Name: a low-cost method of measuring the in situ primary productivity of periphyton communities of lentic waters
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Description: Watch this Scientific Journal Video about A Low-Cost Method of Measuring the In Situ Primary Productivity of Periphyton Communities of Lentic Waters at JoVE.com

This study was supported by the Czech Science Foundation (GACR 19-05791S), RVO 67985939, and by the CAS within the program of the Strategy AV 21, Land save and recovery. Many thanks to Ond&#345;ej Sihelsk&#253; for taking the shots in the field - without him, the filming would have been complete hell. The project would not be possible without tight cooperation with companies, Palivov&#253; Kombin&#225;t &#218;st&#237; s.p. and Sokolovsk&#225; Uheln&#225;, who provided access to the studied localities.

<ol>
	<li>Blachart, J. L., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Potential+consequences+of+climate+change+for+primary+production+and+fish+production+in+large+marine+ecosystems.">Potential consequences of climate change for primary production and fish production in large marine ecosystems.</a> Philosophical Transactions of the Royal Society B: Biological Sciences. 367 (1605), 2979-2989 (2012).</li><li>Howarth, R. W., Michaels, A. F., Sala, O. E., Jackson, R. B., Mooney, H. A., Howarth, R. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+Measurement+of+primary+production+in+aquatic+ecosystems.">The Measurement of primary production in aquatic ecosystems.</a> Methods in Ecosystem Science. , 72-85 (2000).</li><li>Vadenbecouer, Y. E. G., Peterson, M. J., Vander, Z., Kalff, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Benthic+algal+production+across+lake+size+gradients:+Interactions+among+morphometry,+nutrients,+and+light.">Benthic algal production across lake size gradients: Interactions among morphometry, nutrients, and light.</a> Ecology. 89 (9), 2542-2552 (2008).</li><li>Reimer, A., Landmann, G., Kempe, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Lake+Van,+eastern+Anatolia,+hydrochemistry+and+history.">Lake Van, eastern Anatolia, hydrochemistry and history.</a> Aquatic Geochemistry. 15 (1), 195-222 (2009).</li><li>Cantonati, M., Lowe, R. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Lake+benthic+algae:+toward+an+understanding+of+their+ecology.">Lake benthic algae: toward an understanding of their ecology.</a> Freshwater Sciences. 33 (2), 475-486 (2014).</li><li>Gaarder, T., Gran, H. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Investigation+of+the+production+of+plankton+in+the+Oslo+Fjord.+Rapports+et+Proces-verbaux+des+R&#233;unions.">Investigation of the production of plankton in the Oslo Fjord. Rapports et Proces-verbaux des R&#233;unions.</a> Conseil International pour l'&#201;xploration de la Mer. 42, 1-48 (1927).</li><li>Hall, R. O., Thomas, S., Gaiser, E. E., Fahey, T. J., Knapp, A. K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Measuring+Freshwater+Primary+Productivity+and+Respiration.">Measuring Freshwater Primary Productivity and Respiration.</a> Principles and Standards for Measuring Primary Productivity. , (2007).</li><li>Howart, R., Michaels, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Chapter+6+The+Measurement+of+Primary+Production+in+Aquatic+Ecosystems.">Chapter 6 The Measurement of Primary Production in Aquatic Ecosystems.</a> Springer Science and Business Media LLC. , (2000).</li><li>Kop&#225;&#269;ek, J., Hejzlar, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Semi-micro+determination+of+total+phosphorus+in+soils,+sediments,+and+organic+materials:+a+simplified+perchloric+acid+digestion+procedure.">Semi-micro determination of total phosphorus in soils, sediments, and organic materials: a simplified perchloric acid digestion procedure.</a> Communications in Soil Science and Plant Analysis. 26 (11-12), 1935-1946 (1995).</li><li>Benson, B. B., Krause, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+concentration+and+isotopic+fractionation+of+oxygen+dissolved+in+freshwater+and+seawater+in+equilibrium+with+the+atmosphere1.">The concentration and isotopic fractionation of oxygen dissolved in freshwater and seawater in equilibrium with the atmosphere1.</a> Limnology and Oceanography. 29 (3), 620-632 (1984).</li><li>Dodds, W. K., Biggs, B. J., Lowe, R. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Photosynthesis-irradiance+patterns+in+benthic+microalgae:+variations+as+a+function+of+assemblage+thickness+and+community+structure.">Photosynthesis-irradiance patterns in benthic microalgae: variations as a function of assemblage thickness and community structure.</a> Journal of Phycology. 35 (1), 42-53 (1999).</li><li>Bott, T. L., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=An+evaluation+of+techniques+for+measuring+periphyton+metabolism+in+chambers.">An evaluation of techniques for measuring periphyton metabolism in chambers.</a> Canadian Journal of Fisheries and Aquatic Sciences. 54 (3), 715-725 (1997).</li><li>Blankenship, R. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Structural+and+functional+dynamics+of+photosynthetic+antenna+complexes.">Structural and functional dynamics of photosynthetic antenna complexes.</a> Proceedings of the National Academy of Sciences. 112 (45), 13751-13752 (2015).</li><li>Hawes, I., Schwartz, A. -. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Photosynthesis+in+an+extreme+shade+environment,+benthic+microbial+mats+from+Lake+Hoare,+a+permanently+ice-covered+Antarctic+lake.">Photosynthesis in an extreme shade environment, benthic microbial mats from Lake Hoare, a permanently ice-covered Antarctic lake.</a> Journal of Phycology. 35 (3), 448-459 (1999).</li><li>Aristegui, J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Planktonic+primary+production+and+microbial+respiration+measured+by+14C+assimilation+and+dissolved+oxygen+changes+in+coastal+waters+of+the+Antarctic+peninsula+during+austral+summer:+Implications+for+carbon+flux+studies.">Planktonic primary production and microbial respiration measured by 14C assimilation and dissolved oxygen changes in coastal waters of the Antarctic peninsula during austral summer: Implications for carbon flux studies.</a> Marine Ecology-Progress Series. 132, 191-201 (1996).</li><li>Steemann-Nielsen, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+use+of+radioactive+carbon+(14C)+for+measuring+organic+production+in+the+sea.">The use of radioactive carbon (14C) for measuring organic production in the sea.</a> Journal of Experimental Marine Biology and Ecology. 18 (2), 117-140 (1952).</li><li>Sanz-Mart&#237;n, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Relationship+between+carbon-and+oxygen-based+primary+productivity+in+the+Arctic+Ocean,+svalbard+archipelago.">Relationship between carbon-and oxygen-based primary productivity in the Arctic Ocean, svalbard archipelago.</a> Frontiers in Marine Science. 6, 468 (2019).</li><li>Nielsen, E. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Measurement+of+the+production+of+organic+matter+in+the+sea+by+means+of+carbon-14.">Measurement of the production of organic matter in the sea by means of carbon-14.</a> Nature. 167 (4252), 684-685 (1951).</li><li>J&#246;nsson, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+14C-incubation+technique+for+measuring+microphytobenthic+primary+productivity+in+intact+sediment+cores.">A 14C-incubation technique for measuring microphytobenthic primary productivity in intact sediment cores.</a> Limnology and Oceanography. 36 (7), 1485-1492 (1991).</li><li>Bender, M. L., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+comparison+of+four+methods+for+determining+planktonic+community+production.">A comparison of four methods for determining planktonic community production.</a> Limnology and Oceanography. 32 (5), 1085-1098 (1987).</li><li>&#352;imek, K., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Spatio-temporal+patterns+of+bacterioplankton+productivity+and+community+composition+related+to+phytoplankton+composition+and+protistan+bacterivory+in+a+dam+reservoir.">Spatio-temporal patterns of bacterioplankton productivity and community composition related to phytoplankton composition and protistan bacterivory in a dam reservoir.</a> Aquatic Microbial Ecology. 51 (3), 249-262 (2008).</li></ol>

The authors confirm that they have no conflicts of interest to disclose.

The methodology described in this paper is based on the principle of the light and dark bottle oxygen technique in combination with the noninvasive technique of measuring O2 concentration using optical oxygen sensors. This system allows the parallel measurement of different incubation settings as the optical fiber for measuring O2 can be moved quickly from bottle to bottle. The benthic communities from various depths can differ in taxonomic composition and productivity; simultaneously measuring them in a parallel manner can save time and make the results more precise, enabling the calculation of whole-depth profiles where periphyton grow. Non-invasive oxygen sensors integrated into shake flasks enable real-time oxygen flux monitoring.
The designed experimental setup allows measurement of the primary productivity of microbial communities at respective depths where the communities naturally occur, which is difficult to simulate in the laboratory (appropriate pressure, track the changes in the diurnal course of temperature and light intensity in situ, different nutrient availability). The major improvements in the proposed method are noninvasiveness, cost-effectiveness, and the possibility of multiple simultaneous measurements at a single coordinate location near the place of sample collection. The method is environmentally friendly as all materials can be used repeatedly. It also enables the collection of large datasets without additional expenses. The introduction of highly sensitive O2 probes (i.e., detection limit 15 ppb, resolution 0.1% of 0.04 mg O2 L-1) also shifts the usage of the method to less productive aquatic environments.
Nevertheless, the proposed method also has limitations that are associated with the nature of the experimental arrangement. The first limitation is the possibility of O2 supersaturation of the closed bottles. This occurs especially during sunny days if the highly productive biomass is incubated. It can result in an underestimation of photosynthesis caused by the incomplete dissolution of O2 in the water7. High concentrations of O2 can also deactivate PSII11. To avoid supersaturation, preliminary experiments focused on biomass amount optimization are strongly recommended. Second, the exchange of produced gases can be limited by the thick diffusive boundary layer of the periphytic mat. Together with the stagnant conditions within incubation bottles, it can lead to underestimation of productivity. However, within the framework of the described methodology, the incubation bottles are manipulated every hour when pulled out from the water during the measuring time and lowered back then. During incubation, the autotrophic community could also become limited by dissolved inorganic carbon (DIC), causing a deceleration in production. Therefore, the increase in O2 is not linear over the entire incubation time. Usually, the problem is solved by selecting only the linear region of the O2 to time relationship for rate calculation. Here, we used a third-degree polynomial regression, which serves as an approximation of any differentiable function defining the rate of change of O2 over time. It allows for calculation of the rate of change at time zero; hence, the community is not yet limited by the DIC at time zero.
Studies have shown that minimal agitation for homogenization of gases between periphytic mats and overlying water is necessary12. In lentic environments, the hydrodynamics of littoral zones is not strong anyway, so mixing the water within the incubation bottle during the individual measurements can be a sufficient compromise to natural homogenization processes. Another problematic step is pulling out the periphytic sample from the physical context of the bottom, which may lead to the alteration of light exposure and gas penetration to the mat. The quantity of illumination of periphyton can be slightly modulated, especially at the edges of the collected mats. However, photosynthetic organisms face light intensity fluctuations naturally, so they are adapted to maintain such changes13,14. To avoid a greater degree of disturbance, it is recommended to carefully sample one compact piece of the periphytic mat for incubation. Samples may be exposed to more intense radiation when they are pulled out from the water during measurements. Nevertheless, the incubation time of the bottles in the respective depths is much larger than the time required to conduct O2 concentration reading. The incubation time is 4 h, and one reading takes approximately 30 s.
The presented method was invented for the lentic water ecosystems. It can also be used in running waters, but some minor technical problems must be solved. In running water, the incubation bottles can be broken more easily and anchoring the whole system would be challenging in the flow. Nevertheless, even in stagnant water, incubation bottles must be handled carefully so they do not break each other during the manipulation. Special attention must be paid if the incubation bottles include microbial mats growing on stones. It is critical not to sample big stones to avoid breaking the glass bottles.
As noted in protocol section 7, the described method provides estimates of net ecosystem productivity, ecosystem respiration rate, and their product-gross ecosystem productivity. In this method, respiration rates of heterotrophic and autotrophic parts of the microbial community cannot be distinguished. Hence, the gross ecosystem productivity is lower than gross primary productivity by a rate equal to heterotrophic respiration. Therefore, this method is not directly comparable to the frequently used method of primary productivity estimation by 14C isotope labeling15,16,17. This method is based on introducing a known amount of labeled 14C-CO2 into incubation bottles filled with water of known dissolved inorganic C concentration18. The radioactivity loss rate over time is directly proportional to gross primary productivity. The proposed method, in fact, complements the radiolabeling method because the combination of both methods theoretically allows for distinguishing all carbon fluxes, including heterotrophic and autotrophic respiration. However, the radiolabeling method also has several limitations. For example, it relies on the assumption of rapid and homogeneous incorporation of the labeled element into the periphytic mat19, which is not usually possible without the mechanical disruption of the periphyton matrix. To obtain accurate daily estimates, samples must be collected, incubated, and sampled several times throughout the day, and the method should be run under laboratory conditions to avoid radioactive contamination7. Continuous and noninvasive measurement is not possible.
Other commonly used methods for estimating primary productivity in aquatic ecosystems have the same experimental design but include different types of oxygen probes, which are less convenient, especially for estimating the primary productivity of periphyton20. The use of some types of oxygen probes can help detect the actual oxygen concentration within the periphytic mat but cannot estimate the rate of change over time unless sealed in a closed system. Sealing the oxygen probe is more prone to leakage than noninvasive optical sensors. Due to their need for wired connections, other bottle-based metabolism methods do not allow measurement of oxygen concentrations at greater depths. This makes the proposed method, which does not require a wired connection, more capable of providing accurate estimates of deep-water benthic metabolism. As each optical sensor is an independent unit, it is possible to simultaneously measure the change in oxygen concentration over time at multiple depths at a single coordinate location. As a result, the method could provide a robust dataset that even allows upscaling to the entire water body if required. The remote sensing technique, for example, is not similarly applicable because this method cannot be used at varying depths below the surface.
Detailed seasonal in situ measurements of gross productivity of periphyton (or other benthic organisms) can improve current knowledge of the processes controlling primary productivity dynamics in lentic waters. Estimating the total primary productivity in each lake&#39;s littoral zone can give a better idea of its relative importance for the whole-lake carbon metabolism. However, this method is unsuitable for measuring plankton primary productivity in oligotrophic waters. Compared to benthic communities, the plankton biomass in open water is too low. Thus, the changes in O2 concentration would be too slow to be detected simultaneously during the experiment with benthic biomass. In this case, the metabolic activity of plankton can be measured using the 14C-bicarbonate method as described by &#352;imek et al.21 in the same sampling time. In eutrophic waters, it is possible to use this method to measure the primary productivity of plankton as well. Each technique for measuring primary productivity mentioned above has advantages and disadvantages, which have been discussed. The best technique should be selected according to the scientific questions being investigated. Further, the method should mimic the ecological parameters of studied ecosystems as much as possible.

Primary productivity is the only entry of autochthonous carbon into the aquatic systems that form the entire system food web1. Hence, the accurate estimation of primary productivity is an essential step toward understanding the functioning of aquatic ecosystems. Littoral zones are areas of high primary productivity and biodiversity. In addition to phytoplankton, periphyton (hereafter referred to as microbial mats) and macroalgae are assumed to significantly contribute to primary productivity in littoral zones2. Due to their sessile lifestyle and significant spatial heterogeneity, quantification of primary productivity is not trivial. 
Primary productivity is driven mainly by light intensity, temperature, availability of nutrients, and distribution of the ionic species of the carbonate system in the respective depths of euphotic zones3,4. The depth markedly influences the spatial distribution of microbial mats. Microbial communities must cope with the adverse effects of high irradiation and pronounced seasonal temperature variations in shallow depths and with lower light intensity at greater depths. In addition to the depth gradient, dynamic trophic interactions generate multiple and complex spatial patterns at different scales5. This complex system is complicated to simulate in the laboratory. The most accurate way to infer the metabolic activity of individual primary producers from littoral zones is to set up in situ experiments. 
The methodology introduced in this paper is based on the traditional chamber method2,6,7, together with a transportable and easy-to-build low-cost floating barge. This allows the measurement of primary productivity at different depths under the natural light spectrum, temperature, and different distribution of the ionic species of the carbonate system with the depth. The method is based on the principle of light versus dark bottle oxygen, which was first employed to measure phytoplankton photosynthesis6 and is still commonly used6,7. It compares the rate of change in oxygen in bottles kept in the light (which includes the effects of primary productivity and respiration) with those held in the dark (respiration only)8. The method uses oxygen evolution (photosynthesis) as a proxy for primary productivity. The measured variables are net ecosystem productivity (NEP, as a change in O2 concentration over time in light conditions) and ecosystem respiration (RE, as a change in O2 concentration over time in the dark). Gross ecosystem productivity (GEP) is the calculation of the difference between the two (Table 1). The term "ecosystem" is used here to denote that the periphyton is composed of autotrophic and heterotrophic organisms. The most significant improvement of this traditional chamber method is using noninvasive oxygen optical sensors and optimization of this primarily planktonic method for measuring periphytic primary productivity.
The technique is described in the example of measuring microbial mats in the littoral zone of newly emerged post-mining lakes in the Czech Republic-Milada, Most, and Medar. The metabolic activity of microbial mats is determined using direct in situ measurement of O2 fluxes performed directly at specific depths, where the studied communities naturally occur. Heterotrophic and phototrophic activity is measured in closed glass bottles equipped with noninvasive optical oxygen sensors. These sensors detect the partial pressure of oxygen using the fluorescence of light-sensitive dyes. The bottles with microbial mats are suspended and incubated on a floating device at the appropriate depths. The oxygen concentration inside the bottles was continuously measured during the daylight period from the small boat. 
Samples of intact microbial mats are collected and placed in gas-tight incubation bottles at designated depths by scuba divers. Each bottle is equipped with a noninvasive optical oxygen microsensor, which monitors the O2 productivity/consumption over time. All measurements are done in five replicate dark/light pairs in each depth. The temperature and photosynthetically active radiation (PHAR) intensities are measured at respective depths throughout the incubation. After 6 h of in situ incubation (daylight hours), the microbial mats are harvested from the bottles and dried. O2 fluxes are normalized to microbial biomass. As a control, fluxes are corrected for changes in O2 concentration in separate light and dark gas-tight bottles (blank controls) containing lake water without microbial mat biomass. Below are detailed instructions for building the floating barge and performing the whole experiment step-by-step. This paper also presents representative results from the measurements of microbial mats at two depths (1 m and 2 m), with five replicates at each depth. Actual temperature and light intensity were measured during the whole experiment using dataloggers.

<table><tbody><tr><td>Aluminum angle L profile 40 x 40 mm x 3 mm, length 2,000 mm</td><td></td><td></td><td></td></tr><tr><td>Aluminum flat bar 40 x 3 x 350 mm</td><td></td><td></td><td></td></tr><tr><td>Bucket 15 L with concrete infill&amp;nbsp;</td><td></td><td></td><td></td></tr><tr><td>Carabine hook with screw lock 50 x 5 mm</td><td></td><td></td><td></td></tr><tr><td>electric tape black</td><td></td><td></td><td></td></tr><tr><td>Extruded polystyrene (XPS) material 500 x 200 x 150 mm</td><td></td><td></td><td></td></tr><tr><td>Fibox 3 LCD trace</td><td>PreSens Precision Sensing GmbH</td><td></td><td>stand-alone fiber optic oxygen meter</td></tr><tr><td>Hondex PS-7 Portable Depth Sounder</td><td>Hondex&amp;nbsp; - Honda Electronics</td><td></td><td>to measures distances through water - to bottom depth measurement; https://www.honda-el.net/industry/ps-7e</td></tr><tr><td>KORKEN - glass tight-seal jar 0.5 L</td><td>IKEA</td><td></td><td>incubation bottles; https://www.ikea.com/cz/en/p/korken-jar-with-lid-clear-glass-70213545/</td></tr><tr><td>metal hook&amp;nbsp;</td><td></td><td></td><td></td></tr><tr><td>Oxygen Sensor Spot SP-PSt3-NAU-D5</td><td>PreSens Precision Sensing GmbH</td><td></td><td>non-invasive optical oxygen sensor for measurements under Real Conditions</td></tr><tr><td>SCOUT infantable canoe</td><td>GUMOTEX</td><td></td><td>https://www.gumotexboats.com/en/scout-standard#0000-044667-021-13/11C</td></tr><tr><td>Screw 10 x 170 mm with hexagonal nuts</td><td></td><td></td><td></td></tr><tr><td>Screw 4 x 15 mm with hexagonal nuts</td><td></td><td></td><td></td></tr><tr><td>Screw 4 x 15 mm with wing nuts</td><td></td><td></td><td></td></tr><tr><td>Snap hooks 50 x 5 mm</td><td></td><td></td><td></td></tr><tr><td>Steel Carabine hook 50 x 5 mm</td><td></td><td></td><td></td></tr><tr><td>Steel chain with wire diameter 3 mm, inside link 5.5 x 26 mm</td><td></td><td></td><td></td></tr><tr><td>Steel chain, 5 m</td><td></td><td></td><td></td></tr><tr><td>toothbrush</td><td></td><td></td><td></td></tr><tr><td>tweezer</td><td></td><td></td><td></td></tr><tr><td>Washer 10 x 50 mm</td><td></td><td></td><td></td></tr><tr><td>Washer 4 x 10 mm</td><td></td><td></td><td></td></tr><tr><td>Washer 4 x 10 mm</td><td></td><td></td><td></td></tr></tbody></table>

a low-cost method of measuring the in situ primary productivity of periphyton communities of lentic waters

Measuring the in situ primary productivity of periphyton during the growing season gradient can elucidate the quantitative effect of environmental drivers (mainly phosphorus concentration and light intensity) and species composition on primary productivity. Primary productivity is mainly driven by light intensity, temperature, availability of nutrients, and distribution of the ionic species of the carbonate system in the respective depths of the euphotic zone. It is a complex system that is very difficult to simulate in the laboratory. This cheap, transportable, and easy-to-build floating barge allows measuring the primary productivity accurately-directly under the actual natural conditions. The methodology is based on measuring the primary productivity in real time using noninvasive oxygen sensors integrated into tightly sealed glass jars, enabling online oxygen flux monitoring and providing new insights into metabolic activities. Detailed seasonal in situ measurements of gross primary productivity of microbial mats (or other benthic organisms) can improve current knowledge of the processes controlling primary productivity dynamics in lentic waters.

<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/64078/64078fig05.jpg" /> 
Figure 5: Net and gross ecosystem productivity of microbial mats during daylight. (A) Light bottle-net ecosystem productivity: time course data of net oxygen productivity of microbial mats from the light bottles. The oxygen concentration change in incubation bottles was measured after 1 h during daylight. Grey circles: bottles with samples of microbial mats. White circles: blank bottles with lake water only. (B) Dark bottle-respiration: the sum of autotrophic and heterotrophic respiration from the dark bottles. The change in oxygen concentration in incubation bottles was measured after 1 h during the daylight period. Grey circles: bottles with samples of microbial mats. White circles: blank bottles with lake water only. (C) Comparison of net ecosystem productivity, respiration, and gross ecosystem productivity: subtracting respiration rates from net ecosystem productivity rates results in gross ecosystem productivity of the microbial mats biomass. Abbreviations: NEP = net ecosystem productivity; RE = respiration; GEP = gross ecosystem productivity. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/64078/64078fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
A schematic for the experimental floating barge and incubation bottles setup is shown in Figure 1, Figure 2,&#160;and Figure 3. Figure 4 demonstrates how the experiment can be set up in the field. Figure5 shows the representative data set used to calculate the final net and gross ecosystem productivity. In Figure 5A, the change in O2 concentration (i.e., recalculated to mmol (O2) g(OM)-1) over time is shown for bottles with (denoted as &#34;sample&#34;) or without (denoted as &#34;control&#34;) the microbial mats that were exposed to light. The increase in O2 concentration is apparent in the control as well as in the sample. Nevertheless, the slope of increase is significantly higher in bottles with microbial mats. It shows that the signal of the microbial mat activity is detectable against the background.
The same applies to data from bottles kept in the dark, which are shown in Figure 5B. In this case, however, the slope of O2 concentration change over time in the control is not significantly different from zero. Figure 5C shows the products of data shown in Figure 5A,B, the net ecosystem productivity (NEP) and respiration (RE), respectively. By definition, the first must be positive and the latter negative. The data comply with the definition well. The sum of NPP and RE is then the gross ecosystem productivity (GEP). Notice that the error of GEP increases because it is the square root of the squared sums of errors calculated for NEP and RE. Every measurement event produces a series of O2 concentrations over daylight time for each of the monitored incubation bottles, which is the basis for estimating net and gross ecosystem productivity. Figure 5A shows the time course data of net oxygen productivity (photosynthesis and respiration) of microbial mats from the light bottles. Figure 5B shows the time course data of respiration of microbial mats from dark bottles. Subtracting respiration rates from net ecosystem productivity yields the rates of gross ecosystem productivity (Figure 5C). Figure 6 shows the results obtained from the practical application of the method in the field. The Gross ecosystem productivity of the periphytic community was measured in three post-mining lakes in the Czech Republic during the vegetation season.
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/64078/64870fig06v2.jpg" /> 
Figure 6. Gross ecosystem productivity of periphyton in three post-mining lakes. Example of seasonal variation of gross ecosystem productivity (GEP) of periphyton biomass in three post-mining lakes in the Czech Republic (Milada, Most, Medard). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/64078/64870fig06largev2.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td></td>
			<td></td>
			<td>Estimate</td>
			<td></td>
		</tr>
		<tr>
			<td>Experimental unit</td>
			<td>O2 flux proxy</td>
			<td>&#181;mol O2 g OM-1 h-1</td>
			<td>SE</td>
		</tr>
		<tr>
			<td>Light bottle</td>
			<td>Net ecosystem productivity (NEP)</td>
			<td>38.6</td>
			<td>1.98</td>
		</tr>
		<tr>
			<td>Dark bottle</td>
			<td>Ecosystem respiration (RE)</td>
			<td>-18.1</td>
			<td>2.3</td>
		</tr>
		<tr>
			<td colspan="2">Gross ecosystem productivity (GEP)</td>
			<td>53.8</td>
			<td>13.6</td>
		</tr>
	</tbody>
</table>
Table 1: A summary table of results. The results of the linear regression calculated as mean and standard error. Average estimation values from five replicates of dark and light incubation bottles. SE = standard error.

Watch this Scientific Journal Video about A Low-Cost Method of Measuring the In Situ Primary Productivity of Periphyton Communities of Lentic Waters at JoVE.com

A Low-Cost Method of Measuring the In Situ Primary Productivity of Periphyton Communities of Lentic Waters

Presented here is a cost-effective and transportable method/facility for measuring the primary productivity of microbial mats under actual in situ environmental temperature and light conditions. The experimental setup is based on widely available materials and can be used under various conditions while offering the advantages of laboratory-based models.

A Low-Cost Method of Measuring the In Situ Primary Productivity of Periphyton Communities of Lentic Waters

Measuring the in situ primary productivity of periphyton during the growing season gradient can elucidate the quantitative effect of environmental drivers ...

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Nanyang Technological University

Research

JoVE Journal

Environment

1.8K Views.  Institute of Hydrobiology. Gedetailleerde in-situ meting van de bruto productiviteit van periphyton of een watermicro-organisme op een geschikte locatie kan de huidige kennis van de processen die de primaire productiviteitsdynamiek in lenige wateren regelen, verbeteren. De belangrijkste verbeteringen in de voorgestelde methoden zijn niet-invasiviteit, kosteneffectiviteit en de mogelijkheid van gelijktijdige metingen op verschillende locaties. Het maakt ook het verzamelen van grote gegevensassets mogelijk zonder extra k...

Video: A Low-Cost Method of Measuring the In Situ Primary Productivity of Periphyton Communities of Lentic Waters

Laboratory-determined Phosphorus Flux from Lake Sediments as a Measure of Internal Phosphorus Loading

Eutrophication is a water quality issue in lakes worldwide, and there is a critical need to identify and control nutrient sources. Internal phosphorus (P) loading from lake sediments can account for a substantial portion of the total P load in eutrophic, and some mesotrophic, lakes. Laboratory determination of P release rates from sediment cores is one approach for determining the role of internal P loading and guiding management decisions. Two principal alternatives to experimental determination of sediment P release exist for estimating internal load: in situ measurements of changes in hypolimnetic P over time and P mass balance. The experimental approach using laboratory-based sediment incubations to quantify internal P load is a direct method, making it a valuable tool for lake management and restoration.
Laboratory incubations of sediment cores can help determine the relative importance of internal vs. external P loads, as well as be used to answer a variety of lake management and research questions. We illustrate the use of sediment core incubations to assess the effectiveness of an aluminum sulfate (alum) treatment for reducing sediment P release. Other research questions that can be investigated using this approach include the effects of sediment resuspension and bioturbation on P release.
The approach also has limitations. Assumptions must be made with respect to: extrapolating results from sediment cores to the entire lake; deciding over what time periods to measure nutrient release; and addressing possible core tube artifacts. A comprehensive dissolved oxygen monitoring strategy to assess temporal and spatial redox status in the lake provides greater confidence in annual P loads estimated from sediment core incubations.

Lake eutrophication is a water quality issue worldwide, making the need to identify and control nutrient sources critical. Laboratory determination of phosphorus release rates from sediment cores is a valuable approach for determining the role of internal phosphorus loading and guiding management decisions.

Eutrophication is a water quality issue in lakes worldwide, and there is a critical need to identify and control nutrient sources. Internal phosphorus (P) ...

Integrated Field Lysimetry and Porewater Sampling for Evaluation of Chemical Mobility in Soils and Established Vegetation

Potentially toxic chemicals are routinely applied to land to meet growing demands on waste management and food production, but the fate of these chemicals is often not well understood. Here we demonstrate an integrated field lysimetry and porewater sampling method for evaluating the mobility of chemicals applied to soils and established vegetation. Lysimeters, open columns made of metal or plastic, are driven into bareground or vegetated soils. Porewater samplers, which are commercially available and use vacuum to collect percolating soil water, are installed at predetermined depths within the lysimeters. At prearranged times following chemical application to experimental plots, porewater is collected, and lysimeters, containing soil and vegetation, are exhumed. By analyzing chemical concentrations in the lysimeter soil, vegetation, and porewater, downward leaching rates, soil retention capacities, and plant uptake for the chemical of interest may be quantified. Because field lysimetry and porewater sampling are conducted under natural environmental conditions and with minimal soil disturbance, derived results project real-case scenarios and provide valuable information for chemical management. As chemicals are increasingly applied to land worldwide, the described techniques may be utilized to determine whether applied chemicals pose adverse effects to human health or the environment.

Field lysimetry and porewater sampling allow researchers to evaluate the fate of chemicals applied to soils and established vegetation. The goal of this protocol is to demonstrate how to install required instrumentation and collect samples for chemical analysis during integrated field lysimetry and porewater sampling experiments.

Potentially toxic chemicals are routinely applied to land to meet growing demands on waste management and food production, but the fate of these chemicals ...

A Small Volume Bioassay to Assess Bacterial/Phytoplankton Co-culture Using WATER-Pulse-Amplitude-Modulated (WATER-PAM) Fluorometry

Conventional methods for experimental manipulation of microalgae have employed large volumes of culture (20 ml to 5 L), so that the culture can be subsampled throughout the experiment1&#8211;7. Subsampling of large volumes can be problematic for several reasons: 1) it causes variation in the total volume and the surface area:volume ratio of the culture during the experiment; 2) pseudo-replication (i.e., replicate samples from the same treatment flask8) is often employed rather than true replicates (i.e., sampling from replicate treatments); 3) the duration of the experiment is limited by the total volume; and 4) axenic cultures or the usual bacterial microbiota are difficult to maintain during long-term experiments as contamination commonly occurs during subsampling.
The use of microtiter plates enables 1 ml culture volumes to be used for each replicate, with up to 48 separate treatments within a 12.65 x 8.5 x 2.2 cm plate, thereby decreasing the experimental volume and allowing for extensive replication without subsampling any treatment. Additionally, this technique can be modified to fit a variety of experimental formats including: bacterial-algal co-cultures, algal physiology tests, and toxin screening9&#8211;11. Individual wells with an alga, bacterium and/or co-cultures can be sampled for numerous laboratory procedures including, but not limited to: WATER-Pulse-Amplitude-Modulated (WATER-PAM) fluorometry, microscopy, bacterial colony forming unit (cfu) counts and flow cytometry. The combination of the microtiter plate format and WATER-PAM fluorometry allows for multiple rapid measurements of photochemical yield and other photochemical parameters with low variability between samples, high reproducibility and avoids the many pitfalls of subsampling a carboy or conical flask over the course of an experiment.

A Small Volume Bioassay to Assess Bacterial/Phytoplankton Co-culture Using WATER-Pulse-Amplitude-Modulated WATER-PAM Fluorometry

The goal of this procedure is to demonstrate the reproducibility and adaptability of using a microtiter plate format for microalgal screening. This rapid screen combines WATER-Pulse-Amplitude-Modulated (WATER-PAM) fluorometry to measure photosynthetic yield as an indicator of Photosystem II (PSII) health with small volume bacterial-algal co-cultures.

Conventional methods for experimental manipulation of microalgae have employed large volumes of culture (20 ml to 5 L), so that the culture can be ...

VacuSIP, an Improved InEx Method for In Situ Measurement of Particulate and Dissolved Compounds Processed by Active Suspension Feeders

Benthic suspension feeders play essential roles in the functioning of marine ecosystems. By filtering large volumes of water, removing plankton and detritus, and excreting particulate and dissolved compounds, they serve as important agents for benthic-pelagic coupling. Accurately measuring the compounds removed and excreted by suspension feeders (such as sponges, ascidians, polychaetes, bivalves) is crucial for the study of their physiology, metabolism, and feeding ecology, and is fundamental to determine the ecological relevance of the nutrient fluxes mediated by these organisms. However, the assessment of the rate by which suspension feeders process particulate and dissolved compounds in nature is restricted by the limitations of the currently available methodologies. Our goal was to develop a simple, reliable, and non-intrusive method that would allow clean and controlled water sampling from a specific point, such as the excurrent aperture of benthic suspension feeders, in situ. Our method allows simultaneous sampling of inhaled and exhaled water of the studied organism by using minute tubes installed on a custom-built manipulator device and carefully positioned inside the exhalant orifice of the sampled organism. Piercing a septum on the collecting vessel with a syringe needle attached to the distal end of each tube allows the external pressure to slowly force the sampled water into the vessel through the sampling tube. The slow and controlled sampling rate allows integrating the inherent patchiness in the water while ensuring contamination free sampling. We provide recommendations for the most suitable filtering devices, collection vessel, and storing procedures for the analyses of different particulate and dissolved compounds. The VacuSIP system offers a reliable method for the quantification of undisturbed suspension feeder metabolism in natural conditions that is cheap and easy to learn and apply to assess the physiology and functional role of filter feeders in different ecosystems.

VacuSIP, an Improved InEx Method for In Situ Measurement of Particulate and Dissolved Compounds Processed by Active Suspension Feeders

We introduce the VacuSIP, a simple, non-intrusive, and reliable method for clean and accurate point sampling of water. The system was developed and evaluated for the simultaneous collection of the water inhaled and exhaled by benthic suspension feeders in situ, to cleanly measure removal and excretion of particulate and dissolved compounds.

Benthic suspension feeders play essential roles in the functioning of marine ecosystems. By filtering large volumes of water, removing plankton and ...

Characterization of Aquatic Biofilms with Flow Cytometry

Biofilms are dynamic consortia of microorganism that play a key role in freshwater ecosystems. By changing their community structure, biofilms respond quickly to environmental changes and can be thus used as indicators of water quality. Currently, biofilm assessment is mostly based on integrative and functional endpoints, such as photosynthetic or respiratory activity, which do not provide information on the biofilm community structure. Flow cytometry and computational visualization offer an alternative, sensitive, and easy-to-use method for assessment of the community composition, particularly of the photoautotrophic part of freshwater biofilms. It requires only basic sample preparation, after which the entire sample is run through the flow cytometer. The single-cell optical and fluorescent information is used for computational visualization and biological interpretation. Its main advantages over other methods are the speed of analysis and the high-information-content nature. Flow cytometry provides information on several cellular and biofilm traits in a single measurement: particle size, density, pigment content, abiotic content in the biofilm, and coarse taxonomic information. However, it does not provide information on biofilm composition on the species level. We see high potential in the use of the method for environmental monitoring of aquatic ecosystems and as an initial biofilm evaluation step that informs downstream detailed investigations by complementary and more detailed methods.

Flow cytometry in combination with visual clustering offers an easy-to-use and fast method for studying aquatic biofilms. It can be used for biofilm characterization, detection of changes in biofilm community structure, and detection of abiotic particles embedded in the biofilm.

Biofilms are dynamic consortia of microorganism that play a key role in freshwater ecosystems. By changing their community structure, biofilms respond ...

BtM, a Low-cost Open-source Datalogger to Estimate the Water Content of Nonvascular Cryptogams

Communities of nonvascular cryptogams, such as mosses or lichens, are an important part of the Earth's biodiversity, contributing to the regulation of the carbon and nitrogen cycles in many ecosystems. Being poikilohydric organisms, they do not actively control their internal water content and need a humid environment to activate their metabolism. Therefore, studying water relationships of nonvascular cryptogams is crucial to understand both their diversity patterns and their functions in the ecosystems. We present the BtM datalogger, a low-cost open-source platform for the study of the water content of nonvascular cryptogams. The datalogger is designed to measure ambient temperature, humidity, and conductance from up to eight samples simultaneously. We provide a design for a printed circuit board (PCB), a detailed protocol to assemble the components, and the required source code. All this makes the assembly of the BtM datalogger accessible to any research group, even to those without previous specialized knowledge. Therefore, the design presented here has the potential to help popularize the use of this type of device among ecologists and field biologists.

We present a simple and cost-effective method to build an open-source datalogger that measures the conductance of nonvascular cryptogams together with the environmental temperature and humidity. We describe the hardware design of the datalogger and provide step-by-step assembly instructions, the list of required open-source logging software, the code to run the datalogger, and a calibration protocol.

Communities of nonvascular cryptogams, such as mosses or lichens, are an important part of the Earth's biodiversity, contributing to the regulation of the ...

Laser-Induced Fluorescence Emission (L.I.F.E.) as Novel Non-Invasive Tool for In-Situ Measurements of Biomarkers in Cryospheric Habitats

Global warming affects microbial communities in a variety of ecosystems, especially cryospheric habitats. However, little is known about microbial-mediated carbon fluxes in extreme environments. Hence, the methodology of sample acquisition described in the very few studies available implies two major problems: A) high resolution data require a large number of samples, which is difficult to obtain in remote areas; B) unavoidable sample manipulation such as cutting, sawing, and melting of ice cores that leads to a misunderstanding of in situ conditions. In this study, a prototype device that requires neither sample preparation nor sample destruction is presented. The device can be used for in situ measurements with a high spectral and spatial resolution in terrestrial and ice ecosystems and is based on the Laser-Induced Fluorescence Emission (L.I.F.E.) technique. Photoautotrophic supraglacial communities can be identified by the detection of L.I.F.E. signatures in photopigments. The L.I.F.E. instrument calibration for the porphyrin derivates chlorophylla (chla) (405 nm laser excitation) and B-phycoerythrin (B-PE) (532 nm laser excitation) is demonstrated. For the validation of this methodology, L.I.F.E. data were ratified by a conventional method for chla quantification that involved pigment extraction and subsequent absorption spectroscopy. The prototype applicability in the field was proven in extreme polar environments. Further testing on terrestrial habitats took place during Mars analog simulations in the Moroccan dessert and on an Austrian rock glacier. The L.I.F.E. instrument enables high resolution scans of large areas with acceptable operation logistics and contributes to a better understanding of the ecological potential of supraglacial communities in the context of global change.

Laser-Induced Fluorescence Emission L.I.F.E. as Novel Non-Invasive Tool for In-Situ Measurements of Biomarkers in Cryospheric Habitats

Carbon fluxes in the cryosphere are hardly assessed yet but are crucial regarding climate change. Here we show a novel prototype device that captures the phototrophic potential in supraglacial environments based on laser-induced fluorescence emission (L.I.F.E.) technology offering high spectral and spatial resolution data under in situ conditions.

Global warming affects microbial communities in a variety of ecosystems, especially cryospheric habitats. However, little is known about ...

A Practical and Low-Cost Setup to Measure Methane in Aqueous Samples

Measuring Dissolved Methane in Aquatic Ecosystems Using An Optical Spectroscopy Gas Analyzer

Measuring greenhouse gas (GHG) fluxes and pools in ecosystems are becoming increasingly common in ecological studies due to their relevance to climate change. With it, the need for analytical platforms adaptable to measuring different pools and fluxes within research groups also grows. This study aims to develop a procedure to use portable optical spectroscopy-based gas analyzers, originally designed and marketed for gas flux measurements, to measure GHG concentrations in aqueous samples. The protocol involves the traditional headspace equilibration technique followed by the injection of a headspace gas subsample into a chamber connected through a closed loop to the inlet and outlet ports of the gas analyzer. The chamber is fabricated from a generic mason jar and simple laboratory supplies, and it is an ideal solution for samples that may require pre-injection dilution. Methane concentrations measured with the chamber are tightly correlated (r2 &#62; 0.98) with concentrations determined separately through gas chromatography-flame ionization detection (GC-FID) on subsamples from the same vials. The procedure is particularly relevant for field studies in remote areas where chromatography equipment and supplies are not readily available, offering a practical, cheaper, and more efficient solution for measuring methane and other dissolved greenhouse gas concentrations in aquatic systems.

This study demonstrates an approach to measure methane gas concentrations in aqueous samples using portable optical analyzers coupled to an injection chamber in a closed loop. The results are similar to conventional gas chromatography, presenting a practical and low-cost alternative particularly suitable for remote field studies.

Measuring greenhouse gas (GHG) fluxes and pools in ecosystems are becoming increasingly common in ecological studies due to their relevance to climate ...

Linking Predation Risk, Herbivore Physiological Stress and Microbial Decomposition of Plant Litter

The quantity and quality of detritus entering the soil determines the rate of decomposition by microbial communities as well as recycle rates of nitrogen (N) and carbon (C) sequestration1,2. Plant litter comprises the majority of detritus3, and so it is assumed that decomposition is only marginally influenced by biomass inputs from animals such as herbivores and carnivores4,5. However, carnivores may influence microbial decomposition of plant litter via a chain of interactions in which predation risk alters the physiology of their herbivore prey that in turn alters soil microbial functioning when the herbivore carcasses are decomposed6. A physiological stress response by herbivores to the risk of predation can change the C:N elemental composition of herbivore biomass7,8,9 because stress from predation risk increases herbivore basal energy demands that in nutrient-limited systems forces herbivores to shift their consumption from N-rich resources to support growth and reproduction to C-rich carbohydrate resources to support heightened metabolism6. Herbivores have limited ability to store excess nutrients, so stressed herbivores excrete N as they increase carbohydrate-C consumption7. Ultimately, prey stressed by predation risk increase their body C:N ratio7,10, making them poorer quality resources for the soil microbial pool likely due to lower availability of labile N for microbial enzyme production6. Thus, decomposition of carcasses of stressed herbivores has a priming effect on the functioning of microbial communities that decreases subsequent ability to of microbes to decompose plant litter6,10,11.
We present the methodology to evaluate linkages between predation risk and litter decomposition by soil microbes. We describe how to: induce stress in herbivores from predation risk; measure those stress responses, and measure the consequences on microbial decomposition. We use insights from a model grassland ecosystem comprising the hunting spider predator (Pisuarina mira), a dominant grasshopper herbivore (Melanoplus femurrubrum),and a variety of grass and forb plants9.

We present methods to evaluate how predation risk can alter the chemical quality of herbivore prey by inducing dietary changes to meet demands of heightened stress, and how the decomposition of carcasses from these stressed herbivores slows subsequent plant litter decomposition by soil microbes.

The quantity and quality of detritus entering the soil  determines the rate of decomposition by microbial communities as well as  recycle rates of ...

Biology

Operation of Laboratory Photobioreactors with Online Growth Measurements and Customizable Light Regimes

The laboratory study of microalgae can be experimentally challenging. In addition to the cultivation requirements of non-photosynthetic microorganisms, phototrophs also require illumination. Routinely, researchers seek to provide custom light supplies, i.e., vary the light intensity and time over which it is delivered. Such flexibility is difficult with standard benchtop lights. Usually, cultivation studies also require growth comparisons between experimental treatments. Frequently, growth is assessed over an extended duration, e.g., multiple times a day over a week-long trial. Manual measurements can be time-consuming and lack data resolution. Therefore, photobioreactors (PBRs) with automatic growth monitoring and customizable light supply are useful for replicated experiments with multiple treatments. The current work presents the design, construction, and operation of laboratory PBRs. The materials are easily sourced and relatively inexpensive. The design could be duplicated with moderate skill. Each structure has a footprint of ~40 cm2 and hosts three 1 L glass bottles for triplicate replication. Bottles rest upon platforms containing magnetic stirrers and are arranged vertically within a 1 m high and 15 cm diameter polyvinyl chloride (PVC) pipe. The pipe interior is lined with light-emitting diodes (LEDs). These LEDs produce continuous light intensities from 0&#8211;2400 &#181;mol photons m-2 s-1 of photosynthetically active radiation (PAR). Users design a custom lighting program. The light intensity can be adjusted each second or held constant for longer durations. Oxygen produced from photosynthesis exits each bottle via a one-way volumetric gas sensor. Software is used to record gas sensor data. The amount of oxygen produced can be correlated to biomass growth. If biomass samples are required, a syringe can be used to extract culture. The method is suited for microalgae grown with bicarbonate as the carbon source. These PBRs are valuable to a laboratory that requires replicated experiments, light regime flexibility, and continuous high-resolution growth data.

This publication describes the design of laboratory photobioreactors (PBRs) with customizable light regimes. The growth of cyanobacteria or microalgae, using bicarbonate as their carbon source, is monitored continuously by measuring volumetric oxygen production. These PBRs facilitate rapid, replicated laboratory growth comparisons with little user intervention during experiments.

The laboratory study of microalgae can be experimentally challenging. In addition to the cultivation requirements of non-photosynthetic microorganisms, ...

A Low-Cost Method of Measuring the In Situ Primary Productivity of Periphyton Communities of Lentic Waters

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