Determination of the Settling Rate of Clay/Cyanobacterial Floccules

Summary

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.

Abstract

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.

Introduction

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 sediment^1,2,3. 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 blooms^{2,3,4,5,6,7,8,9,10,11,12}. 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.

Protocol

1. Preparing Cyanobacterial Cultures

1. Preparing inoculation cultures using solid media
   1. 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.
   2. Maintain Synechococcus cells on plates containing solid media (A+ liquid media¹⁴ and 1.5% agar¹⁵). These will be referred to hereafter as inoculation cultures.
   3. Incubate the plates at 32 ± 2 °C with continuous light provided at 30–50 µmol photons m^-2 s^-1,15,18.
   4. Repeat steps 1.1–1.3 and replace A+ media with BG-11¹⁶ in step 1.1.2 for freshwater cyanobacterial species, such as Synechocystis sp. PCC 6803^16,17, referred to hereafter as Synechocystis. Refer to Tables 1–4 for the composition of A+ and BG-11 media.
2. Preparing liquid cultures: Each tank experiment requires one 400 mL cyanobacterial culture.
   1. Collect one 250 mL and two 1 L heat-resistant Erlenmeyer flask.
   2. 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
   3. 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).
   4. Seal each flask with a gas permeable foam stopper and cover the foam stopper and flask neck with tinfoil.
   5. Label and autoclave the flasks at 121 °C and 100 kPa for 25 min. Allow the flasks to cool to room temperature.
   6. Prepare a laminar flow hood by sterilizing the surface with 70% alcohol.
   7. 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.
   8. Sterilize the wire inoculation loop briefly using the flame of a Bunsen burner and allow it to cool down.
   9. Drag the cool loop along the surface of the inoculation culture to collect cyanobacterial cells.
   10. 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.
   11. 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.
   12. Remove the inoculation loop and re-sterilize the lip of the flask using the Bunsen burner flame.
   13. Replace the foam stopper and foil cap onto the Erlenmeyer flask (without touching the foam stopper).
   14. Sterilize the inoculation loop briefly within the Bunsen burner flame.
   15. Prepare a growth room (referred to as the growth chamber) where the temperature is maintained at 30 °C^15,18 and light intensity is 30–50 µmol photons m^-2 s^-1,15,18. Humidity is not actively controlled.
   16. Allow the inoculated 50 mL culture to incubate by shaking at 150 rpm in the growth chamber, with a maintained temperature of 30 °C. Keep the mouth of the flask covered during the agitation to control evaporative losses and prevent contamination.
   17. 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 0.4–0.6.
   18. 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.
   19. Repeat step 1.2.10 for both flasks within a laminar flow hood.
   20. 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.
   21. 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.
   22. Agitate the 1 L flask at 150 rpm at 30 °C with the bubbler apparatus attached to an air pump, which circulates a humidified air mixture through the solution^15,18 at 300 mL/min.
   23. Allow the culture(s) to grow within the growth chamber at 30 °C for 120–168 h. A successful culture should appear bright green and have an OD_{750 nm} of 0.4–0.6.
   24. Repeat step 1.2 to produce a control culture, which will not be mixed with the clay.
   25. Experiments using dead biomass require an additional step. Autoclave the 1 L flask culture for 60 min at 121 °C and allow it to cool to room temperature.

2. Experimental Set Up

1. 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.
2. 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
3. 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.
4. 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.
5. Place a black cloth over the light bank, tank, and camera to shield the experiment from outside light sources (Figure 1).

3. Flocculation Experimental Protocol

1. 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).
2. Dilute the 400 mL culture (initial cell concentration of 4–6 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.
3. 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.
4. Begin video recording. In the case of multiple experiments, a sign with the experiment identifier can be temporarily held before the camera.
5. 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 OD_{750 nm} values. Take all samples in triplicate to ensure accurate results.
6. Quickly pour the clay (50 g) into the tank with vigorous agitation using a stirring stick for 10–15 s.
7. Start the timer and take the initial 1 mL sample for Chl a determination using a P1000 pipette (sample Time₀).
8. 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).
9. Once the experiment is finished, save the video file, turn off the video camera, and process all samples for Chl a determination (step 4).
10. 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.
11. 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.
12. If dead biomass is used during the experiment, use the same experimental procedure. However, the disposal procedure of the solution does not require autoclaving.

4. Sample Processing and Evaluation

1. 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.
2. Remove excess media using a P1000 pipette and add 1 mL of 100% methanol (20 °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.
3. Incubate the samples at -20 °C for 24 h to facilitate the extraction of Chl a.
4. 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.
5. Determine Chl a concentration spectrophotometrically by measuring absorbance at 665 nm and 652 nm, and OD at 750 nm. The Chl a concentration is determined using the formula Chl a = 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.
6. Plot calculated Chl a values versus time.

Results

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 5–6, Figure 2–3). Cyanobacte...

Discussion

Flocculation catalyzed by cyanobacterial cell-clay interaction has attracted a lot of interest in the fields of ecology and engineering^{2,3,4,5,6,7,8,9,10,11,1...}

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
cyanobacteria (in this study: Synechococcus sp. PCC 7002 and Synechocystis sp. PCC 6803)	Pasteur Culture Collection	PCC 7002 or PCC 6803	used to inoculate the plates
agar	Thermo Scientific	CM0003	used to fill two petri dishes
Petri plates (standard bacteriology, 100 x 15 mm)	Sarstedt	82.1473.001	2 required
1 L heat resistant Erlenmeyer flask	Pyrex	4980-125	1 required
250 mL heat resistant Erlenmeyer flask	Pyrex	4980-250	1 required
Nichrome inoculating loop with handle	Fisher Scientific	14-956-103	1 required
tinfoil	Reynolds Wrap Aluminum Foil	89079-067	50 cm required; used to cover foam stopper and neck of erlenmeyer flasks
growth media (e.g. A+)			1050 mL required; produced using composition described in tables 1-4
Bunsen Burner	Fisher Scientific	S95941	1 required
plastic tubing	Fisher Scientific	S504591	1 m required; used to create the bubbling apparatus
sponge stopper	Jaece Industries Inc	14-127-40E	1 required; hole made in center for pipette; used for constructin the bubbling apparatus
acrylic sheet	Home Depot	Optix clear acrylic sheet model # MC-102S	1 required; used to construct acrylic tank (20 x 30 x 5.1 cm)
clear waterproof silicone adhesive	Home Depot	Loctite clear silicone model # 908570	1 required; used to construct acrylic tank (20 x 30 x 5.1 cm)
camera or video recorder	Panasonic	HC-V770 HD camcorder	1 required
tripod	Magnus	VT-300	1 required
black cloth	primomart	EAN 0726670162199; Part number 680254blacknappedfr	1 required; duvetyne light block-out cloth; approximatly 152 x 213 cm to cover tank experiment
heat resistant serological pipet	corning incorporated C708510	13-671-101G	1 required; used to create the bubbling apparatus
sample vials	Dynalon	S30467	at least 12 (will vary with time interval chosen)
heat resistant glass pipette	Fisher Scientific	Corning Incorporated C708510, 13-671-101G	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.
microcentrifuge	Eppendorf	22 62 120-3	1 required;Comparable products may be used if capable of centrifuging 1.5 -2 mL microfuge tubes at 13,000 x g
vortex machine (Vortex-Genie 2)	Scientific Industries, Inc	SI-0236	1 required
100% methanol	Fisher Scientific	A412-500 SDS	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.
cuvettes (1.6  mL, polystyrene)	Sarstedt	67.742	at least 12 required
spectrophotometer	Fisher Scientific	222-271600	1 required; Pharmacia Biotech Novaspec ll could also be used.
light bulbs	Home Depot	model # 451807; internet #205477895; store SKU #1001061538	6-8 bulbs required to provide light for the tank experiments
pipette (Pipetman Classic P1000	Gilson	F123602	used to collect samples
37 % Hydrochloric acid	Sigma-Aldrich	258148	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.
Foam stopper (small)	Canlab	T 1385
Foam stopper (large)	Canlab	T 1387	Requires some intact stoppers and some with a single hole through the centre
30 °C incubator/growth room with continuous illumination			1 required
70 % Ethanol	Fisher Scientific	BP8201500	30 mL  required;Caution: Toxic and flammable. Wear lab coat and safety glasses
hydrophobic air filter (Midisart 2000, 0.2 µm)	Sartorius	17805	1 required
clay (e.g. kaolin)	Fisher Scientific	MFCD00062311	at least 50 g required
microfuge tubes (2 mL, polypropylene)	Sarstedt	72.695.500	Comparable products may be used. At least 12 (will vary with time interval chosen)
1000 µL pipet tips	Sarstedt	70.762	1 required