluble protein content of A. pseudosieboldianum seedlings was the lowest when the light intensity was 55%. The free proline content of A. mono seedlings under 20% light intensity was significantly higher than 100%, 75% and 55%, but there was no significant difference among the latter three ( Figure 3B ).

Effect of light intensity on osmoregulation substance ( A , soluble protein; B , free proline contents). Small letters indicate significant differences under different light intensities ( P < 0.05).

Photosynthesis

The light-photosynthetic response curve is the key to understand the photochemical efficiency and photochemical processes of plants ( Loik and Holl, 1999 ; Razzak et al., 2017 ). We found that when PPDF > 400 μmol·m -2 ·s -1 , P n of A. pseudosieboldianum seedlings tended to be stable ( Figure 1A ) while P n of A. mono seedlings tended to be stable when the PPDF>600 μmol·m -2 ·s -1 ( Figure 1B ). This result is consistent with our assumption that as PPDF availability increased, the A. pseudosieboldianum seedlings were difficult to absorb electrons through photochemical processes and on the contrary, A. mono seedlings could deal effectively with the increase in light energy. This variation modes of photosynthetic characteristics may be related to the inherent genetic physiological, and it is also the result of the long-term adaptation of tree species to the environment ( Fariba et al., 2014 ). We also found that A. mono seedlings have higher A max , LSP and lower LCP than A. pseudosieboldianum seedlings in four light gradients ( Table 1 ). This result suggests that the photosynthetic potential for A. mono is high, which may also be the reason why this tree species occupies the forest’s main storey in the natural mixed-broadleaved Korean pine forests. Furthermore, we found that the A max of A. pseudosieboldianum seedlings was the largest at 55% light intensity, while A. mono seedlings exhibited the largest A max at 75% light intensity ( Table 1 ), reflecting that 55% and 75% of full light may be the optimum light levels for the two species respectively. In the field, the optimum light of A. pseudosieboldianum and A. mono is congruent with the habitat choice, which prefers forest gaps, forest edges, and the top of the canopy ( Wu et al., 2006 ; Ye et al., 2014 ).

In this study, the A max of two species under 100% light intensity was significantly lower than 75% and 55% treatments, indicating that the photosynthesis of maple seedlings was limited under strong light. This response to excess light energy is common in other shade tolerant species such as A. Saccharum ( Marilou and Christian, 1998 ), Pinus koraiensis ( Zhu et al., 2014 ), Fagus grandifoli ( Collin et al., 2017 ), and Quercus virginian ( Thyroff et al., 2019 ). Moreover, we found that the LSP and LCP of the two species dropped with the weakening of light intensity, which was consistent with the previous results showing the relatively low LCP and LSP of shade tolerant species were conducive to plants to utilize the light energy more efficiently under weak light environment, thereby increasing the accumulation of organic matter ( Ma et al., 2015 ). Lower R d is generally considered as the adaptive response of plants to cope with shaded conditions and obtain the maximum carbon benefit ( Dias et al., 2018 ). R d of two species under 75%, 55%, and 20% light intensity was lower than that of 100% treatment in our study, although not significant. This suggests that under shading conditions, seedlings reduce the loss of photosynthetic products and maintain the balance of carbon metabolism by decreasing R d , which was also confirmed by Yao et al. (2014) in the study of Abies holophylla .

Seedling growth

Light is a key factor affecting the early growth of tree seedlings in the forest ( Collin et al., 2017 ). Seedling regeneration may fail in shaded habitats with insufficient light ( Dias et al., 2018 ). As a result, seedlings must rely on forest gaps or forest edges to achieve individual regeneration. Previous studies have shown that the greater the light intensity, the better the seedling growth ( Gehring, 2003 ; Kelly et al., 2015 ), however, two maple seedlings exposed to 100% light intensity resulted in significantly lower SH and LAPP compared with the seedlings grown under the 75%, 55%, and 20% light treatments in this study, and BSD, LN and TDW also had a similar trend ( Table 2 ). These results showed that full light has little benefit to the maple seedling growth and is expected that maple trees are reputed to be a late succession and shade tolerant species. Moreover, under the canopy of closed adult plants in the natural mixed-broadleaved Korean pine forests in Northeast China, maples often form a dense seedling bank with a state of growth inhibition, and these seedlings can survive for many years ( Ye et al., 2014 ). Notably, SH, BSD, LN, LAPP, and TDW of A. pseudosieboldianum seedlings were the largest under 55% light intensity, while A. mono seedlings grew best under 75% light intensity ( Table 2 ). The different growth responses of two species to the different light levels may be explained by the photosynthetic variables previously observed in our study, and thus, the optimum light intensity required for seedlings determines their growth. A similar result was also reported in Camptotheca acuminata ( Ma et al., 2015 ) and Tetracentron sinense ( Lu et al., 2020 ).

The modifying of morphological plasticity is an adaptive response of plants to environmental stress (e.g., drought, high salinity and shade) and is also an important way for plants to improve population fitness and resource acquisition ability ( Kitajima, 1994 ; Tripathi et al., 2020 ). In the present study, we found that the SLA of A. pseudosieboldianum seedlings under 20% light intensity was significantly higher than that of 100% treatment, while the RSR under 20% light intensity was significantly lower than that of 100%, 75%, and 55% treatments ( Table 2 ). Similarly, in A. mono seedling, the light of decreased intensity resulted in the increase of SLA and the decrease of RSR ( Table 2 ). This morphological response to variation in light availability has been observed in many other studies ( Popma and Bongers, 1988 ; Avalos and Mulkey, 2014 ; Tang et al., 2015 ). This may be the result of the trade-off between plant biomass aboveground and underground and light stress ( Kitajima, 1994 ). Generally, soil moisture under strong light limits the upward extension of seedlings and eventually affects their growth and survival, thus seedlings allocate more photosynthetic products to the underground to form better developed roots, which is conducive to the absorption of water and nutrition; conversely, the biomass allocation of seedlings under weak light transferred to the aboveground, which can enhance the ability of plants to capture light ( Walters et al., 1993 ; Kaelke et al., 2001 ; Kelly et al., 2015 ; Tang et al., 2015 ). Moreover, we found that SLA and RSR in A. pseudosieboldianum seedlings were higher than that of A. mono seedlings across the light intensity ( Table 2 ). This result is consistent with Canham (1988) which found that shade tolerant species are generally more morphological plastic than less tolerant ones, which helps to improve the resistance and the ability to obtain resources of an individual tree seedling in the weak light environment, hence ensure the long-term reproduction of tree population ( Paquette et al., 2012 ).

Physiological characteristics

In stressful environmental conditions, the imbalance of ROS metabolism and the damage to the cell membrane system can lead to the increase of lipid peroxidation in biomembranes and permeability ( Yi et al., 2020 ), thus resulting in the accumulation of MDA in leaf cells, the product of membrane lipid peroxide, and then decreasing the photosynthetic capacity ( Ozturk et al., 2021 ). In this study, although a significant difference was not observed, the MDA content of two species under 100% and 20% light intensity was higher than that of 75% and 55% treatments ( Figure 2D ). This result agrees well with a recent study that shows full light and deep shade aggravate oxidative damage to lipid membranes ( Wang et al., 2021 ). However, plants have a complete antioxidant enzyme system including SOD, POD, and CAT, which can avoid the damage caused by ROS ( Tang et al., 2015 ). In this study, compared with 75% light intensity, 100% light intensity increased the activities of SOD, POD and CAT of two species ( Figures 2A–C ), indicating that the scavenging ability of ROS was enhanced in the full light environment. This result agrees with the report on olive trees by Sofo et al. (2004) . Similar results were also observed under 20% light intensity and the 20% light intensity enhances antioxidant enzyme activity of two species compared to 55% light intensity ( Figures 2A–C ), which could be due to the fact that the seedlings suffer from light threat under 20% light intensity more grievous than that under 55% light treatment. As a result, seedlings are bound to improve the activity of antioxidant enzymes to resist light stress and reduce light damage ( Ozturk et al., 2021 ).

Another immediate response of plants to cope with light stress is osmotic regulation ( Ozturk et al., 2021 ; Wang et al., 2021 ). For example, free proline can stabilize the construction of membranes and protein by eliminating ROS ( Bates et al., 1973 ; Kishor et al., 2005 ), and soluble protein protects cells against structural-metabolic disruptions and maintain osmolarity ( Ozturk et al., 2021 ). In the present study, an obvious rise in the content of soluble protein and free proline of two species was observed under 100% and 20% light intensities ( Figures 3A and B ), which was consistent with the lower photosynthetic capacity under these two light intensities, indicating that the seedlings increase osmotic regulators to adapt full light and deep shade. Similar results were reported that high levels of soluble protein and free proline maintain cell stability and reduce high/low photo damage ( Wang et al., 2021 ). It is worth noting that the proline and soluble protein content, as well as the above-mentioned three enzyme activities of A. mono seedlings, were the lowest at 75% light intensity, while these of A. pseudosieboldianum seedlings displayed a minimum at 55% light treatment ( Figures 2A–C and Figures 3A, B ), thus the subtle difference supporting their shade tolerance in the plasticity physiological shows that A. pseudosieboldianum more so than A. mono .

Our work demonstrates that full light and deep shade limited the growth of two maple seedlings, the optimum light intensity for the growth of the A. mono and A. pseudosieboldianum seedlings was 75% and 55% of full light, respectively, which can account for the niche of two maple trees in the natural mixed-broadleaved Korean pine forests in Changbai Mountains, Northeast, China. On the other hand, the differentiation in light requirements improves a theoretical basis that in artificial seedling raising and management, appropriate shading should be given to ensure that they are in an optimal light environment. Moreover, while marked differences in growth exist in two maple species, the response in shade conditions is similar, such as increasing antioxidant enzyme activity or osmoregulation substance content, or increasing SLA and reducing RSR, and these responses guarantee the establishment of two tree species in long-term shaded environments. Future studies need to focus on the expression level of photosynthesis-related genes and cell structure, to better understand the adaptation mechanism of higher plants to light variation. Such information expands our understanding of the light-regulating mechanism of endangered plant species and contributes to develop management practices to promote natural forest regeneration.

Data availability statement

Author contributions.

JL designed the research project and provided theoretical guidance. JZ collected and analyzed the data. JZ, JG, and BD wrote the manuscript. All authors contributed to the article and approved the submitted version.

This research was funded by the National Science and Technology Basic Resources Survey Project (SQ2019FY101602).

Acknowledgments

We would like to thank Shixiong Wu for his technical assistance.

Conflict of interest

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.

Biology archive

Course: biology archive   >   unit 11.

• Conceptual overview of light dependent reactions
• Light dependent reactions actors
• Photosynthesis: Overview of the light-dependent reactions

Light and photosynthetic pigments

• The light-dependent reactions

Introduction

What is light energy, pigments absorb light used in photosynthesis, chlorophylls, carotenoids, what does it mean for a pigment to absorb light, attribution:.

• “ The light-dependent reactions of photosynthesis ,” by OpenStax College ( CC BY 3.0 ). Download the original article for free at http://cnx.org/contents/f829b3bd-472d-4885-a0a4-6fea3252e2b2@11 .
• " Bis2A 06.3 Photophosphorylation: the light reactions of photosynthesis ," by Mitch Singer ( CC BY 4.0 ). Download the original article for free at http://cnx.org/contents/c8fa5bf4-1af7-4591-8d76-711d0c1f05f9@2 .

Works cited:

• Chlorophyll a. (2015, October 11). Retrieved October 22, 2015 from Wikipedia: https://en.wikipedia.org/wiki/Chlorophyll_a .
• Speer, B.R., (1997, July 9) Photosynthetic pigments. In UCMP glossary . Retrieved from http://www.ucmp.berkeley.edu/glossary/gloss3/pigments.html .
• Bullerjahn, G. S. and A. F. Post. (1993). The prochlorophytes: are they more than just chlorophyll a/b-containing cyanobacteria? Crit. Rev. Microbiol. 19(1), 43. http://dx.doi.org/10.3109/10408419309113522 .
• Reece, J. B., Urry, L. A., Cain, M. L., Wasserman, S. A., Minorsky, P. V., and Jackson, R. B. (2011). Photosynthesis. In Campbell biology (10th ed.). San Francisco, CA: Pearson, 193.

Additional references:

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Laboratory Manual For SCI103 Biology I at Roxbury Community College

9 photosynthesis.

In this lab, we will study the effect of light intensity and quality (wave length - color) on photosynthesis . As a measure of the rate of photosynthesis, we will monitor the rate of oxygen production. When plants that spend their life submerged in water release oxygen it forms bubbles, which we can count over a period of time to determine photosynthesis rate.

Photosynthesis is a process used by plants and other organisms to convert light energy into chemical energy that can later be released to fuel the organisms’ activities (energy transformation). This chemical energy is stored in carbohydrate molecules, such as sugars, which are synthesized from carbon dioxide and water - hence the name photosynthesis, from the Greek phōs, “light”, and synthesis, “putting together”. In most cases, oxygen is also released as a waste product. Most plants, most algae, and cyanobacteria perform photosynthesis; such organisms are called photoautotrophs. Photosynthesis is largely responsible for producing and maintaining the oxygen content of the Earth’s atmosphere, and supplies all of the organic compounds and most of the energy necessary for life on Earth.

Although photosynthesis is performed differently by different species, the process always begins when energy from light is absorbed by proteins called reaction centers that contain green chlorophyll pigments. In plants, these proteins are held inside organelles called chloroplasts, which are most abundant in leaf cells, while in bacteria they are embedded in the plasma membrane. In these light-dependent reactions, some energy is used to strip electrons from suitable substances, such as water, producing oxygen gas. The hydrogen freed by the splitting of water is used in the creation of two further compounds that act as an immediate energy storage means: reduced nicotinamide adenine dinucleotide phosphate (NADPH) and adenosine triphosphate (ATP), the “energy currency” of cells.

In plants, algae and cyanobacteria, long-term energy storage in the form of sugars is produced by a subsequent sequence of light-independent reactions called the Calvin cycle; some bacteria use different mechanisms, such as the reverse Krebs cycle, to achieve the same end. In the Calvin cycle, atmospheric carbon dioxide is incorporated into already existing organic carbon compounds, such as ribulose bisphosphate (RuBP). Using the ATP and NADPH produced by the light-dependent reactions, the resulting compounds are then reduced and removed to form further carbohydrates, such as glucose.

The first photosynthetic organisms probably evolved early in the evolutionary history of life and most likely used reducing agents such as hydrogen or hydrogen sulfide, rather than water, as sources of electrons. Cyanobacteria appeared later; the excess oxygen they produced contributed directly to the oxygenation of the Earth, which rendered the evolution of complex life possible. Today, the average rate of energy capture by photosynthesis globally is approximately 130 terawatts which is about three times the current power consumption of human civilization. Photosynthetic organisms also convert around 100-115 thousand million metric tons of carbon into biomass per year.

The main source of light on Earth is the Sun. Sunlight provides the energy that green plants use to create sugars mostly in the form of starches, which release energy into the living things that digest them. This process of photosynthesis provides virtually all the energy used by living things. The primary properties of visible light are intensity, propagation direction, frequency or wavelength spectrum, and polarization, while its speed in a vacuum, 299,792,458 meters per second, is one of the fundamental constants of nature. Visible light, as with all types of electromagnetic radiation (EMR), is experimentally found to always move at this speed in a vacuum.

9.1 Intensity of light

Light is electromagnetic radiation within a certain portion of the electromagnetic spectrum (Figure 9.1 ). The word usually refers to visible light, which is visible to the human eye and is responsible for the sense of sight. Visible light is usually defined as having wavelengths in the range of 400-700 nanometres (nm), or 400 × 10 -9 to 700 × 10 -9 m, between the infrared (with longer wavelengths) and the ultraviolet (with shorter wavelengths). This wavelength means a frequency range of roughly 430-750 terahertz (THz).

Figure 9.1: Spectrum of light. V, violet; B, blue; G, green Y, yellow; O, orange; R, red

In this experiment (Figure 9.2 ), we will study the effect of light intensity on the photosynthetic activity of Elodea canadensis . We will vary the light intensity by changing the distance between the light source and the plant. We will count the emerging oxygen bubbles as an indicator of the photosynthetic activity of the plant.

Figure 9.2: Setup for photosynthesis experiment.

9.1.1 Experimental procedures

Before you begin with the actual experiment, write down in your own words the hypothesis for this experiment:

• Obtain a cylindrical test tube.
• Fill test tube with 0.3% sodium bicarbonate.
• Select a fresh, crisp sprig of Elodea about 15 cm in length.
• While the plant is still submerged, cut 2-3 mm from its base.
• Place the sprig upside down into the test tube filled with sodium bicarbonate. The sodium bicarbonate will absorb anu toxic materials that are released by the plant during photosynthesis.
• Keeping the plant submerged, position a light source 10 cm away and adjust so the light shines directly on the plant.
• Place the test tube in a beaker of water as shown in Fig. 9.2 to prevent overheating the plant. 1. 1. Allow the system to stand 7-10 minutes, or until bubbles begin to appear regularly.
• Count the bubbles produced each minute for a 5-minute period and average them. Record your findings in the table.
• Move the light back 20 cm from the plant, wait 5 minutes, and repeat counting. Record your findings in Table 9.1 .
• Move the light back 40 cm from the plant and repeat counting the bubbles.
• When you have finished recording your data, calculate the average number of bubbles for each 5 minute period and enter the result into the table.

Do the data support or contradict your hypothesis?

Figure 9.3: Appearance of bubbles indicates active photosynthesis.

9.2 Color of light

In this experiment, we will study the effect of the color of light on the photosynthetic activity of Elodea canadensis . We will use filter to expose the plant to light of only a limited range of wavelengths. We will again count the emerging oxygen bubbles as an indicator of the photosynthetic activity of the plant.

9.2.1 Experimental procedures

• Empty the test tube that you used in the previous experiment.
• Fill the test tube with fress 0.3% sodium bicarbonate.
• Place the Elodea sprig into the test tube and submerge it completely in the bicarbonate.
• Place the red colored filter between the test tube and the heat shield beaker and allow it to sit for 5 minutes.
• Count bubbles for 5 minutes as in the previous experiment. Record your findings in Table 9.2 .
• Remove the color filter and expose the plant to white light. Count bubbles again for 5 minutes in 1 minute intervals. Record your findings in Table 9.2 .
• Place the green colored filter between the test tube and the heat shield beaker and allow it to sit for 5 minutes.
• Count bubbles for 5 minutes. Record your findings in Table 9.2 . Table: (#tab:color) Color of light.

9.3 Determination of the light absorption spectrum of dye solutions

In this experiment, we will use a spectrophotometer to measure the differential absorption of light of different wavelength by water stained with food dyes.

Figure 9.4: Spectrophotometer and cuvettes with dye solutions.

9.3.1 Experimental procedures

• Take six cuvettes.
• Fill one cuvette with water.
• Fill each of the remaining five cuvettes with one of the color solutions listed in Table 9.3 .
• Insert the cuvette with water into the slot marked “B”.
• Insert the other cuvettes into the slots marked 1 to 5 and write down which color is in which slot.
• Following the instructions posted on the spectrophotometer, program the machine to take absorption measurements at wavelengths between 380-740 nm in 20 nm steps.
• Once the measurements are completed, write down the absorption number for each dye and wavelength.
• Use a spreadsheet program to graph your results.
• Compare your curves with the data shown in Figure 9.6 .

Figure 9.5: Cuvettes placed in the spectrophotometer.

Figure 9.6: Normalized absorption of red, green and blue dye solutions. Compare these data with your own results.

9.4 Chromatography

Chromatography is a laboratory technique for the separation of a mixture. The mixture is dissolved in a fluid called the mobile phase, which carries it through a structure holding another material called the stationary phase. The various constituents of the mixture travel at different speeds, causing them to separate. The separation is based on differential partitioning between the mobile and stationary phases. Subtle differences in a compound’s partition coefficient result in differential retention on the stationary phase and thus affect the separation. Chromatography may be preparative or analytical. The purpose of preparative chromatography is to separate the components of a mixture for later use and is thus a form of purification. Analytical chromatography is done normally with smaller amounts of material and is for establishing the presence or measuring the relative proportions of analytes in a mixture.

In this experiment, we separate a mixture of food dyes (a dark brown liquid). The mobile phase (separation buffer) is 1% NaCl in water, the stationary phase is chromatography paper.

9.4.1 Experimental procedures

• Obtain a small beaker.
• Add NaCl running buffer to the beaker until it reaches a height of about 5 mm.
• Obtain a strip of chromatography paper and put it down on the bench.
• Obtain the bottle containing the dark green food dye mixture.
• Obtain a glass capillary and insert the tip of the capillary into the food dye mixture liquid. A little bit of dye will ascend into the capillary.
• Remove the capillary and apply.
• Touch the left side of the chromatography paper about 1 cm above its lower end with the tip of the capillary. A little bit of green liquid will spread out on the paper. Lift the capillary and touch the paper again just to the right of the dye you just applied. Repeat this until you have a horizontal line of dye from the left to the right side of the paper.
• Place the chromatography paper into the beaker as shown below.
• Observe how the running buffer moves up the paper and separates the dye mixture into three components (red, yellow and blue.

Figure 9.7: Result of the Chromatography experiment.

9.5 Review Questions

• What is light?
• In your own words, describe the endproducts of photosynthesis.
• In your own words, describe what happens in photosynthesis.
• What is chlorophyll and what does it do?
• Where inside of plant cells does photosynthesis happen?
• What is chromatography and what is it used for?

• Science is LIT

Explore How Light Affects Photosynthesis

Algae are aquatic, plant-like organisms that can be found in oceans, lakes, ponds, rivers, and even in snow. But don’t worry, if you’re not near a waterway, it can easily be ordered from Amazon or Carolina Biological. Algae range from single-celled phytoplankton (microalgae) to large seaweeds (macroalgae). Phytoplanktons can be found drifting in water and are usually single-celled. They can also grow in colonies (group of single-cells) that are large enough to see with the naked eye. The specific types of algae that can be used in this experiment are  Scenedesmus, Chlamydomonas, or  Chlorella , all of which are phytoplanktons or microalgae. 

Experimental variables

• Color filter paper
• Table/desk lamp
• Light bulbs (varying intensities and colors)

Laboratory Supplies

• Transfer pipettes
• Vials with caps
• Freshwater Algae ( Scenedesmus , Chlorella , or Chlamydomonas )
• Small beakers or cups

Laboratory Solutions

• 2% Calcium Chloride
• 2% Sodium alginate
• Cresol red/thymol blue pH indicator solution

Solution Preparations

2% calcium chloride (cacl 2 ).

• 20 g of CaCl 2
• Fill to 1000 mL with water

2% CaCl 2 is stable at room temperature indefinitely.

2% Sodium alginate (prepared in advance)

• 2 g sodium alginate
• Fill to 100 mL with water

It takes a while for the alginate to go into solution. We recommend to dissolve by stirring using a magnetic stir bar overnight at room temperature. Store at 4 °C for up to 6 months or use immediately.

Cresol red/Thymol blue pH indicator solution (10x)

• 0.1 g cresol red
• 0.2 g thymol blue
• 0.85 g sodium bicarbonate (NaHCO 3 )
• 20 mL ethanol
• Fill to 1L with fresh boiled water

Measure indicators and mix with ethanol. Measure sodium bicarbonate and mix with warm/hot water. Mix the solutions together and fill with remaining freshly boiled water up to 1L final solution. The 10x stock solution is stable for at least a year.

In preparation for doing the experiment, prepare 1x indicator solution by diluting the 10x indicator solution with distilled water (e.g. 20 ml 10x into 200 mL final solution).

Experimental Bench Set-Up

• ~10 mL of 2% CaCl 2 in a cup or beaker
• ~3-5 mL of sodium alginate in cup or beaker
• Cup with ~10 mL of water
• Empty cup or beaker that holds a minimum of 30 mL

Preparing Algae for Experiment

• Prepare a concentrated suspension of algae. Without centrifuge : leave ~50 mL of algae suspension to settle (preferably overnight), then carefully pour off the supernatant to leave ~3-5 mL of concentrated algae. With centrifuge : Centrifuge ~50 mL of algae suspension at low speed for 10 minutes and then carefully pour off the supernatant, leaving behind ~3-5 mL of concentrated algae.
• In a small beaker, add equal volumes of sodium alginate and then add in the concentrated algae. Gently mix algae and sodium alginate together using a transfer pipette until its evenly distributed.
• Using the transfer pipette, carefully add single drops of the algae/sodium alginate mixture into the CaCl 2 to make little “algae balls”
• Once all of the “algae balls” are in the CaCl 2 solution, allow them to harden for 5 minutes
• Place the strainer over the empty cup or beaker, and pour over the entire solution of “algae balls” and CaCl 2 into the strainer allowing the CaCl 2 to pass through, leaving just the algae in the strainer
• Keeping the strainer over the container, pour the water over the “algae balls” to rinse the remain CaCl 2
• Transfer your newly made “algae balls” to a new cup or beaker

Setting up Photosynthesis Experiment

• Distance from light (using ruler) – group can set up vials different distances from one light source
• Different color lights (using color filter paper or different color light bulbs) – group can set up by covering the vials with different colored films and arrange them the same distance away from the light source or set up 1 vial in front of a different colored lamp same distance away.
• With or without light – group places 1 vial in front of an illuminated lamp and another has the vial or lamp covered with black paper the same distance away

• When starting your experiment, be sure to take note of the time that you placed your vial in front of the light source. Vials should be left for ~1-2 hours.

What would happen if the algae photosynthesizes (increase O2) in a solution that started at pH8.2?

Analyzing photosynthesis results

• After 1-2 hours, return to the experiment. Without disturbing the vials, analyze and take pictures of results. Have students write down the time that their experiment ended.
• Using the color chart above, determine which pH matches your sample the closest.
• Have students determine if they got what they expected and discuss amongst their group members.

Explain how the rate of photosynthesis is affected by their different variables.

What were your conclusions from this experiment? If you were to repeat the experiment, what would you change and why? What’s the relationship with O2 and CO2 during the process of photosynthesis? Is there a “best” source of light that allowed the algae to photosynthesize better?

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Practical Biology

A collection of experiments that demonstrate biological concepts and processes.

Observing earthworm locomotion

Practical Work for Learning

Published experiments

Investigating photosynthesis using immobilised algae, class practical.

This procedure offers a method for measuring the rate of photosynthesis which depends directly on the rate of uptake of carbon dioxide by the photosynthetic organism. Hydrogencarbonate indicator changes colour with pH, which is determined by the concentration of carbon dioxide in solution.

The photosynthetic organism is a fast-growing green alga – such as Scenedesmus quadricauda – immobilised in alginate beads. These algal balls make it easy to standardise the amount of photosynthetic tissue in any investigation.

There is scope for students to develop the protocol to investigate a range of factors.

This protocol is adapted with permission from information on the Science and Plants for Schools (SAPS) website (see www.saps.org.uk ).

Lesson organisation

You can make up the algal balls in one lesson, discuss how to set up the main procedure, and carry it through in the next lesson. A preliminary investigation by teacher/ technician will allow you to estimate the amount of algal material and indicator to use to get a result with your equipment in the time available.

Apparatus and Chemicals

For each group of students:.

Transparent containers (glass bottles are ideal) with sealable lids, around 10 cm 3 , 6–12

Clamp stand, boss and clamp

Syringe barrel, 10 cm 3

Beaker, 100 cm 3 , 2

Cocktail stick to stir alginate

150 W lamp ( Note 5 )

Container of water as heat filter ( Note 6 )

Ruler/ tape measure

For the class – set up by technician/ teacher:

Colorimeter ( Note 1 )

Algal suspension, 2.5 cm 3 concentrated for each group ( Note 2 )

Sodium alginate, 2–3%, 100 cm 3 for a class of 30 students ( Note 3 )

Calcium chloride solution, 2%, 50 cm 3 per group

Hydrogencarbonate indicator, 50–100 cm 3 per group ( Note 4 )

Hydrogencarbonate indicator, standard colour scale, if colorimeter not available ( Note 4 )

Light meters, if available

Health & Safety and Technical notes

Do not look directly into the lamps. Do not touch the lamps while hot. Keep flammable material away from the lamps in use.

Read our standard health & safety guidance

1 Colorimeter. There is a linear relationship between absorbance of the indicator (at 550 nm – bright green filter) and pH over the range studied in this procedure.

2 Growing your alga. Prepare a culture of green alga such as unicellular Scenedesmus quadricauda . Make up a solution of algal enrichment medium, and subculture the alga into this. Aerate gently and keep at temperatures between 18–22 °C. Constant illumination ensures faster growth of the alga. After 3–4 weeks, the culture should have a green ‘pea soup’ colour. Subculture the alga again to maintain a healthy culture. You could use other algae, but Scenedesmus should produce 2 to 3 litres of dark green ‘soup’ in about 4 weeks from 50 cm 3 of original culture. (Details from SAPS Sheet 23).

3 Preparing solutions to make alginate beads (Refer to Recipe card 2):

• Dissolve 3 g of sodium alginate in 100 cm 3 of cold, pure water. Stir with a spatula every half hour or so. Leave overnight and stir in the morning.
• Dissolve 4 g of calcium chloride-6-water in 200 cm 3 of pure water in a 250 cm 3 beaker.

4 Hydrogencarbonate indicator. Refer to Recipe card 34 and Hazcard 32. Low hazard once made; must be made fresh by qualified staff using fume cupboard. The indicator is very sensitive to changes in pH, so rinse all apparatus with the indicator before use. Avoid exhaling over open containers of the indicator. Make up a ‘standard colour scale’ of reaction bottles containing buffers from pH 7.6- pH 9.2 with hydrogencarbonate indicator if students will not have access to a colorimeter.

5 Lamps. You need a brighter light than a standard 40 W or 60 W bench light. Low energy bulbs produce too limited a spectrum of light for full activity. 150 W tungsten or halogen lamps are best. 150 W portable halogen lamps have a stand and handle separate from the body of the lamp which makes them safer to handle. But they do produce heat, so you will need a heat filter for the investigation ( Note 6 ).

6 Heat filter. Use a large flat-sided glass vase (available from Ikea or Homebase or other domestic suppliers) or a medical ‘flat’ filled with water. With a high power lamp, the small volume in a medical ‘flat’ may get too hot for comfort.

7 Making alginate beads:

• When making up the alginate or diluting the algal culture it is essential to use pure water; otherwise calcium ions in the water will cause the alginate to 'set' prematurely.
• If your beads are not the size and texture you want, try different mixes with your active material, or different concentrations of sodium alginate (around 2–3%), or make the syringe nozzle narrower (with glass capillary tubing) or wider (by sawing off and adding a plastic tube). Different brands of alginate have different consistencies. You need a viscous mixture that will drip steadily through the syringe. Keep a note of what worked for this supply and keep your syringe barrels with nozzles for next time.

8 Neutral density filters: can be sourced as a film from photographic suppliers (for example, Lee filters www.leefilters.com . A neutral density filter reduces the amount of light transmitted by the same amount at all wavelengths.

Ethical issues

There are no ethical issues associated with this procedure.

SAFETY: Take precautions to avoid burns, fires and dazzle caused by hot, bright lamps. Do not leave the apparatus unattended overnight as the lamps are so hot.

Preparation of algal beads

a Concentrate your active algal culture ( Note 2 ). Do this by leaving 50 cm 3 to settle for 30 minutes in a measuring cylinder until you have a darker green sediment. Carefully pour off the pale green suspension to leave just 5 cm 3 of concentrated culture.

b Make up the solutions you need to prepare alginate beads ( Note 3 ).

c Mix 5 cm 3 of the algal culture with 5 cm 3 of the 3% sodium alginate solution ( Note 3 ) in a very small beaker.

d Clamp a syringe barrel above a beaker of calcium chloride solution (see diagram and Note 7 ) – making sure the tip of the syringe is well above the solution in the beaker.

e Pour the alga/alginate mixture through the syringe barrel so it drips through and forms beads in the beaker. Swirl the beaker gently as the drops fall. ( Note 7 .)

f Allow the beads to harden for a few minutes before straining them out of the beaker through a tea strainer.

g Rinse the beads in distilled water. The algae in the beads will stay alive for several months in a stoppered bottle of distilled water in the refrigerator.

Other materials

h Make up hydrogencarbonate indicator (see Note 4 and Recipe card 34). It is important to make up enough for the whole investigation as the depth of colour of this indicator is so variable. Make a few litres and keep for the duration of the investigations, aerating before each lesson. Make up a ‘standard colour scale’ of reaction bottles containing buffers from pH 7.6- pH 9.2 with hydrogencarbonate indicator if students will not have access to a colorimeter.

a Rinse 6 translucent bottles with hydrogencarbonate indicator solution.

b Add equal numbers of algal balls to each container – around 20.

c Add a standard volume of indicator to each container (7–10 cm 3 ).

d Replace the lid.

e Note the colour of the solution against a standard set of bottles of indicator, and/or measure the absorbance at 550nm using a colorimeter.

f Put one container in the dark. Set up the other containers at different light intensities – either by placing at different distances from the lamp, or by wrapping different neutral density filters ( Note 8 ) around the bottles.

g Place a heat filter between the light and the containers to absorb heat from the lamp. ( Note 6 .)

h After 30 minutes, note the colours of the solution at each distance.

i Once the bottles cover a range of colours, which will probably take more than 60 minutes, take samples from each bottle and measure the absorbance at 550 nm using a colorimeter. Start with the bottle that has changed most – the one at highest light intensity.

j Plot absorbance against light intensity.

k You could measure light intensity at each distance if a light meter is available, or calculate light intensity (1 ÷ distance 2 ).

Teaching notes

It takes at least an hour for colour changes to happen – so students will need to return to the lab at break or after lessons to ‘read’ the results.

Plotting absorbance against light intensity for the above procedure will show a proportional relationship at low light intensities, levelling off at higher intensities which indicates another limiting factor. The method with neutral density filters (rather than changing distance from the lamp) is reportedly more accessible for some students.

Carbon dioxide is unlikely to be the limiting factor here as the indicator contains a relatively high concentration of hydrogencarbonate ions.

Other factors to investigate include:

• amount of algal material: Shake 1 litre of algal culture, and leave 500 cm 3 , 250 cm 3 or 100 cm 3 to settle in measuring cylinders, saving the bottom 5 cm 3 each time. Make up into balls as above – each batch will have a different amount of algae, but the same surface area to volume ratio for the balls.
• surface area to volume ratio: Make smaller balls using a narrower bore to drip the alginate through.
• wavelength of light: Use coloured filters wrapped around the containers, or try different lamps (if you can identify the spectrum of light produced). Coloured filters will also alter the light intensity.

This practical reportedly works well for coursework investigations as students can suggest improvements or developments after some preliminary work with the technique.

Set aside a container of indicator and algal beads to keep in the lab for a few days. Students will be able to see changes to the colour of indicator according to the time of day – algae use up more carbon dioxide than they produce during the day and release carbon dioxide at night when they are not taking up any.

There is a linear response between absorbance of the indicator (at 550nm) and pH over the range studied in this procedure. Using a colorimeter (at 550nm) is one way to quantify this procedure. Alternatively you could set up a ‘standard colour scale’ with buffer solutions and indicator at the same concentration as the reagent in the investigation. Students can then compare their reaction vessels with the standards and interpolate to estimate pH.

You can also use this technique of immobilisation in an alginate bead to study enzymes or yeast cells. The beads can be packed into a column (for example, a syringe barrel) and a suitable substrate passed over the beads. Collect the products at the bottom of the column and use the immobilised enzymes or cells again and again without the need to separate them from the reactants.

Health and safety checked, September 2008

Related experiments

Investigating factors affecting the rate of photosynthesis

Investigating the light dependent reaction in photosynthesis

Adapted with permission from information on the Science and Plants for Schools (SAPS) website (see www.saps.org.uk ).

Investigating the Rate of Photosynthesis ( AQA A Level Biology )

Revision note.

Biology & Environmental Systems and Societies

Apparatus & Techniques: Investigating the Rate of Photosynthesis

• Investigations to determine the effects of light intensity, carbon dioxide concentration and temperature on the rate of photosynthesis can be carried out using aquatic plants , such as Elodea or Cabomba (types of pondweed )
• Light intensity – change the distance ( d ) of a light source from the plant (light intensity is proportional to 1/ d 2 )
• Carbon dioxide concentration – add different quantities of sodium hydrogencarbonate (NaHCO 3 ) to the water surrounding the plant, this dissolves to produce CO 2
• Temperature (of the solution surrounding the plant) – place the boiling tube containing the submerged plant in water baths of different temperatures
• For example, when investigating the effect of light intensity on the rate of photosynthesis, a glass tank should be placed in between the lamp and the boiling tube containing the pondweed to absorb heat from the lamp – this prevents the solution surrounding the plant from changing temperature
• Distilled water
• Aquatic plant, algae or algal beads
• Sodium hydrogen carbonate solution
• Thermometer
• Test tube plug
• This will ensure oxygen gas given off by the plant during the investigation form bubbles and do not dissolve in the water
• This will ensure that the plant contains all the enzymes required for photosynthesis and that any changes of rate are due to the independent variable
• Ensure the pondweed is submerged in sodium hydrogen carbonate solution (1%) – this ensures the pondweed has a controlled supply of carbon dioxide (a reactant in photosynthesis)
• Cut the stem of the pondweed cleanly just before placing into the boiling tube
• Measure the volume of gas collected in the gas-syringe in a set period of time (eg. 5 minutes)
• Change the independent variable (ie. change the light intensity, carbon dioxide concentration or temperature depending on which limiting factor you are investigating) and repeat step 5
• Record the results in a table and plot a graph of volume of oxygen produced per minute against the distance from the lamp (if investigating light intensity), carbon dioxide concentration, or temperature

The effect of light intensity on an aquatic plant is measured by the volume of oxygen produced

Results - Light Intensity

• The closer the lamp, the higher the light intensity (intensity ∝ 1/ d 2 )
• Therefore, the volume of oxygen produced should increase as the light intensity is increased
• This is when the light stops being the limiting factor and the temperature or concentration of carbon dioxide is limiting the rate of photosynthesis
• The effect of these variables could then be measured by increasing the temperature of water (by using a water bath) or increasing the concentration of sodium hydrogen carbonate respectively
• Rate of photosynthesis = volume of oxygen produced ÷ time elapsed

Limitations

• Immobilised algae beads are beads of jelly with a known surface area and volume that contain algae, therefore it is easier to ensure a standard quantity
• Immobilised algae beads are easy and cheap to grow, they are also easy to keep alive for several weeks and can be reused in different experiments
• The method is the same for algae beads though it is important to ensure sufficient light coverage for all beads

Light intensity – the distance of the light source from the plant (intensity ∝ 1/ d 2 )

Temperature - changing the temperature of the water bath the test tube sits in

Carbon dioxide - the amount of NaHCO 3 dissolved in the water the pondweed is in

Also remember that the variables not being tested (the control variables) must be kept constant.

Required Practical: Affecting the Rate of Dehydrogenase Activity

• The light-dependent reactions of photosynthesis take place in the thylakoid membrane and involve the release of high-energy electrons from chlorophyll a molecules
• These electrons are picked up by the electron acceptor NADP in a reaction catalysed by dehydrogenase
• However, if a redox indicator (such as DCPIP or methylene blue ) is present, the indicator takes up the electrons instead of NADP
• DCPIP: oxidised ( blue ) → accepts electrons → reduced ( colourless )
• Methylene blue: oxidised ( blue ) → accepts electrons → reduced ( colourless )
• The colour of the reduced solution may appear green because chlorophyll produces a green colour
• When light is at a higher intensity, or at more preferable light wavelengths, the rate of photoactivation of electrons is faster, therefore the rate of reduction of the indicator is faster

Light activates electrons from chlorophyll molecules during the light-dependent reaction. Redox indicators accept the excited electrons from the photosystem, becoming reduced and therefore changing colour.

• Isolation medium
• Pestel and mortar
• Aluminium Foil

Method - Measuring light as a limiting factor

• This produces a concentrated leaf extract that contains a suspension of intact and functional chloroplasts
• The medium must have the same water potential as the leaf cells so the chloroplasts don’t shrivel or burst and contain a buffer to keep the pH constant
• The medium should also be ice-cold (to avoid damaging the chloroplasts and to maintain membrane structure)
• The room should be at an adequate temperate for photosynthesis and maintained throughout, as should carbon dioxide concentration
• If different intensities of light are used, they must all be of the same wavelength (same colour of light) - light intensity is altered by changing the distance between the lamp and the test tube
• If different wavelengths of light are used, they must all be of the same light intensity - the lamp should be the same distance in all experiments
• DCPIP of methylene blue indicator is added to each tube, as well as a small volume of the leaf extract
• A control that is not exposed to light (wrapped in aluminium foil) should also be set up to ensure the affect on colour is due to the light
• This is a measure of the rate of photosynthesis
• A graph should be plotted of absorbance against time for each distance from the light
• This is because the lowered light intensity will slow the rate of photoionisation of the chlorophyll pigment, so the overall rate of the light dependent reaction will be slower
• This means that less electrons are released by the chlorophyll, hence the DCPIP accepts less electrons. This means that it will take longer to turn from blue to colourless
• A higher rate of decrease, shown by a steep gradient on the graph, indicates that the dehydrogenase is highly active.
• This experiment is not measuring the rate of dehydrogenase activity directly (through measuring the rate of substrate use or product made) but is instead predicting what the rate would be by measuring the rate of electron transfer from the photosystems
• It is therefore important to control the amount of leaf used to produce the chloroplast sample and also how much time is spent crushing the leaf to release the chloroplast
• It is also a good idea to measure a specific wavelength absorption by each sample on the colorimeter before and after the experiment so you can get a more accurate change in oxidised DCPIP concentration
• Results should also be repeated and the mean value calculated
• The time taken to go colourless is subjective to each person observing and therefore one person should be assigned the task of deciding when this is

In chemistry the acronym ‘OILRIG’ is used to remember if something is being oxidised or reduced. Oxidation Is Loss (of electrons) and Reduction Is Gain (of electrons). Therefore the oxidised state is when it hasn’t accepted electrons and the reduced state has accepted electrons.

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Author: Alistair

Alistair graduated from Oxford University with a degree in Biological Sciences. He has taught GCSE/IGCSE Biology, as well as Biology and Environmental Systems & Societies for the International Baccalaureate Diploma Programme. While teaching in Oxford, Alistair completed his MA Education as Head of Department for Environmental Systems & Societies. Alistair has continued to pursue his interests in ecology and environmental science, recently gaining an MSc in Wildlife Biology & Conservation with Edinburgh Napier University.

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As light intensity increases, what happens to the rate of photosynthesis?

The following image provides more nuance. It shows that light intensity and the rate of photosynthesis increase with one another. It also shows that the rate at which photosynthesis levels out is dependent upon other factors—both plants in 0.1% CO₂, they cannot photosynthesize at nearly the rate of the plants in 0.4% CO₂.

Similarly, plants photosynthesize at a greater rate in higher temperatures (generally—not in temperatures that are too hot—this is also dependent upon species).

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