Measurement of Chlorophyll Loss Due to ... - ACS Publications

Laboratory Experiment pubs.acs.org/jchemeduc

Measurement of Chlorophyll Loss Due to Phytoremediation of Ag Nanoparticles in the First-Year Laboratory Kurt Winkelmann,* Leonard Bernas, Brendan Swiger, and Shannon Brown Department of Chemistry, Florida Institute of Technology 150 West University Boulevard, Melbourne, Florida 32901, United States S Supporting Information *

ABSTRACT: A two-week experiment is presented in which students can observe the impact of nanoparticles on the concentration of chlorophyll in plants. First-year students in an introductory nanotechnology laboratory course and a general chemistry laboratory course synthesized silver nanoparticles and then exposed stalks of Egeria densa (E. densa), a common waterweed, to the nanoparticle solution for 1 week. In the following session, students extracted chlorophyll from the plants and measured its concentration using a visible spectrometer. Compared to other, similar lab activities, this experiment generates a lower amount of waste, requires a shorter duration of plant growth, and involves the measurement of chemical species in order to determine the toxicological effects of nanomaterials. Additional ideas are discussed for implementing the experiment in high school, general chemistry, and other courses.

KEYWORDS: First-Year Undergraduate/General, Laboratory Instruction, Hands-On Learning/Manipulatives, Nanotechnology

■

INTRODUCTION Nanoscale materials are becoming an increasingly prevalent component of new technology for medicine and health, consumer goods, electronics, and industrial processes. In order to meet the current and expected demand for workers with nanotechnology-related skills, educators are beginning to teach this subject at all grade levels.1,2 Besides preparing students for their careers, educators must help students learn about nanotechnology so that they are better-informed citizens in a highly technological world. Researchers are concerned about the toxicity of nanoparticles. Their novel properties, so useful for new materials, may also lead to unwanted chemical reactions when nanomaterials are released into the environment. Small nanoparticles may enter biological cells disrupting cell functions, and they may decompose into metal ions, which are themselves toxic. Negative effects of a nanomaterial may vary depending on the target organism,3 the particular environment around the nanoparticle,4 particle size,3 coating,5 and chemical composition.6 As primary producers in the food chain, the impact of pollution on plants is a significant concern. An effective remediation strategy involves the intentional introduction of plants into a polluted area to absorb pollutants. The plants are then harvested for later treatment. This process, known as phytoremediation, is relatively low-cost and generates less chemical waste. For these reasons, plants are frequently studied in order to measure the environmental impact of chemical pollutants and to determine the best option for removal of contaminants. One such plant found to be effective for phytoremediation is Egeria densa (E. densa), also known as Elodea densa.7 © XXXX American Chemical Society and Division of Chemical Education, Inc.

Silver nanoparticles (Ag NPs) are known to exhibit biocidal properties, leading to their use in medical equipment, water purification, and personal care products. They are among the most widely used nanomaterials,8 and they may also form in the natural environment from soluble silver salts.9 The fate of Ag NPs in the environment depends on their stability. Nanoparticles may agglomerate to form micron-sized (bulk) silver particles which precipitate. Bulk silver is less toxic (though not nontoxic) because the larger particles have less surface area and they would be too big to enter cells of plants and animals. Silver nanoparticles may be oxidized to form insoluble silver salts such as AgCl or Ag2O. Again, the silver compound would precipitate from the aqueous environment. The silver has not disappeared, but it is less accessible when it is part of the sediment. Alternatively, nanoparticles may exhibit increased stability if they are coated in natural organic matter. This enables them to spread through waterways far from their original source of pollution.10 Silver nanoparticles can disrupt the biological functions of a cell through two mechanisms. The nanoparticle may adsorb to the cell and release silver cations as a product of various redox reactions. Silver cations enter the cell through sodium and copper ion channels. The cell may also completely absorb the nanoparticle through a process called endocytosis. This is less common for larger nanoparticles due to their size. Once in the cell, the nanoparticle may release Ag+ or silver atoms on the Received: September 13, 2016 Revised: March 14, 2017

A

DOI: 10.1021/acs.jchemed.6b00694 J. Chem. Educ. XXXX, XXX, XXX−XXX

Journal of Chemical Education

■

Laboratory Experiment

PROCEDURE Working in pairs, students synthesize silver nanoparticles during the first 3 h lab session of this two-week experiment. Students combine equal volumes (e.g., 10 mL) of 4.7 × 10−3 M silver nitrate and 0.20 M sodium citrate solutions in an Erlenmeyer flask and then heat the solution to 80 °C for 15−20 min in a water bath with occasional stirring. Students remove the solution when it begins to turn a light yellow color. The solution color continues to change to a deeper yellow as it cools to room temperature.20 Students measure the visible spectrum of their solution using a visible spectrophotometer such as those offered by Vernier. The instructor should acquire E. densa plants prior to the first week of the experiment. The authors have successfully performed this experiment using E. densa purchased from local aquarium stores or ordered from Carolina Biological Company. The authors acclimate their plants for 1 week in an aquarium filled with dechlorinated tap water (water dechlorinator is available at pet stores). E. densa will survive and grow for several months as it floats on the water in an aquarium equipped with a light and water filter. The authors remove any roots which have formed on the selected stalks, cut them into 10 cm portions, and place the stalks in a beaker of aquarium water immediately before the lab session begins. Each pair of students needs four of these pieces. Since E. densa absorbs water through the stalk, it does not matter if students use the top, middle, or bottom of a long stalk. Students prepare 40 mL solutions for the E. densa plants in each 50 mL test tube. Two test tubes contain just DI water (>1 MΩ) for the controls. Students can choose a volume of nanoparticle solution to add (a range 1−10 mL works well) to the two other test tubes and then dilute the solution to 40 mL, yielding concentrations between 6 × 10−5 and 5.9 × 10−4 M (6−63 ppm) total silver. A single laboratory session provides enough time for students to extract chlorophyll from a total of four plant samples during the following week of the experiment. The authors placed the students’ test tubes, uncovered, on a window sill for 1 week. Use of a greenhouse is unnecessary. During the second week of the experiment, students extract chlorophyll. They place each plant in a small beaker inside a drying oven at 110 °C for about 15 min and then remove the leaves from the dried plants. They weigh the dried leaves and then transfer leaves from a single stalk to a mortar and grind them into a powder. Students transfer the leaf powder into a small test tube with the aid of small portions of cold 95% ethanol. Students add a total of 15−20 mL of cold 95% ethanol to the test tube. Chlorophyll dissolves in the ethanol for 10 min. During that time, students can set up a Buchner funnel with filter paper, a side arm flask, and a vacuum hose for filtration. One student can filter the solution while another student begins the extraction procedure for the leaves of another stalk. Some ethanol will evaporate so students should record the final volume of filtrate recovered. Students measure an absorption spectrum of the filtrate at 649, 664, and 750 nm. Chlorophyll does not absorb at 750 nm; this value accounts for any turbidity of the solution, which is assumed to cause a constant absorbance at the other wavelengths measured. Before calculating the concentration of chl a and chl b, students should subtract the absorbance at 750 nm from the absorbance values at 649 and 664 nm to determine the corrected absorbance values. The absorbance at 664 and 649 nm should be below 1 absorbance unit. If the

nanoparticle surface that can bind with proteins and negatively affect their activities.11 Chlorophyll (chl), the green pigment involved in plant photosynthesis, includes two conjugated molecules, chl a and chl b. Chlorophyll compounds absorb blue and red light, creating excited π electrons. The light energy is converted into chemical energy through a reaction with an electron acceptor. Chlorophyll a and b molecules have absorption maxima at slightly different wavelengths so their individual concentrations can be detected spectroscopically. They are found primarily in the leaves of the plant. Chlorophyll molecules degrade when plants are under stress, such as when they absorb toxins from their environment. Thus, a lower chlorophyll concentration can indicate that the plant is under stress due to its environment or toxins. Due to their widespread uses, vivid size-dependent colors, and ease of synthesis, Ag NPs are featured in many nanotechnology lab experiments described in this Journal and elsewhere.12−18 Most procedures are appropriate for a general chemistry lab course, and some can be performed by high school students. Students can observe the antimicrobial properties of Ag NPs19,20 and their detrimental effects on plants21 and brine shrimp.22 A recent article in this Journal shows a simple method for illustrating the toxicological effects of other nanoparticles on beans.23 The laboratory experiment reported here allows students to synthesize Ag NPs and measure their toxicological effects on stalks of E. densa, a waterweed that is inexpensive and readily available. In the subsequent laboratory session, students extract chlorophyll from the leaves. Recent volumes of this Journal have frequently described the extraction and study of plant pigments in general chemistry.24−27 The loss of chlorophyll pigments is a measure of the harm caused by the silver nanoparticles. This has several advantages over the previously published silver nanoparticle experiments: • E. densa stalks react within 1 week of exposure to Ag NPs, so this experiment spans only two lab sessions, unlike other experiments which can require three or more weeks of plant growth. • Plants used in this experiment float in water and do not require soil. This greatly reduces the amount of solid waste generated. • Students perform a chemical extraction of chlorophyll in order to quantify the damage caused by the silver nanoparticles, rather than simply measuring the physical properties of plants, such as the lengths of stalks and roots. • Both the plant species used and the extraction of biomolecules from plants make this experiment similar to those conducted in nanomaterial phytoremediation research. The goals of this experiment are for students to synthesize silver nanoparticles and measure their effect on the natural environment. Their performance during the lab session and their answers to laboratory report questions provide a means for assessing the achievement of these goals. Students have successfully performed this experiment as part of an introductory nanotechnology lab course28,29 and in a General Chemistry 2 lab session. In both cases, most students were first-year engineering and science majors who had completed only one semester of general chemistry. B




Figure 1. Visible absorbance spectrum of silver nanoparticles prepared by a student.

used when performing this experiment. Silver nanoparticles in the chemical waste can be converted to bulk silver by adding excess NaCl.

absorbance values are not, students can dilute their extract solution with more ethanol. The relevant absorbance peaks are 664 nm for chl a and 649 nm for chl b. The absorption spectrum of each pigment overlaps with the other.24 The concentration of each pigment is determined from the absorbance of one wavelength subtracted from the other with the coefficients of the absorbance based on Beer’s Law constants for the two pigments. Equations 1 and 2 illustrate the relationship between the absorbance values at both wavelengths and pigment concentrations. Both equations include corrected absorbance values (A) at wavelengths of 649 and 664 nm to determine the concentration of chlorophyll a and b.30 [chl a] (μg/mL) = 13.36A 664 − 5.19A 649

(1)

[chla] (μg/mL) = 27.43A 649 − 8.12A 664

(2)

■

Concentrations of chl a and chl b are frequently reported in the research literature with units of mg of pigment per g of plant sample. Results of eqs 1 and 2 are expressed as dry weight concentrations using the conversion shown in eq 3. ⎛ mg ⎞ ⎛ μg ⎞ filtrate volume (mL) [chl]⎜ DW⎟ = [chl]⎜ ⎟ × dried leaf mass (g) ⎝ g ⎠ ⎝ g ⎠ 1 mg × 100 μg

RESULTS AND DISCUSSION

After the light yellow solution cools, students have finished preparing Ag nanoparticles. Figure 1 shows a typical spectrum of a student’s solution with λmax = 411 nm. A TEM image of Ag NPs prepared by the authors with a particle size distribution is shown in Figure 2.31 Particle size is bimodal, with the mean diameter equal to 5.7 ± 2.5 nm for the 95% of particles below 15 nm in diameter and a mean diameter of 27 ± 8 nm for the larger particles. Particle size analysis is based on measurements of 469 particles. These results are consistent with the previously published particle sizes of Ag NPs formed from a citrate reduction.20 Heating the silver nitrate and citrate solution at a temperature that is too high leads to rapid nanoparticle formation. In that scenario, it can be difficult for students to remove the flask from the water bath before the solution turns a brown color, indicating that the nanoparticles may be too large. Heating at too low a temperature can cause the formation to occur too slowly. The authors have found that a temperature range 75−80 °C yields satisfactory results. Nanoparticle solution concentrations of less than 30 ppm are sufficient to cause visually obvious, negative effects on the plants. Students typically recovered between 0.04 and 0.06 g of dried leaves, which provides a sufficient mass for chlorophyll extraction. If the dry weight was too low, students could combine plants from similar samples to create a single sample with greater mass. Students’ largest source of error was the extent to which they grind the leaves. Thoroughly grinding the leaves exposes more leaf surface area, increasing the amount of chlorophyll which students extract. Differences in the quality of grinding led to large variations in results among different lab groups, but since each group tended to grind each of their four samples the same

(3)

Students record their values on a data sheet and report their values to the instructor. The combined class data is reported here. A copy of the experiment handout and instructors’ notes are included in Supporting Information.

■

HAZARDS Students should wear goggles, closed-toe shoes, and lab coats or aprons. Hot plates used to heat the water bath will be hot so students should exercise caution when handling them. Ethanol is a flammable liquid. No open flames or heat sources should be C




Figure 2. (a) TEM image of silver nanoparticles formed by the reduction of citrate at 80 °C. (b) Particle size distribution.

way, each group could observe differences between their control and treatment plant stalks. Although some students did not grind their dried leaves as thoroughly as instructed, all students achieved at least acceptable results. In a class with multiple groups treating plants with the same concentrations of nanoparticles, their varying results tended average out. Figure 3 shows a graph of pooled student data collected in an introductory nanotechnology lab course. It illustrates the decline in concentration of chlorophyll a and b in Egeria densa plants exposed to various concentrations of silver nanoparticles. Silver species do not specifically target

chlorophyll. Rather, silver nanoparticles can bind to proteins, enzymes, and other biomolecules, preventing them from functioning correctly. If the nanoparticle is oxidized and releases Ag+, cations can cause oxidative stress within the cell. Even prior to death, these conditions can diminish the amount of chlorophyll that the plant produces. This results in the plant showing brown or pale green leaves. Figure 4 shows the deteriorated condition of plants after 1 week of exposure to Ag nanoparticles prepared according to the laboratory handout. The plant stalk used for the control (left-most in Figure 4) D




Figure 3. Pooled student data showing the loss of chlorophyll a and b in the E. densa plants as a function of silver nanoparticle concentration in solution. Error bars represent standard deviation values. Five of the original 80 data points were excluded from the analysis as outliers based on the Q test (95% confidence).

Figure 4. Elodea densa plants removed from test tubes after exposure to (from left to right) 0, 13, 25, 38, and 51 ppm Ag NP solutions.

appears unchanged. Note the gradual loss of green color from the leaves and stem as the concentration of Ag NPs increases. In addition to grinding the leaves into a powder, some students failed to remove the flask of light yellow solution from

the heat at the proper time. By letting the solution heat too long, the nanoparticles continue to grow. When cooled, the solution appears brown, not yellow, indicating that the nanoparticles are larger. Instructors can assess the quality of E



■

students’ nanoparticle synthesis based on the absorption spectrum of the Ag nanoparticle solution, but students can continue to use their nanoparticles for the second part of the experiment. Despite their larger size, the nanoparticles still have similar toxicological effects on the plant. Laboratory reports were collected for students in the General Chemistry 2 lab course offered during the summer term. These reports revealed that students learned about the importance of particle size with regard to nanoparticle properties. In response to a discussion question, students correctly explained that bulk particles would have less of an effect on the plant. They also surmised that using the fresh weight of the plants, rather than the dry weight, would potentially lead to inconsistent results due to the variable amount of water in the samples. Students correctly discussed their data in terms of the negative impact that silver nanoparticles had on their plants. Students’ views of the experiment were positive. They appreciated that the experiment linked laboratory work with a larger environmental issue. Several students remarked in their lab reports that they enjoyed working with plants. Others noted that this was the first time they had learned anything about nanotechnology. These comments indicate that the experiment intertwines a variety of topics which appeal to students. The authors have tested different versions of this experiment during its development, and author L.B. has performed a simplified version with high school students. Descriptions of these can be found in the instructors’ notes (Supporting Information).

CONCLUSIONS As nanotechnology becomes more ubiquitous, attention is being paid to teaching students about the environmental impact of nanomaterials. Measurement of extracted chlorophyll from plants is a chemically relevant method of evaluating plant health and is appropriate for first-year students. Students learned about nanotechnology and its effect on plants, and their feedback indicates that they enjoyed studying the chemistry of a living system. The experiment could be expanded in a variety of ways, as discussed in the instructors’ notes. ASSOCIATED CONTENT

S Supporting Information *

The Supporting Information is available on the ACS Publications website at DOI: 10.1021/acs.jchemed.6b00694. Laboratory handout (PDF) (DOCX) Instructors’ notes (PDF) (DOCX)

■

REFERENCES

(1) Nanoscale Science and Engineering Education; Sweeney, A., Seal, S., Eds.; American Scientific Publishers: Stevenson Ranch, CA, 2008. (2) Global Perspectives of Nanoscience and Engineering Education; Winkelmann, K., Bhushan, B., Eds.; Springer: Berlin, 2016. (3) Glenn, J. B.; White, S. A.; Klaine, S. J. Interactions of gold nanoparticles with freshwater aquatic macrophytes are size and species dependent. Environ. Toxicol. Chem. 2012, 31, 194−201. (4) Chinnapongse, S. L.; MacCuspie, R. I.; Hackley, V. A. Persistence of singly dispersed silver nanoparticles in natural freshwaters, synthetic seawater, and simulated estuarine waters. Sci. Total Environ. 2011, 409, 2443−2450. (5) Colman, B. P.; Espinasse, B.; Richardson, C. J.; Matson, C. W.; Lowry, G. V.; Hunt, D. E.; Wiesner, M. R.; Bernhardt, E. S. Emerging contaminant or an old toxin in disguise? Silver nanoparticle impacts on ecosystems. Environ. Sci. Technol. 2014, 48, 5229−5236. (6) Van Koetsem, F.; Xiao, Y.; Luo, Z.; Du Laing, G. Impact of water composition on association of Ag and CeO2 nanoparticles with aquatic macrophyte Egeria canadensis. Environ. Sci. Pollut. Res. 2016, 23, 5277−5287. (7) Romero-Oliva, C. S.; Contardo-Jara, V.; Pflugmacher, S. Time dependent uptake, bioaccumulation and biotransformation of cell free crude extract microcystins from Lake Amatitlán, Guatemala by Ceratophyllum demersum, Egeria densa and Hydrilla verticillata. Toxicon 2015, 105, 62−73. (8) Vance, M. E.; Kuiken, T.; Vejerano, E. P.; McGinnis, S. P.; Hochella, M. F. J.; Rejeski, D.; Hull, M. S. Nanotechnology in the real world: Redeveloping the nanomaterial consumer products inventory. Beilstein J. Nanotechnol. 2015, 6, 1769−1780. (9) Akaighe, N.; Maccuspie, R. I.; Navarro, D. A.; Aga, D. S.; Banerjee, S.; Sohn, M.; Sharma, V. K. Humic acid-induced silver nanoparticle formation under environmentally relevant conditions. Environ. Sci. Technol. 2011, 45, 3895−3901. (10) Levard, C.; Hotze, E. M.; Lowry, G. V.; Brown, G. E. Environmental transformations of silver nanoparticles: Impact on stability and toxicity. Environ. Sci. Technol. 2012, 46, 6900−6914. (11) Wigginton, N. S.; de Titta, A.; Piccapietra, F.; Dobias, J.; Nesatyy, V. J.; Bernier-Latmani, R.; Suter, M. J. F. Binding of silver nanoparticles to bacterial proteins depends on surface modifications and inhibits enzymatic activity. Environ. Sci. Technol. 2010, 44, 2163− 2168. (12) Paniconi, G.; Chung, S.-J.; Liou, S.-C.; Kim, S.; Kim, L.; Kang, S.; Kim, D. Three Green Approaches to the Synthesis of Silver Nanoparticles Using Pentose and Hexose Carbohydrates. Chem. Educator 2016, 21, 73−76. (13) Cooke, J.; Hebert, D.; Kelly, J. A. Sweet nanochemistry: A fast, reliable alternative synthesis of yellow colloidal silver nanoparticles using benign reagents. J. Chem. Educ. 2015, 92, 345−349. (14) Paluri, S.; Edwards, M.; Lam, N.; Williams, E.; Meyerhoefer, A.; Sizemore, I. E. P. Introducing “Green” and “Nongreen” Aspects of Noble Metal Nanoparticle Synthesis: An Inquiry-Based Laboratory Experiment for Chemistry and Engineering Students. J. Chem. Educ. 2015, 92, 350−354. (15) Dong, Z.; Richardson, D.; Pelham, C.; Islam, M. R. Rapid Synthesis of Silver Nanoparticles using a Household Microwave and their Characterization: A Simple Experiment for Nanoscience. Chem. Educator 2008, 4171, 240−243. (16) Soukupova, J.; Kvitek, L.; Kratochvilova, M.; Panacek, A.; Prucek, R.; Zboril, R. Silver voyage from macro- to nanoworld. J. Chem. Educ. 2010, 87, 1094−1097. (17) Mulfinger, L.; Solomon, S. D.; Bahadory, M.; Jeyarajasingam, A. V.; Rutkowsky, S. A.; Boritz, C. Synthesis and Study of Silver Nanoparticles. J. Chem. Educ. 2007, 84, 322−325. (18) Richardson, A.; Janiec, A.; Chan, B. C.; Crouch, R. D. Synthesis of silver nanoparticles: an undergraduate laboratory using green approach. Chem. Educator 2006, 11, 331−333. (19) Gardner, G. E.; Jones, M. G. Bacteria Buster: Testing Antibiotic Properties of Silver Nanoparticles. Am. Biol. Teach. 2009, 71, 231−234.

■

■


AUTHOR INFORMATION

Corresponding Author

*E-mail: kwinkel@fit.edu. ORCID

Kurt Winkelmann: 0000-0002-2016-602X Notes

The authors declare no competing financial interest.

■

ACKNOWLEDGMENTS This material is based upon work supported by the National Science Foundation under Grant 1140841. F




(20) Bauer, C.; Garner, O.; Saldarriaga, L.; Rauda, I.; Chia, H. Silver Nanoparticle Biotoxicity Experiment. California NanoSystems Institute, 2012. Available upon request from publishers at http://cnsi. ctrl.ucla.edu/nanoscience/pages/biotoxicity (accessed March 2017). (21) Metz, K. M.; Sanders, S. E.; Miller, A. K.; French, K. R. Uptake and Impact of Silver Nanoparticles on Brassica rapa: An Environmental Nanoscience Laboratory Sequence for a Nonmajors Course. J. Chem. Educ. 2014, 91, 264−268. (22) Maurer-Jones, M. A.; Love, S. A.; Meierhofer, S.; Marquis, B. J.; Liu, Z.; Haynes, C. L. Toxicity of nanoparticles to brine shrimp: An introduction to nanotoxicity and interdisciplinary science. J. Chem. Educ. 2013, 90, 475−478. (23) Ross, S. S.; Owen, M. J.; Pedersen, B. P.; Liu, G.; Miller, J. W. Using Mung Beans as a Simple, Informative Means To Evaluate the Phytotoxicity of Engineered Nanomaterials and Introduce the Concept of Nanophytotoxicity to Undergraduate Students. J. Chem. Educ. 2016, 93, 1428−1433. (24) Israsena Na Ayudhya, T.; Posey, F. T.; Tyus, J. C.; Dingra, N. N. Using a Microscale Approach To Rapidly Separate and Characterize Three Photosynthetic Pigment Species from Fern. J. Chem. Educ. 2015, 92, 920−923. (25) Dias, A. M.; Ferreira, L. S. Supermarket Column Chromatography of Leaf Pigments” Revisited: Simple and Ecofriendly Separation of Plant Carotenoids, Chlorophylls, and Flavonoids from Green and Red Leaves. J. Chem. Educ. 2015, 92, 189−192. (26) Sjursnes, B. J.; Kvittingen, L.; Schmid, R. Normal and ReversedPhase Thin Layer Chromatography of Green Leaf Extracts. J. Chem. Educ. 2015, 92, 193−196. (27) Wesolowski, M. C. Integrating Biology into the General Chemistry Laboratory: Fluorometric Analysis of Chlorophyll a. J. Chem. Educ. 2014, 91, 1224−1227. (28) Winkelmann, K.; Mantovani, J.; Brenner, J. In Interdisciplinary Lab Course in Nanotechnology for Freshmen at the Florida Institute of Technology; Sweeney, A. E., Seal, S., Eds.; American Scientific Publishers: Stevenson Ranch, CA, 2008; pp 269−291. (29) Winkelmann, K. Practical Aspects of Creating an Interdisciplinary Nanotechnology Laboratory Course for Freshmen. J. Nano Educ. 2009, 1, 34−41. (30) Lichtenthaler, H. K. Chlorophylls and Carotenoids: Pigments of Photosynthetic Biomembranes. Methods Enzymol. 1987, 148, 350− 382. (31) Brenner, J. R. Incorporation of Data Analysis Throughout the Chemical Engineering Curriculum Made Easy with DataFit. Chem. Eng. Educ. 2007, 253−257.

G


Measurement of Chlorophyll Loss Due to ... - ACS Publications

Recommend Documents