The authors gratefully acknowledge funding from the Natural Sciences and Engineering Research Council of Canada (05448, 165831 and 213411).

1. Preparing Cyanobacterial Cultures
<ol>
	<li>Preparing inoculation cultures using solid media
	<ol>
		<li>Obtain axenic cyanobacterial cells from the American Type Culture Collection or Pasteur Culture Collection. For example, the unicellular, marine Synechococcus sp. PCC 7002 was obtained from the Pasteur Culture Collection, it will be referred to hereafter as Synechococcus.</li>
		<li>Maintain Synechococcus cells on plates containing solid media (A+ liquid media14 and 1.5% agar15). These will be referred to hereafter as inoculation cultures.</li>
		<li>Incubate the plates at 32 &#177; 2 &#176;C with continuous light provided at 30&#8211;50 &#181;mol photons m-2 s-1,15,18.</li>
		<li>Repeat steps 1.1&#8211;1.3 and replace A+ media with BG-1116 in step 1.1.2 for freshwater cyanobacterial species, such as Synechocystis sp. PCC 680316,17, referred to hereafter as Synechocystis. Refer to Tables 1&#8211;4 for the composition of A+ and BG-11 media.</li>
	</ol>
	</li>
	<li>Preparing liquid cultures: Each tank experiment requires one 400 mL cyanobacterial culture.
	<ol>
		<li>Collect one 250 mL and two 1 L heat-resistant Erlenmeyer flask.</li>
		<li>Rinse each flask with a 4 M hydrochloric acid (HCl) solution (30 mL of HCl solution for each flask), followed by distilled water. 
		Caution: Hydrochloric acid is corrosive and toxic. Ensure that a lab coat, safety goggles, and acid-resistant gloves are worn while using the 4 M HCl solution. Perform this step in a laboratory sink and follow local regulations regarding the disposal of the 4 M HCl solution</li>
		<li>Fill the 250 mL flask with 50 mL of liquid media (A+ or BG-11). Fill one 1 L flask with 400 mL of liquid media (A+ or BG-11) and the other 1 L flask with 600 mL of media (A+ or BG-11).</li>
		<li>Seal each flask with a gas permeable foam stopper and cover the foam stopper and flask neck with tinfoil.</li>
		<li>Label and autoclave the flasks at 121 &#176;C and 100 kPa for 25 min. Allow the flasks to cool to room temperature.</li>
		<li>Prepare a laminar flow hood by sterilizing the surface with 70% alcohol.</li>
		<li>Collect a wire inoculation loop, a Bunsen burner, the cooled 50 mL liquid media flask (from step 1.2.5), and the inoculation culture (from step 1.1). Place these items in the sterilized flow hood.</li>
		<li>Sterilize the wire inoculation loop briefly using the flame of a Bunsen burner and allow it to cool down.</li>
		<li>Drag the cool loop along the surface of the inoculation culture to collect cyanobacterial cells.</li>
		<li>Remove the foam stopper and foil cap from the 250 mL flask (without touching the foam stopper) and sterilize the rim of the Erlenmeyer flask by passing it briefly through the flame.</li>
		<li>Tilt the flask so the media is near the neck of the flask and insert the inoculation loop coated with cells into the flask. Once the loop is submerged into the media, stir the inoculation loop to remove the cells.</li>
		<li>Remove the inoculation loop and re-sterilize the lip of the flask using the Bunsen burner flame.</li>
		<li>Replace the foam stopper and foil cap onto the Erlenmeyer flask (without touching the foam stopper).</li>
		<li>Sterilize the inoculation loop briefly within the Bunsen burner flame.</li>
		<li>Prepare a growth room (referred to as the growth chamber) where the temperature is maintained at 30 &#176;C15,18&#160;and light intensity is 30&#8211;50 &#181;mol photons m-2 s-1,15,18. Humidity is not actively controlled.</li>
		<li>Allow the inoculated 50 mL culture to incubate by shaking at 150 rpm in the growth chamber, with a maintained temperature of 30 &#176;C. Keep the mouth of the flask covered during the agitation to control evaporative losses and prevent contamination.</li>
		<li>Allow the cyanobacterial culture(s) to grow for 96 h. A successful culture should appear bright green and have an optical density (OD) at 750 nm of 0.4&#8211;0.6.</li>
		<li>Once the culture has grown sufficiently, place the 50 mL culture and the 1 L flask containing 400 mL of liquid media (from step 1.2.5) in the laminar flow hood. Ensure the flow hood is sterilized following step 1.2.6.</li>
		<li>Repeat step 1.2.10 for both flasks within a laminar flow hood.</li>
		<li>Pour the 50 mL culture into the 400 mL of media in the 1 L flask and re-sterilize the lip of the 1 L flask using the Bunsen burner flame.</li>
		<li>In the place of the foam stopper and foil cap, attach a sterile bubbling apparatus consisting of a foam stopper, pipette, plastic tubing, in-line filter, and foil cover to the mouth of the flask.</li>
		<li>Agitate the 1 L flask at 150 rpm at 30 &#176;C with the bubbler apparatus attached to an air pump, which circulates a humidified air mixture through the solution15,18 at 300 mL/min.</li>
		<li>Allow the culture(s) to grow within the growth chamber at 30 &#176;C for 120&#8211;168 h. A successful culture should appear bright green and have an OD750 nm of 0.4&#8211;0.6.</li>
		<li>Repeat step 1.2 to produce a control culture, which will not be mixed with the clay.</li>
		<li>Experiments using dead biomass require an additional step. Autoclave the 1 L flask culture for 60 min at 121 &#176;C and allow it to cool to room temperature.</li>
	</ol>
	</li>
</ol>
2. Experimental Set Up
<ol>
	<li>Place a rectangular acrylic tank (length of 20 cm, width of 5.1 cm, and height of 30 cm) in front of a bank of fluorescent lights (nine T8 bulbs, at least 2,600 lumens; Figure 1)19.</li>
	<li>Place a translucent, white plastic sheet between the bank of lights and the tank to diffuse the light, which shines through the tank. This set-up was adapted from Sutherland et al.19</li>
	<li>Mark the acrylic tank at a height of 10 cm, measured vertically from the base. All measurements should be done at this height in the water column to ensure the consistency of all sampling.</li>
	<li>Place a camera on a tripod, 1 m in front of the tank to record each experiment. A video camera is recommended as images can be extracted from video files, and by using mathematical modeling software, video files can be used to model settling dynamics19.</li>
	<li>Place a black cloth over the light bank, tank, and camera to shield the experiment from outside light sources (Figure 1).</li>
</ol>
3. Flocculation Experimental Protocol
<ol>
	<li>Collect the 400 mL culture (step 1.2), a large graduated cylinder, and 600 mL of extra growth media in a 1 L flask (step 1.2.5).</li>
	<li>Dilute the 400 mL culture (initial cell concentration of 4&#8211;6 mg/mL) with additional growth media (A+ or BG-11) to final volume of 1 L using the graduated cylinder. Add this solution to the acrylic tank.</li>
	<li>Prepare 2 mL microfuge tubes by labeling and placing them in a convenient location near the acrylic tank. Measure 50 g of clay for the experiment.</li>
	<li>Begin video recording. In the case of multiple experiments, a sign with the experiment identifier can be temporarily held before the camera.</li>
	<li>Using a pipette, take a 1 mL sample and place it in a labeled microfuge tube. Use this sample to determine the initial cyanobacterial cell count by measuring the OD750 nm values. Take all samples in triplicate to ensure accurate results.</li>
	<li>Quickly pour the clay (50 g) into the tank with vigorous agitation using a stirring stick for 10&#8211;15 s.</li>
	<li>Start the timer and take the initial 1 mL sample for Chl a determination using a P1000 pipette (sample Time0).</li>
	<li>Take additional samples at appropriate time intervals (e.g., 1 min, 2 min, 3 min, 4 min, 5 min, 10 min, 15 min, 30 min, 60 min, 180 min, and 240 min).</li>
	<li>Once the experiment is finished, save the video file, turn off the video camera, and process all samples for Chl a determination (step 4).</li>
	<li>Autoclave the remaining clay/cyanobacterial solution. Depending on cyanobacterial species, local regulations, and lab policy, dispose of the solution using appropriate methods. Clean the tank with soap and water, rinse it with distilled water, and air dry it.</li>
	<li>Prepare a control experiment with no clay added. Take samples at the same time points. This data can be compared to the other experimental data to confirm that settling rates are enhanced due to the flocculation with clay, not a result of natural settling.</li>
	<li>If dead biomass is used during the experiment, use the same experimental procedure. However, the disposal procedure of the solution does not require autoclaving.</li>
</ol>
4. Sample Processing and Evaluation
<ol>
	<li>Determine Chl a concentration in each cell sample using a procedure modified from Owttrim16. Once collected, pellet the cells from each sample for 3 min at 13,000 x g using a microcentrifuge at room temperature.</li>
	<li>Remove excess media using a P1000 pipette and add 1 mL of 100% methanol (20 &#176;C). Vortex the sample at full speed for 1 min to resuspend the cell pellet. 
	CAUTION: Methanol is toxic and flammable. Wear gloves and safety glasses, and keep methanol away from ignition sources.</li>
	<li>Incubate the samples at -20 &#176;C for 24 h to facilitate the extraction of Chl a.</li>
	<li>After incubation, pellet the cellular debris as described in 4.1, transfer the green supernatant to a cuvette using a pipette, and place the cuvette into the spectrophotometer. Allow the sample to rest in the spectrophotometer for 1 min to ensure that any remaining clay within the sample settles and does not interfere with the measurement.</li>
	<li>Determine Chl&#160;a&#160;concentration spectrophotometrically by measuring absorbance at 665 nm and 652 nm, and OD at 750 nm. The Chl&#160;a concentration is determined using the formula Chl&#160;a&#160;= 16.29 x (A665 - OD750) - 8.54 x (A652 - OD750) 20. Chl a concentration is used as a proxy for cell concentration. Additionally, a conversion factor of 7.4 x 1010 cells/L = 10g/L21 can be used to calculate the cell concentration in grams of some cyanobacterial species.</li>
	<li>Plot calculated Chl a values versus time.</li>
</ol>

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The authors have nothing to disclose.

Flocculation catalyzed by cyanobacterial cell-clay interaction has attracted a lot of interest in the fields of ecology and engineering2,3,4,5,6,7,8,9,10,11,1...

Using present environmental conditions and processes to infer past depositional mechanisms has long been an underpinning of sedimentology. While modern depositional analogues, such as the Black Sea, have been used to understand the deposition of organic-rich, fine-grained deposits, laboratory experiments have the potential to shed additional light on the origin of shale deposits. One area of inquiry in the genesis of black shales is the deposition rate and mechanism of original formation. Traditionally, it has been hypothesized that black shales formed in environments where the sedimentation rate, primary productivity, and organic matter respiration rates promote the preservation of organic matter in the sediment1,2,3. However, the role of cyanobacterial and clay flocculation has largely remained unconsidered. This mechanism of flocculation would allow for rapid deposition of organic-rich, fine-grained sediments to occur, and does not necessitate low-oxygen. Considering this premise, this protocol has two goals: 1) measure the sedimentation rate of cyanobacterial/clay floccules, and 2) visualize the sedimentation process in real time. This methodology, in addition to geochemical analysis, has been used to demonstrate that cyanobacterial/clay flocculation may in fact be an important mechanism for shale formation1. While originally intended for modelling shale deposition, this method is applicable to other disciplines such as biology and environmental remediation where the influence of clay input on bacterial metabolism and population need to be measured.
Numerous studies have been conducted to observe the flocculation of cyanobacteria and clay, for mitigating harmful algal blooms2,3,4,5,6,7,8,9,10,11,12. However, while measuring cell concentration over time, these studies have not applied cyanobacteria/clay flocculation to modelling the deposition of the rock record. As such, these studies lack a visual component, which can be critical when modelling past sedimentological processes. Additionally, the majority of studies utilize cell-counting (e.g., Pan et al.11), which can be laborious. Our method, with recent advances in measuring cyanobacterial flocculation, determines the changes in cyanobacterial cell concentration by measuring chlorophyll a (Chl a) at discrete time intervals. Pairing Chl a measurement with visual data is a new approach, which can be used to infer depositional conditions. The images generated can also be used to calculate sedimentation rate after the work from Du et al.13. The combination of visual and numerical data strengthens the reliability of the results. Furthermore, we outline additional protocols allowing for the sedimentation of dead biomass and clay to also be observed. This is important when considering past sedimentological environments, where live and dead biomass may have co-occurred. Differences in the behavior of dead biomass during the flocculation (for example, decrease in flocculation rate) would potentially have sedimentological implications.

<table><tbody><tr><td>cyanobacteria (in this study: &lt;em&gt;Synechococcus&lt;/em&gt; sp. PCC 7002 and &lt;em&gt;Synechocystis&lt;/em&gt; sp. PCC 6803)</td><td>Pasteur Culture Collection</td><td>PCC 7002 or PCC 6803</td><td>used to inoculate the plates</td></tr><tr><td>agar</td><td>Thermo Scientific</td><td>CM0003</td><td>used to fill two petri dishes</td></tr><tr><td>Petri plates (standard bacteriology, 100 x 15 mm)</td><td>Sarstedt</td><td>82.1473.001</td><td>2 required</td></tr><tr><td>1 L heat resistant Erlenmeyer flask</td><td>Pyrex</td><td>4980-125</td><td>1 required</td></tr><tr><td>250 mL heat resistant Erlenmeyer flask</td><td>Pyrex</td><td>4980-250</td><td>1 required</td></tr><tr><td>Nichrome inoculating loop with handle</td><td>Fisher Scientific</td><td>14-956-103</td><td>1 required</td></tr><tr><td>tinfoil</td><td>Reynolds Wrap Aluminum Foil</td><td>89079-067</td><td>50 cm required; used to cover foam stopper and neck of erlenmeyer flasks</td></tr><tr><td>growth media (e.g. A+)</td><td></td><td></td><td>1050 mL required; produced using composition described in tables 1-4</td></tr><tr><td>Bunsen Burner</td><td>Fisher Scientific</td><td>S95941</td><td>1 required</td></tr><tr><td>plastic tubing</td><td>Fisher Scientific</td><td>S504591</td><td>1 m required; used to create the bubbling apparatus</td></tr><tr><td>sponge stopper</td><td>Jaece Industries Inc</td><td>14-127-40E</td><td>1 required; hole made in center for pipette; used for constructin the bubbling apparatus</td></tr><tr><td>acrylic sheet&amp;nbsp;</td><td>Home Depot</td><td>Optix clear acrylic sheet model # MC-102S</td><td>1 required; used to construct acrylic tank (20 x 30 x 5.1 cm)</td></tr><tr><td>clear waterproof silicone adhesive</td><td>Home Depot</td><td>Loctite clear silicone model # 908570</td><td>1 required; used to construct acrylic tank (20 x 30 x 5.1 cm)</td></tr><tr><td>camera or video recorder</td><td>Panasonic</td><td>HC-V770 HD camcorder</td><td>1 required</td></tr><tr><td>tripod</td><td>Magnus</td><td>VT-300</td><td>1 required</td></tr><tr><td>black cloth</td><td>primomart</td><td>&amp;nbsp;EAN 0726670162199; Part number 680254blacknappedfr</td><td>1 required; duvetyne light block-out cloth; approximatly 152 x 213 cm to cover tank experiment</td></tr><tr><td>heat resistant serological pipet</td><td>corning incorporated C708510</td><td>13-671-101G</td><td>1 required; used to create the bubbling apparatus</td></tr><tr><td>sample vials&amp;nbsp;</td><td>Dynalon</td><td>S30467</td><td>at least 12 (will vary with time interval chosen)</td></tr><tr><td>heat resistant glass pipette</td><td>Fisher Scientific</td><td>Corning Incorporated C708510, 13-671-101G</td><td>1 required; used to create the bubbling apparatus; Polystyrene serological pipet would also work, but should be connected to the tubing and stopper after the rest of the apparatus is autoclaved.</td></tr><tr><td>microcentrifuge</td><td>Eppendorf</td><td>22 62 120-3</td><td>&amp;nbsp;1 required;Comparable products may be used if capable of centrifuging 1.5 -2 mL microfuge tubes at 13,000 x g</td></tr><tr><td>vortex machine (Vortex-Genie 2)</td><td>Scientific Industries, Inc</td><td>SI-0236</td><td>1 required</td></tr><tr><td>100% methanol</td><td>Fisher Scientific</td><td>A412-500 SDS</td><td>at least 12 mL (1mL per sample) required; Caution: Flammable, toxic. Wear gloves and safety glasses. Do not use or store near ignition source. Alternate sources may be used.</td></tr><tr><td>cuvettes (1.6&amp;nbsp; mL, polystyrene)</td><td>Sarstedt</td><td>67.742</td><td>at least 12 required</td></tr><tr><td>spectrophotometer</td><td>Fisher Scientific</td><td>222-271600</td><td>1 required; Pharmacia Biotech Novaspec ll could also be used.</td></tr><tr><td>light bulbs</td><td>Home Depot</td><td>model # 451807; internet #205477895; store SKU #1001061538</td><td>6-8 bulbs required to provide light for the tank experiments</td></tr><tr><td>pipette (Pipetman Classic P1000</td><td>Gilson</td><td>F123602</td><td>used to collect samples</td></tr><tr><td>37 % Hydrochloric acid</td><td>Sigma-Aldrich</td><td>258148</td><td>Caution: Corrosive and toxic. Wear lab coat, safety glasses and acid-resistant gloves while using. Prepared to 4 N before use by dilution into deionized water in a chemical fumehood.</td></tr><tr><td>Foam stopper (small)</td><td>Canlab</td><td>T 1385</td><td></td></tr><tr><td>Foam stopper (large)</td><td>Canlab</td><td>T 1387</td><td>Requires some intact stoppers and some with a single hole through the centre</td></tr><tr><td>30 &amp;deg;C incubator/growth room with continuous illumination</td><td></td><td></td><td>1 required</td></tr><tr><td>70 % Ethanol</td><td>Fisher Scientific</td><td>BP8201500</td><td>30 mL&amp;nbsp; required;Caution: Toxic and flammable. Wear lab coat and safety glasses</td></tr><tr><td>hydrophobic air filter (Midisart 2000, 0.2 &amp;micro;m)</td><td>Sartorius</td><td>17805</td><td>1 required</td></tr><tr><td>clay (e.g. kaolin)</td><td>Fisher Scientific</td><td>MFCD00062311</td><td>at least 50 g required</td></tr><tr><td>microfuge tubes (2 mL, polypropylene)</td><td>Sarstedt</td><td>72.695.500</td><td>Comparable products may be used. At least 12 (will vary with time interval chosen)</td></tr><tr><td>1000 &amp;micro;L pipet tips</td><td>Sarstedt</td><td>70.762</td><td>1 required</td></tr></tbody></table>

determination of the settling rate of clay/cyanobacterial floccules

The mechanisms underpinning the deposition of fine-grained, organic-rich sediments are still largely debated. Specifically, the impact of the interaction of clay particles with reactive, planktonic cyanobacterial cells to the sedimentary record is under studied. This interaction is a potentially major contributor to shale depositional models. Within a lab setting, the flocculation and sedimentation rates of these materials can be examined and measured in a controlled environment. Here, we detail a protocol for measuring the sedimentation rate of cyanobacterial/clay mixtures. This methodology is demonstrated through the description of two sample experiments: the first uses kaolin (a dehydrated form of kaolinite) and Synechococcus sp. PCC 7002 (a marine coccoid cyanobacteria), and the second uses kaolin and Synechocystis sp. PCC 6803 (a freshwater coccoid cyanobacteria). Cyanobacterial cultures are mixed with varying amounts of clay within a specially designed tank apparatus optimized to allow continuous, real-time video and photographic recording. The sampling procedures are detailed as well as a post-collection protocol for precise measurement of chlorophyll a from which the concentration of cyanobacterial cells remaining in suspension can be determined. Through experimental replication, a profile is constructed that displays sedimentation rate.

When exposed to clay, cyanobacterial cells are brought out of suspension22. This is demonstrated in the representative results given here. To determine the effect of clay on cyanobacterial populations and to observe the sedimentation rates, two experiments were conducted during which Synechococcus and Synechocystis were exposed to 50 g/L kaolin clay (Table 5&#8211;6, Figure 2&#8211;3)...

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Determination of the Settling Rate of Clay/Cyanobacterial Floccules

The interaction and sedimentation of the clay and bacterial cells within the marine realm, observed in natural environments, can be best investigated in a controlled lab environment. Here, we describe a detailed protocol, which outlines a novel method for measuring the sedimentation rate of clay and cyanobacterial floccules.

The mechanisms underpinning the deposition of fine-grained, organic-rich sediments are still largely debated. Specifically, the impact of the interaction ...

determination-of-the-settling-rate-of-claycyanobacterial-floccules

Universidad San Sebastian

Research

JoVE Journal

Environment

6.9K Views.  University of Alberta. De interactie en sedimentatie van klei en bacteriële cellen binnen de mariene realm, waargenomen in de natuurlijke milieus, kunnen het beste worden onderzocht in een gecontroleerde testomgeving. Hier beschrijven we een gedetailleerd protocol, dat een nieuwe methode voor het meten van de snelheid van sedimentatie van klei en cyanobacteriën floccules schetst.

Video: Determination of the Settling Rate of Clay/Cyanobacterial Floccules

Laboratory Simulation of an Iron(II)-rich Precambrian Marine Upwelling System to Explore the Growth of Photosynthetic Bacteria

A conventional concept for the deposition of some Precambrian Banded Iron Formations (BIF) proceeds on the assumption that ferrous iron [Fe(II)] upwelling from hydrothermal sources in the Precambrian ocean was oxidized by molecular oxygen [O2] produced by cyanobacteria. The oldest BIFs, deposited prior to the Great Oxidation Event (GOE) at about 2.4 billion years (Gy) ago, could have formed by direct oxidation of Fe(II) by anoxygenic photoferrotrophs under anoxic conditions. As a method for testing the geochemical and mineralogical patterns that develop under different biological scenarios, we designed a 40 cm long vertical flow-through column to simulate an anoxic Fe(II)-rich marine upwelling system representative of an ancient ocean on a lab scale. The cylinder was packed with a porous glass bead matrix to stabilize the geochemical gradients, and liquid samples for iron quantification could be taken throughout the water column. Dissolved oxygen was detected non-invasively via optodes from the outside. Results from biotic experiments that involved upwelling fluxes of Fe(II) from the bottom, a distinct light gradient from top, and cyanobacteria present in the water column, show clear evidence for the formation of Fe(III) mineral precipitates and development of a chemocline between Fe(II) and O2. This column allows us to test hypotheses for the formation of the BIFs by culturing cyanobacteria (and in the future photoferrotrophs) under simulated marine Precambrian conditions. Furthermore we hypothesize that our column concept allows for the simulation of various chemical and physical environments &#8212; including shallow marine or lacustrine sediments.

Laboratory Simulation of an IronII-rich Precambrian Marine Upwelling System to Explore the Growth of Photosynthetic Bacteria

We simulated a Precambrian ferruginous marine upwelling system in a lab-scale vertical flow-through column. The goal was to understand how geochemical profiles of O2 and Fe(II) evolve as cyanobacteria produce O2. The results show the establishment of a chemocline due to Fe(II) oxidation by photosynthetically produced O2.

A conventional concept for the deposition of some Precambrian Banded Iron Formations (BIF) proceeds on the assumption that ferrous iron [Fe(II)] upwelling ...

Determination of the Glycogen Content in Cyanobacteria

Cyanobacteria accumulate glycogen as a major intracellular carbon and energy storage&#160;during photosynthesis. Recent developments in research have highlighted complex mechanisms of glycogen metabolism, including the diel cycle of biosynthesis and catabolism, redox regulation, and the involvement of non-coding RNA. At the same time, efforts are being made to redirect carbon from glycogen to desirable products in genetically engineered cyanobacteria to enhance product yields. Several methods are used to determine the glycogen contents in cyanobacteria, with variable accuracies and technical complexities. Here, we provide a detailed protocol for the reliable determination of the glycogen content in cyanobacteria that can be performed in a standard life science laboratory. The protocol entails the selective precipitation of glycogen from the cell lysate and the enzymatic depolymerization of glycogen to generate glucose monomers, which are detected by a glucose oxidase-peroxidase (GOD-POD) enzyme coupled assay. The method has been applied to Synechocystis sp. PCC 6803 and Synechococcus sp. PCC 7002, two model cyanobacterial species that are widely used in metabolic engineering. Moreover, the method successfully showed differences in the glycogen contents between the wildtype and mutants defective in regulatory elements or glycogen biosynthetic genes.

Here, we present a reliable and easy assay to measure the glycogen content in cyanobacterial cells. The procedure entails precipitation, selectable depolymerization, and the detection of glucose residues. This method is suitable for both wildtype and genetically engineered strains and can facilitate the metabolic engineering of cyanobacteria.

Cyanobacteria accumulate glycogen as a major intracellular carbon and energy storage&#160;during photosynthesis. Recent developments in research have ...

Biochemistry

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 ...

Combined Size and Density Fractionation of Soils for Investigations of Organo-Mineral Interactions

Combined size and density fractionation (CSDF) is a method used to physically separate soil into fractions differing in particle size and mineralogy. CSDF relies on sequential density separation and sedimentation steps to isolate (1) the free light fraction (uncomplexed organic matter), (2) the occluded light fraction (uncomplexed organic matter trapped in soil aggregates) and (3) a variable number of heavy fractions (soil minerals and their associated organic matter) differing in composition. Provided that the parameters of the CSDF (dispersion energy, density cut-offs, sedimentation time) are properly selected, the method yields heavy fractions of relatively homogeneous mineral composition. Each of these fractions is expected to have a different complexing ability towards organic matter, rendering this a useful method to isolate and study the nature of organo-mineral interactions. Combining density and particle size separation brings an improved resolution compared to simple size or density fractionation methods, allowing the separation of heavy components according to both mineralogy and size (related to surface area) criteria. As is the case for all physical fractionation methods, it may be considered as less disruptive or aggressive than chemically-based extraction methods. However, CSDF is a time-consuming method and furthermore, the quantity of material obtained in some fractions can be limiting for subsequent analysis. Following CSDF, the fractions may be analyzed for mineralogical composition, soil organic carbon concentration and organic matter chemistry. The method provides quantitative information about organic carbon distribution within a soil sample and brings light to the sorptive capacity of the different, naturally-occurring mineral phases, thus providing mechanistic information about the preferential nature of organo-mineral interactions in soils (i.e., which minerals, what type of organic matter).

Combined size and density fractionation (CSDF) is a method to physically separate soil into fractions differing in texture (particle size) and mineralogy (density). The purpose is to isolate fractions with different reactivities towards soil organic matter (SOM), in order to better understand organo-mineral interactions and SOM dynamics.

Combined size and density fractionation (CSDF) is a method used to physically separate soil into fractions differing in particle size and mineralogy. CSDF ...

Measurement of the Potential Rates of Dissimilatory Nitrate Reduction to Ammonium Based on 14NH4+/15NH4+ Analyses via Sequential Conversion to N2O

The importance of understanding the fate of nitrate (NO3&#8722;), which is the dominant N species transferred from terrestrial to aquatic ecosystems, has been increasing because global nitrogen loads have dramatically increased following industrialization. Dissimilatory nitrate reduction to ammonium (DNRA) and denitrification are both microbial processes that use NO3&#8722; for respiration. Compared to denitrification, quantitative determinations of the DNRA activity have been carried out only to a limited extent. This has led to an insufficient understanding of the importance of DNRA in NO3&#8722; transformations and the regulating factors of this process. The objective of this paper is to provide a detailed procedure for the measurement of the potential DNRA rate in environmental samples. In brief, the potential DNRA rate can be calculated from the 15N-labeled ammonium (15NH4+) accumulation rate in 15NO3&#8722; added incubation. The determination of the 14NH4+ and 15NH4+ concentrations described in this paper is comprised of the following steps. First, the NH4+ in the sample is extracted and trapped on an acidified glass filter as ammonium salt. Second, the trapped ammonium is eluted and oxidized to NO3&#8722; via persulfate oxidation. Third, the NO3&#8722; is converted to N2O via an N2O reductase deficient denitrifier. Finally, the converted N2O is analyzed using a previously developed quadrupole gas chromatography&#8211;mass spectrometry system. We applied this method to salt marsh sediments and calculated their potential DNRA rates, demonstrating that the proposed procedures allow a simple and more rapid determination compared to previously described methods.

Measurement of the Potential Rates of Dissimilatory Nitrate Reduction to Ammonium Based on 14NH4+/15NH4+ Analyses via Sequential Conversion to N2O

A series of methods to determine the potential DNRA rate based on 14NH4+/15NH4+ analyses is provided in detail. NH4+ is converted into N2O via several steps and analyzed using quadrupole gas chromatography&#8211;mass spectrometry.

The importance of understanding the fate of nitrate (NO3&#8722;), which is the dominant N species transferred from terrestrial to aquatic ecosystems, has ...

A Small-Scale Setup for Algal Toxicity Testing of Nanomaterials and Other Difficult Substances

Ecotoxicity data is a requirement for pre- and post-market registration of chemicals by European and international regulations (e.g., REACH). The algal toxicity test is frequently used in regulatory risk assessment of chemicals. In order to achieve high reliability and reproducibility the development of standardized guidelines is vital. For algal toxicity testing, the guidelines require stable and uniform conditions of parameters such as pH, temperature, carbon dioxide levels and light intensity. Nanomaterials and other so-called difficult substances can interfere with light causing a large variation in results obtained hampering their regulatory acceptance. To address these challenges, we have developed LEVITATT (LED Vertical Illumination Table for Algal Toxicity Tests). The setup utilizes LED illumination from below allowing for a homogenous light distribution and temperature control while also minimizing intra-sample shading. The setup optimizes the sample volume for biomass quantification and does at the same time ensure a sufficient influx of CO2 to support exponential growth of the algae. Additionally, the material of the test containers can be tailored to minimize adsorption and volatilization. When testing colored substances or particle suspensions, the use of LED lights also allows for increasing the light intensity without additional heat generation. The compact design and minimal equipment requirements increase the possibilities for implementation of the LEVITATT in a wide range of laboratories. While compliant with standardized ISO and OECD guidelines for algal toxicity testing, LEVITATT also showed a lower inter-sample variability for two reference substances (3,5-Dicholorophenol and K2Cr2O7) and three nanomaterials (ZnO, CeO2, and BaSO4) compared to Erlenmeyer flasks and microtiter plates.

We demonstrate algal toxicity testing for difficult substances (e.g., colored substances or nanomaterials) using a setup illuminated vertically with an LED.

Ecotoxicity data is a requirement for pre- and post-market registration of chemicals by European and international regulations (e.g., REACH). The algal ...

Methods for Characterizing the Co-development of Biofilm and Habitat Heterogeneity

Biofilms are surface-attached microbial communities that have complex structures and produce significant spatial heterogeneities. Biofilm development is strongly regulated by the surrounding flow and nutritional environment. Biofilm growth also increases the heterogeneity of the local microenvironment by generating complex flow fields and solute transport patterns. To investigate the development of heterogeneity in biofilms and interactions between biofilms and their local micro-habitat, we grew mono-species biofilms of Pseudomonas aeruginosa and dual-species biofilms of P. aeruginosa and Escherichia coli under nutritional gradients in a microfluidic flow cell. We provide detailed protocols for creating nutrient gradients within the flow cell and for growing and visualizing biofilm development under these conditions. We also present protocols for a series of optical methods to quantify spatial patterns in biofilm structure, flow distributions over biofilms, and mass transport around and within biofilm colonies. These methods support comprehensive investigations of the co-development of biofilm and habitat heterogeneity.

Biofilms have complex interactions with their surrounding environment. To comprehensively investigate biofilm-environment interactions, we present here a series of methods to create heterogeneous chemical environment for biofilm development, to quantify local flow velocity, and to analyze mass transport in and around biofilm colonies.

Biofilms are surface-attached microbial communities that have complex structures and produce significant spatial heterogeneities. Biofilm development is ...

Bioengineering

Early Detection of Cyanobacterial Blooms and Associated Cyanotoxins using Fast Detection Strategy

Fast detection of cyanobacteria and cyanotoxins is achieved using a Fast Detection Strategy (FDS). Only 24 h are needed to unravel the presence of cyanobacteria and related cyanotoxins in water samples and in an organic matrix, such as bivalve extracts. FDS combines remote/proximal sensing techniques with analytical/bioinformatics analyses. Sampling spots are chosen through multi-disciplinary, multi-scale, and multi-parametric monitoring in a three-dimensional physical space, including remote sensing. Microscopic observation and taxonomic analysis of the samples are performed in the laboratory setting, which allows for the identification of cyanobacterial species. Samples are then extracted with organic solvents and processed with LC-MS/MS. Data obtained by MS/MS are analyzed using a bioinformatic approach using the online platform Global Natural Products Social (GNPS) to create a network of molecules. These networks are analyzed to detect and identify toxins, comparing data of the fragmentation spectra obtained by mass spectrometry with the GNPS library. This allows for the detection of known toxins and unknown analogues that appear related in the same molecular network.

A fast-multidisciplinarystrategy for early detection of cyanobacterial blooms and associated cyanotoxins is described here. It allows for the detection of cyanobacteria and related cyanotoxins in water samples and in organic matrices, such as bivalve samples, in 24 h.

Fast detection of cyanobacteria and cyanotoxins is achieved using a Fast Detection Strategy (FDS). Only 24 h are needed to unravel the presence of ...

Chemistry

Protocol for Co-Culturing Sucrose-Secreting Cyanobacteria with Fungal Partners

Assembly and Quantification of Co-Cultures Combining Heterotrophic Yeast with Phototrophic Sugar-Secreting Cyanobacteria

With the increasing demand for sustainable biotechnologies, mixed consortia containing a phototrophic microbe and heterotrophic partner species are being explored as a method for solar-driven bioproduction. One approach involves the use of CO2-fixing cyanobacteria that secrete organic carbon to support the metabolism of a co-cultivated heterotroph, which in turn transforms the carbon into higher-value goods or services. In this protocol, a technical description to assist the experimentalist in the establishment of a co-culture combining a sucrose-secreting cyanobacterial strain with a fungal partner(s), as represented by model yeast species, is provided. The protocol describes the key prerequisites for co-culture establishment: Defining the media composition, monitoring the growth characteristics of individual partners, and the analysis of mixed cultures with multiple species combined in the same growth vessel. Basic laboratory techniques for co-culture monitoring, including microscopy, cell counter, and single-cell flow cytometry, are summarized, and examples of nonproprietary software to use for data analysis of raw flow cytometry standard (FCS) files in line with FAIR (Findable, Accessible, Interoperable, Reusable) principles are provided. Finally, commentary on the bottlenecks and pitfalls frequently encountered when attempting to establish a co-culture with sugar-secreting cyanobacteria and a novel heterotrophic partner is included. This protocol provides a resource for researchers attempting to establish a new pair of co-cultured microbes that includes a cyanobacterium and a heterotrophic microbe.

This protocol provides a comprehensive guideline for the setup and quantitative monitoring of co-cultures, including photoautotrophic sugar-secreting cyanobacteria and heterotrophic yeasts.

With the increasing demand for sustainable biotechnologies, mixed consortia containing a phototrophic microbe and heterotrophic partner species are being ...

Procedure to Evaluate the Efficiency of Flocculants for the Removal of Dispersed Particles from Plant Extracts

Plants are important to humans not only because they provide commodities such as food, feed and raw materials, but increasingly because they can be used as manufacturing platforms for added-value products such as biopharmaceuticals. In both cases, liquid plant extracts may need to be clarified to remove particulates. Optimal clarification reduces the costs of filtration and centrifugation by increasing capacity and longevity. This can be achieved by introducing charged polymers known as flocculants, which cross-link dispersed particles to facilitate solid-liquid separation. There are no mechanistic flocculation models for complex mixtures such as plant extracts so empirical models are used instead. Here a design-of-experiments procedure is described that allows the rapid screening of different flocculants, optimizing the clarification of plant extracts and significantly reducing turbidity. The resulting predictive models allow the identification of robust process conditions and sets of polymers with complementary properties, e.g. effective flocculation in extracts with specific conductivities. The results presented for tobacco leaf extracts can easily be adapted to other plant species or tissues and will thus facilitate the development of more cost-effective downstream processes for commodities and plant-derived pharmaceuticals.

The design-of-experiments procedure presented here allows the evaluation of different flocculants in terms of their ability to aggregate dispersed particles in plant extracts, thus reducing turbidity and the costs of downstream processing.

Plants are important to humans not only because they provide commodities such as food, feed and raw materials, but increasingly because they can be used ...

Determination of the Settling Rate of Clay/Cyanobacterial Floccules

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