This study was supported by the Natural Sciences and Engineering Research Council (NSERC), Canada Foundation for Innovation (CFI), Canada First Research Excellence Fund (CFREF), Alberta Innovates, General Sir John Monash Foundation, the Government of Alberta, and the University of Calgary. Thanks is extended to Mark Toonen for the electrical work and William Richardson for solubility calculations.

1. Construction of the PBR stand
<ol>
	<li>With a handheld hacksaw, cut five 380 mm lengths and two 200 mm lengths of angled slotted steel. Fasten together with bolts and large corner braces to make a stand&#39;s base (Figure 1A). Glue on safety end caps.</li>
	<li>Connect two uncut (1220 mm) vertical lengths of angled slotted steel to the base. Secure with bolts and metal corner gussets (Figure 1B). Glue on safety end caps.</li>
	<li>Cut four 65 mm flat lengths of slotted steel. Bolt these at 90&#176; angles onto the vertical supports&#8212;attach two to each support, one 130 mm up from the base (Figure 1C) and one 60 mm down from the top.</li>
	<li>Fasten vertical supports across their top with a horizontal 140 mm length of flat slotted steel (crossbeam) bolted to the back of the frame (Figure 2A).</li>
</ol>
2. Construction of the light chamber
<ol>
	<li>Cut a white 153 mm diameter polyvinyl chloride (PVC) pipe to a length of 1070 mm. Cut the pipe in half lengthwise with a bandsaw. Sand all edges.</li>
	<li>Evenly space and center four aluminum heat sink channels vertically along with the interior of the pipe. Do not attach channels within 20 mm from the pipe&#39;s cut edge. With small bolts, secure the channels in place at their top and bottom (Figure 2B).</li>
	<li>Bolt one-half of the pipe to the stand using the horizontal supports fashioned in step 1.3.</li>
	<li>Lay the reactor down and reunite the pipe halves by taping them together. Center the piano hinge along one cutline. Trace the hinge holes and drill the pipe accordingly. Use a rivet gun and medium-length rivets to fasten the hinge to the pipe.</li>
	<li>Use a small bungee cord (see Table of Materials) to hold the pipe shut (Figure 1C).</li>
	<li>Consult an electrician to wire in the LED lights and install the following four components: LED driver, digital multiplex (DMX) decoder, DMX lighting controller, and switch box (see Table of Materials). Fix all the components to the rear of the PBR according to Figure 2A.</li>
</ol>
3. Construction of the bottle platforms
<ol>
	<li>Cut platform shapes (Figure 3A) out of hard plastic, e.g., high density polyethylene (HDPE) (see Table of Materials), using computer numerical control (CNC) milling. Make three of each shape. 
	NOTE: Cutting spare top and bottom layers is recommended.</li>
	<li>Tape the bottom and top layer together. Mark and drill five small holes, 6 mm in diameter, through both shapes (Figure 3A). Using a larger drill bit, carefully expand the surface of these holes so that bolt heads can be recessed (Figure 3B).</li>
	<li>For each of the three platforms, bolt two small corner braces to the back half of the pipe. 
	NOTE: The distance between the top of the braces should be 350 mm.</li>
	<li>Center each bottom layer on top of its braces. Mark the location of the drill holes from beneath the braces. Drill two 6 mm diameter holes. Using a larger drill bit, carefully expand the surface of the holes so that bolts can be recessed.</li>
	<li>Bolt bottom layers to their braces (Figure 3B&#8211;C).</li>
	<li>Cut fifteen pieces of 12 mm long 6.35 mm outside diameter (OD) rigid tubing. Sandwich five pieces of the rigid tube between each top and bottom layer. Fasten layers and tubing together with long narrow bolts according to Figure 3B&#8211;D.</li>
	<li>Drill a large hole in the PVC pipe behind each platform. Insert each micro magnetic stirrer into its platform. Thread each stirrer&#39;s electrical cable through these newly cut holes (Figure 3C&#8211;E). Connect each stirrer to its respective control unit as well as a power outlet.</li>
	<li>Place a 1 L bottle on each platform. Add eye bolts in the rear pipe at bottle neck level. Wrap a small bungee cord around each bottle neck to add stability (Figure 4A). 
	&#8203;NOTE: Throughout the protocol, color coding platforms, bottles, magnetic stirrers, and all associated cables and sensors will be helpful.</li>
</ol>
4. Construction of the liquid sampling ports (optional)
<ol>
	<li>Cut three 60 mm lengths of 6.35 mm OD rigid tubing. Using a 5 mm drill bit, drill holes through each rubber stopper. Push rigid tube lengths through the stopper.</li>
	<li>Cut three 60 mm lengths of 3.18 mm OD rigid tubing. Connect these with a straight reducer to the protruding tube on the underside of each stopper.</li>
	<li>Insert a one-way stopcock valve (e.g., port 1 = female Luer, port 2 = male slip Luer) in the protruding tube on each stopper surface (Figure 4A).</li>
	<li>Cut three 30 mm lengths of 3.18 mm OD flexible tubing. Insert Luer fittings (e.g., male Luer to hose barb and female Luer to hose barb) on either end.</li>
	<li>Connect the pieces made in step 4.4 to the stopcock valves on the surface of each rubber stopper (Figure 4A). 
	&#8203;NOTE: Many combinations of stopcocks and Luer fittings can produce the same result. The design should allow liquid to be drawn out or inserted via a syringe.</li>
</ol>
5. Hooking up volumetric gas sensors
<ol>
	<li>Prepare gas sensors according to the manufacturer&#39;s instructions. 
	NOTE: This mainly involves filling gas sensors with packing liquid (Figure 4B).</li>
	<li>To make the gas lines, cut three 1000 mm lengths of 3.18 mm OD flexible tubing.</li>
	<li>Drill three 4 mm diameter holes in the rear PVC pipe. Position holes next to the hinge at bottle neck height. Thread gas lines through these holes (Figure 4A).</li>
	<li>To the end of the gas line inside the PVC pipe, add a Luer fitting (e.g., hose barb to male Luer) and connect a one-way stopcock valve (e.g., port 1 = female Luer, port 2 = male Luer). 
	NOTE: The valve is only required when liquid sampling ports are also installed.</li>
	<li>Join the other end of the gas line to the inlet port of the gas sensor using a straight reducer. Secure this connection with a zip tie.</li>
	<li>Connect all gas sensors to the digital input module (DIM) with jack plug cables and the DIM to a nearby computer.</li>
	<li>Install the data acquisition software (see Table of Materials) on a Windows operating system and plug in the licensing key dongle. Add sensor calibration files to the software&#39;s calibration directory.</li>
</ol>
6. Programming of the light regime
<ol>
	<li>Using the rear on/off switch, turn on the PBR and connect the DMX lighting controller to a computer via a micro-USB cable.</li>
	<li>Download Store Upgrade Tools (SUT) and the LED control software (see Table of Materials). Register the DMX lighting controller online.</li>
	<li>Open the LED control software and select Click here to work with the USB-DMX interface: SUSHI-RB-RJ.</li>
	<li>Under the Set-up tab in the ScanLibrary box, select the Generic folder and Single Channel. Change ScanLibrary settings to DMX universe 1, the number of fixtures to 4, and the index number to 1. In the upper-right corner, change the drop-down box to List View. Lastly, click on Patch (Supplementary Figure 1). 
	NOTE: In the DMX universe, one box tests each LED strip&#39;s control by sliding the dimmer buttons or entering a numerical value into the text box.</li>
	<li>Make a standard curve that relates the digital light setting in the lighting control software to the light intensity experienced in the center of the PVC tube (Figure 5). Measure internal light intensity with a small spherical probe (see Table of Materials) suspended in the center of the PVC pipe.</li>
	<li>Advance to the Editor tab. To build a custom light program, create a new Scene and begin adding Steps. Refer to Supplementary Table 1 for an example 16:8 h diurnal program. Set the scene to loop. 
	NOTE: Steps break down scenes into blocks of time, each of which can be set to different light intensity. Steps range from 1 s to 43 min. Here, 30 min steps are most convenient. Multiple scenes can be loaded onto one DMX lighting controller device.</li>
	<li>Create an additional helper scene that is immediately recognizable, e.g., two of the four LEDs on. 
	NOTE: Scenes can be cycled through manually using the button on the side of the DMX lighting controller. If the desired light program begins during the night, it will be impossible to distinguish whether the light program has already started. The helper scene serves as an indicator that the DMX lighting controller is functioning correctly.</li>
	<li>Save the scenes and advance to the Stand Alone tab. Write the memory of the DMX lighting controller and disconnect the device from the computer.</li>
	<li>Connect the DMX lighting controller to its power source using a micro-USB.</li>
	<li>Before starting an experiment, test the light program by logging the internal light intensity for a 24 h duration. If the liquid temperature is of interest, log this simultaneously with a submerged temperature probe (Figure 6).</li>
</ol>
7. Starting an experiment
<ol>
	<li>Sterilize media, bottles, stir bars, rubber stoppers, sampling ports, threaded aperture screw caps, and tubing. 
	NOTE: All components used in this design are autoclavable except the valves and Luer fittings&#8212;there are autoclavable alternatives from other manufacturers.</li>
	<li>Open the data acquisition software and fill in the configuration page (Supplementary Figure 2). Assign calibration files to their respective sensors.</li>
	<li>Under Directory File Name select the corresponding DIM port number folder. Click on Current Folder&#160;and repeat for all ports.</li>
	<li>Click on OK to move on to the logging page.</li>
	<li>Fill bottles to the desired volume with cultivation medium (Supplementary Tables 2,3). 
	NOTE: Each of these bottles will hold a maximum of ~1.1 L with a small headspace (~80 mL, including the gas line).</li>
	<li>Centrifuge the stock culture in three balanced 50 mL tubes for 15 min at 4500 x g to yield three pellets. Add one pellet to each bottle&#8211;wash in pellets with a serological pipette and fresh medium. 
	NOTE: The culture density of day 0 is also known as the initial biomass concentration (IBC). To measure the IBC in gAFDW.L-1, an additional 50 mL tube may be centrifuged in step 7.6. The resulting pellet can then be dried and combusted15,19. Step 7.6 will likely require modification based on the individual aims of users and their experiments.</li>
	<li>Drop a magnetic stirrer, 25 x 8 mm, into each bottle.</li>
	<li>Seal each bottle opening with a rubber stopper and threaded aperture screw cap (Figure 4A). If optional sampling ports are installed, close the valves.</li>
	<li>Locate the end of each gas line inside the PVC pipe (built in step 5.4) and attach a needle to the valve&#39;s male Luer port.</li>
	<li>Connect each bottle to its gas sensor by piercing each rubber stopper with the corresponding needle.</li>
	<li>Launch each gas sensor individually by checking the tick box&#160;on the left side of the screen, clicking on Start, and inputting a file name. Click OK and repeat for all sensors&#160;(Supplementary Figure 3). 
	NOTE: While logging, do not exit the data acquisition window. Set power and sleep settings of the computer to never and postpone computer updates for the duration of the experiment.</li>
	<li>Switch on the PBR and ensure that the DMX lighting controller is plugged into a power supply. The first programmed scene will automatically commence. Refer to step&#160;6.7&#160;to confirm that the DMX lighting controller is functioning correctly.</li>
</ol>
8. Bottle sampling (optional)
<ol>
	<li>Prepare an additional 500 mL of the fresh medium before commencing the experiment (Supplementary Table 2). 
	NOTE: If a 24 h light program with a 16:8 h diurnal cycle was started at 9 am, then sampling times before dusk and dawn would fall at 8 am and 4 pm (Supplementary Table 1). Here, dawn and dusk refer to 30 min steps that transition lights from ON to OFF and vice versa.</li>
	<li>Close the valve on the gas line.</li>
	<li>Connect a syringe (10 mL) to the sampling port valve (Figure 4A).</li>
	<li>Open the sampling port valve and withdraw 8 mL of culture. 
	NOTE: Between 5&#8211;10 mL is recommended. Removing liquid generates a vacuum in the headspace, making volumes &#62;10 mL difficult to extract.</li>
	<li>Close the sampling port valve and disconnect the syringe.</li>
	<li>Connect a syringe containing 8 mL of fresh medium (from step 8.1) to the sampling port valve.</li>
	<li>Open the sampling port valve and inject fresh medium. 
	NOTE: Replacing the volume of sampled culture with fresh medium serves to maintain an equal headspace volume and pressure and flush the sampling port line.</li>
	<li>Close the sampling port valve before disconnecting the syringe.</li>
	<li>Repeat steps 8.2&#8211;8.8 at each sampling time.</li>
</ol>
9. Ending an experiment
<ol>
	<li>Check all active port tick boxes in the data acquisition window and click on Stop.</li>
	<li>To export data, select File and Offline Data. Select all relevant log files. Export the data to spreadsheet software and save.</li>
	<li>For each bottle, convert the total volume of oxygen measured in mL to moles using the ideal gas law. Predict the weight of biomass grown (gAFDW) if 1.05 mole of O2 is generated for each mole of biomass produced. Take the molar weight of biomass as 24.6 g mol-1.</li>
	<li>Manually curate the flow rate data. Use units of mL/h and a 3-point moving average.</li>
</ol>

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H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Growth+of+the+microalgae+Neochloris+oleoabundans+at+high+partial+oxygen+pressures+and+sub-saturating+light+intensity.">Growth of the microalgae Neochloris oleoabundans at high partial oxygen pressures and sub-saturating light intensity.</a> Bioresource Technology. 104, 565-570 (2012).</li><li>Latifi, A., Ruiz, M., Zhang, C. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Oxidative+stress+in+cyanobacteria.">Oxidative stress in cyanobacteria.</a> FEMS microbiology reviews. 33 (2), 258-278 (2009).</li><li>Morillas-Espa&#241;a, A., Lafarga, T., G&#243;mez-Serrano, C., Aci&#233;n-Fern&#225;ndez, F. G., Gonz&#225;lez-L&#243;pez, C. V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Year-long+production+of+Scenedesmus+almeriensis+in+pilot-scale+raceway+and+thin-layer+cascade+photobioreactors.">Year-long production of Scenedesmus almeriensis in pilot-scale raceway and thin-layer cascade photobioreactors.</a> Algal Research. 51, (2020).</li><li>Melis, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Solar+energy+conversion+efficiencies+in+photosynthesis:+minimizing+the+chlorophyll+antennae+to+maximize+efficiency.">Solar energy conversion efficiencies in photosynthesis: minimizing the chlorophyll antennae to maximize efficiency.</a> Plant Science. 177 (4), 272-280 (2009).</li></ol>

The authors have nothing to disclose.

Within this protocol, focus on the following steps increases the likelihood of generating reproducible, high-quality data. When constructing the reactor stand (step 1), the base must be sturdy with well-alligned vertical supports. Slotted steel has sharp edges, so the addition of safety caps is essential. Bottle platform surfaces need to be completely flat, the magnetic stirrer and bolt heads should both sit below the top layer surface (steps 3.2&#8211;3.6). According to the manufacturer&#39;s instruction, the gas sensor packing liquid should be filled to the &#34;tracing screw for liquid level&#34; for accurate oxygen measurements. This liquid level should be checked regularly as evaporation of the packing liquid can short-circuit the measurement cell. All three gas lines made in step 5.2 should be the same length; this enures that replicates have identical headspace volumes. Before commencing an experiment, it is advisable to test the programmed light regime by logging light intensity over a 24 h period (step 6.11). If increases in liquid temperature are of concern, this test should also include a sealed bottle with an internal temperature probe (step 6.11). When logging, do not exit the data acquisition software window; this will terminate logging. If taking culture samples, be careful not to release headspace gas by opening valves in the wrong sequence (steps 8.2-8.8). When reviewing experimental data be advised that the data acquisition software automatically generates a moving average of the flow rate. This inflates the value of the one or two flow rate readings generated overnight. Curate gas sensor logs manually to rectify this.
The most common setback with this method is the potential to short circuit the gas sensor if the liquid packing level declines. There are two ways this can occur. Firstly, evaporation can slowly reduce the liquid level. However, this is unlikely over a short-term (&#60;7 day) experiment29. Secondly, high respiration rates can pull oxygen into the solution and generate a headspace under-pressure. When light energy is unavailable, microalgae use aerobic respiration to supply the energy required for cellular maintenance and repair28. Hence, in dense cultures during non-illuminated hours, oxygen consumption, and the resulting under-pressure, can be substantial. This sucks packing liquid from the gas sensors into the gas line. The distance the packing liquid travels is proportional to the amount of nighttime respiration. If the packing liquid enters the bottles, this generates an oil slick on the liquid surface.
If high nighttime respiration rates are expected, modifications to the protocol can be made. The simplest way to avoid under-pressure is to leave the bottle headspaces open overnight. This also has the advantage of easing DO levels by decreasing the partial headspace pressure of O2. High DO concentrations are believed to be detrimental to growth as O2 can impede the activity of Rubisco and may trigger oxidative stress30,31. It is not uncommon for culture suspensions to reach 4x oversaturation even when in contact with the atmosphere25,32. To open the headspace, disconnect the gas line from the needle spanning the rubber stopper. Nighttime hours can serve as a window to top up gas sensor packing liquid or manipulate continuous experiments with little impact on data collection. For example, one may alter the culture density, refresh nutrients, add an amendment, or introduce a pathogen. Bottles must be re-sealed, and the gas sensor line reconnected before lights turn back on. The oxygen measurements collected from experiments with closed versus open nighttime headspaces will differ.
When bottles remain sealed, nighttime oxygen consumption reduces the number of moles of O2 in the headspace. This causes packing liquid to creep up the gas sensor line to maintain headspace pressure. When lights turn on, oxygen production resumes. Packing liquid must be pushed back into the gas sensor before flow rate readings commence. This lag is therefore proportional to the degree of nighttime respiration. In this way, when the headspace remains closed, O2 readings represent net O2 production (photosynthetic production - respiratory consumption). Conversely, when the headspace is open at night, atmospheric gas replaces what headspace O2 is consumed, and no packing liquid enters the gas line. The result is that respiratory O2 consumption is not accounted for in O2 production data. This may reduce the accuracy of AFDW biomass growth estimates. However, it should not impact the utility of using daytime O2 production as a metric to compare growth between treatments.
All laboratory PBRs are afflicted by the same limitation; artificial lights cannot replicate the solar spectrum. Microalgae use wavelengths of light between 400&#8211;700 nm for photosynthesis. This region is referred to as photosynthetically active radiation (PAR)33. Sunlight and artificial light vary in their relative contribution of wavelengths within this range. This, alongside favorable temperatures and a constant light supply, means that laboratory growth data often cannot be reliably extrapolated to outdoor conditions. These PBRs can, however, address one of the limitations of laboratory PBR light supply. Sunlight intensity is highly variable throughout the day, with cloud cover generating transient fluctuations in incident PAR. The lighting control software and DMX lighting controller can provide light intensities from 0 to 2400 &#181;molphotons m-2 s-1 and beyond. Light regimes can be broken down into individual increments as short as 1 s. Tunable light intensity allows the user to mimic outdoor light patterns more closely than standard PBR setups. Here, simulated 30 min dawn and dusk intervals fade day and night cycles together (Supplementary Table 1).
Although AFDW density has become the standard measure of growth, this method can require substantial culture volumes, a 2&#8211;3-day processing period, and generates one data point at a time. Further, if conditions become unfavorable and cells die, AFDW density does not discriminate between actively photosynthesizing cells and those that are decomposing. Quantifying the rate of photosynthetic oxygen production serves as an alternative growth proxy. This PBR design can record oxygen production continuously with little user intervention while conserving culture volume. Data resolution could be improved by selecting a gas sensor with a lower measurement cell volume, e.g., 1 mL. Further, if cultures are well-mixed, users may decide to install a spectrophotometer for continuous optical density readings. If temperature control of the medium is desired, a recirculating chiller could be added. These PBRs are a valuable addition to a laboratory looking to expand its microalgal research capacity without heavy financial investment. They are particularly suitable for those working with high alkalinity, high pH species like Spirulina. These PBRs offer light regime flexibility and are valid for rapid, replicated, laboratory growth comparisons.

Microalgae and cyanobacteria, collectively called microalgae for simplicity, are championed for their potential in sustainable biotechnology. They are attractive candidates due to their rapid growth, ability to be cultivated on non-arable land, and for their use of sunlight to drive the conversion of carbon dioxide to biomass1,2,3. Microalgal biomass can be converted into products such as bioenergy in the form of oil or gas, food dyes and nutritional supplements, and materials such as biopolymers1,4,5,6,7. In addition, they can be used to treat wastewater or remediate waterbodies by consuming excess nutrients8,9. Given this, microalgal research is widespread and established. The field grows as society reconsiders the carbon intensity and environmental sustainability of current manufacturing and energy generation approaches.
Three fundamental requirements of laboratory-based microalgal studies are a culture vessel, light source, and method to quantify growth. The term photobioreactor (PBR) describes a set-up in which culture vessels are illuminated10. Commonly, studies of microalgae aim to compare growth between two or more treatments, e.g., different growth media, light regimes, or species11,12,13. For statistical relevance, each condition, e.g., treatment and control, should be replicated. If control and treatment are run simultaneously, this means many PBRs must be monitored and sampled for the duration of an experiment. The challenge with operating multiple PBRs is two-fold. Firstly, supplying a uniform light intensity to each PBR is essential for reproducibility but can be difficult. The amount of light incident on the vessel surface is influenced by its distance from the light source, shading from adjacent vessels, and background light fluctuations14. Secondly, a method to accurately quantify growth must be selected.
Growth is commonly measured by cell count, optical density (OD), chlorophyll A content, dry weight (DW) density, and ash-free dry weight (AFDW) density15. Cell counts, chlorophyll A content, and gravimetric methods are manual processes that produce discrete data points. OD can be measured continuously and non-invasively with a spectrophotometer, provided it is well-calibrated against another method such as AFDW density15. However, OD measurements and Chlorophyll A content can be unreliable as results vary under different culture conditions, e.g., between species and throughout the growth cycle15,16. For Chlorophyll A, the extraction method can also impact pigment yield17. Chlorophyll A content is particularly useful in tracking the growth of microalgae within microbial communities which also contain non-photosynthetic organisms17,18. When choosing a method to determine growth, it is essential to consider the morphology of the suspension. When organisms clump and are not well mixed, OD and cell counts aren&#39;t possible15. A single method is not suited to all experimental applications&#8212;researchers must decide which methods are practical and relevant to their experimental aims.
AFDW is a reliable method that enables growth comparisons among various culture conditions, notably, between species and culture media15,19,20. To calculate AFDW, a sample of microalgal culture is first concentrated, either by filtration or centrifugation, and dried. At this stage, the DW can be determined. Usually, the DW sample contains at least 8&#8211;10% ash-inorganic material such as salts and particulate matter15. DW tracks growth trends but can be skewed if the contribution of inorganics varies. To determine AFDW density, dry biomass is combusted at high temperature; this vaporizes the organic or useful portion while leaving ash (inorganics) behind19. To calculate the AFDW, the weight of the ash fraction is subtracted from that of the DW fraction. Typically, in microalgal suspensions, AFDW ranges from 0.1&#8211;3 g/L12,21,22. Small volumes of dilute suspensions yield little dry biomass, &#60;10 mg. After combustion, ash may only weigh 1 mg. Therefore, depending on the culture density, this method requires volumes between 5&#8211;100 mL and analytical scales accurate to 0.1 mg12,15,19,22. Laboratory PBRs are typically small, a couple of liters at most, hence every liquid sample depletes culture volume. Further, the AFDW method is manual and takes 2&#8211;3 days. For replicated and repetitive experiments, an automated and continuous process is preferable.
For microalgae that use bicarbonate as the carbon source, two additional growth metrics can be measured continuously. Photosynthesis consumes bicarbonate and produces oxygen. Bicarbonate consumption drives up medium pH23. An immersed pH probe can measure this change. Photosynthetic oxygen production increases the medium&#39;s dissolved oxygen (DO) concentration until the medium is saturated. Beyond saturation, oxygen exists as bubbles. Oxygen production is measured by many different techniques: probes gauge DO concentration, manometric devices assess headspace pressure, gas chromatography measures headspace composition, and volumetric sensors record gas outflow24,25,26,27. When oxygen is used as a growth proxy, culture vessels must be fully sealed or allow only gas outflow. For pH and oxygen measurements, carbon must be supplied in the form of bicarbonate, not by CO2 sparging. CO2 sparging decreases the medium pH23 and, as a gas, can disturb oxygen measurements. One advantage of pH and oxygen over optical density is that the method is not compromised if microalgae form clumps. Although indirect, both pH and oxygen are effective in comparing growth between treatments.
PBRs in use today range in complexity. Laboratories may use simple benchtop flasks, custom prototypes, or commercially available products. For research groups seeking to upgrade from flasks, the cost of commercial PBRs or technical skill and part fabrication required to build many prototypes may be a barrier. This manuscript aims to describe the step-by-step design, construction, and operation of laboratory PBRs that bridge this gap. These PBRs have a customizable light regime and monitor growth continuously by recording volumetric oxygen production. This design houses three culture vessels for triplicate replication and can be built with moderate skill and easily accessible materials. This PBR is a valuable addition to a laboratory looking to expand its capacity for microalgal research without investing in very technical or expensive products. When choosing to acquire or build a PBR, researchers must consider the suitability of a design to their culture conditions, financial position, and research questions.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Aluminum channels 
			Imperial: 0.90&#8221; x 39.37&#8221; 
			Metric: 2.3 cm x 100 cm 
			Quantity: 4</td>
			<td>LED World</td>
			<td>AC-AR1-1M</td>
			<td>Required as a heat sink</td>
		</tr>
		<tr>
			<td>Bungee cords, small 
			Quantity: 5</td>
			<td>-</td>
			<td>-</td>
			<td>To secure bottles</td>
		</tr>
		<tr>
			<td>Computer - desktop/laptop 
			Quantity: 1</td>
			<td>-</td>
			<td>-</td>
			<td>-</td>
		</tr>
		<tr>
			<td>Data Logger, HOBO U30 USB Weather Station 
			Quantity: 1</td>
			<td>HOBO, Hoskin</td>
			<td>U30-NRC-VIA-10-S100-000</td>
			<td>Records light sensor information</td>
		</tr>
		<tr>
			<td>Digital interface module, Rigamo, 4-channel 
			Quantity: 1</td>
			<td>Ritter</td>
			<td>N/A</td>
			<td>This is to transmit gas sensor data to the computer</td>
		</tr>
		<tr>
			<td>DMX decoder, 12~24 VDC, DMX-CV-4X5A 
			Quantity: 1</td>
			<td>LITECH, LED World</td>
			<td>LT-840-6A</td>
			<td>Transmit messages which alter the light pattern</td>
		</tr>
		<tr>
			<td>DMX lighting controller, SUSHI-RB-RJ 
			Quantity: 1</td>
			<td>Arcolis, Nicolaudie America Inc.</td>
			<td>SUSHI-RB-RJ DMX</td>
			<td>Encodes the lighting program</td>
		</tr>
		<tr>
			<td>Gas sensor packing liquid (Silox) 
			Quantity: 1 L</td>
			<td>Ritter</td>
			<td>https://www.ritter.de/en/data-sheets/silox</td>
			<td></td>
		</tr>
		<tr>
			<td>Gas sensor, volumetric 
			Quantity: 3</td>
			<td>Ritter</td>
			<td>MGC-1 V3.4 PMMA (https://www.ritter.de/downloads/mgc-milligascounter-en)</td>
			<td>Measures oxygen production</td>
		</tr>
		<tr>
			<td>Glass bottles, round 1 L with GL45 neck 
			Quantity: 3</td>
			<td>Corning, Capitol Scientific</td>
			<td>1395-1L</td>
			<td>Culture vessels</td>
		</tr>
		<tr>
			<td>Hardware - end caps for slotted steel 
			Quantity: 10</td>
			<td>Paulin, Home Depot</td>
			<td>142-612</td>
			<td>To cover sharp edges of slotted steel</td>
		</tr>
		<tr>
			<td>Hardware - eye hooks 
			Quantity: 6</td>
			<td>-</td>
			<td>-</td>
			<td>To secure bottles</td>
		</tr>
		<tr>
			<td>Hardware - metal corner braces (large) 
			Imperial: 4&#34; x 4&#34; 
			Metric: 10 cm x 10 cm 
			Quantity: 8</td>
			<td>-</td>
			<td>-</td>
			<td>Larger brackets to construct metal stand</td>
		</tr>
		<tr>
			<td>Hardware - metal corner braces (small) 
			Imperial: 2 1/2&#34; x 2 1/2&#34; 
			Metric: 6.4 cm x 6.4 cm 
			Quantity: 6</td>
			<td>-</td>
			<td>-</td>
			<td>Small brackets to connect bottle platforms to PVC pipe</td>
		</tr>
		<tr>
			<td>Hardware - metal corner gussets 
			Imperial: 3&#34; x 3&#34; 
			Metric: 7.6 cm x 7.6 cm 
			Quantity: 6</td>
			<td>Paulin, Home Depot</td>
			<td>142-616</td>
			<td>Flat brackets to construct metal stand</td>
		</tr>
		<tr>
			<td>Hardware - piano hinge 
			Imperial: 36&#34; 
			Metric: 91 cm 
			Quantity: 1</td>
			<td>-</td>
			<td>-</td>
			<td>Connects two halves of PVC pipe</td>
		</tr>
		<tr>
			<td>Hardware - rivets 
			Quantity: 40</td>
			<td>-</td>
			<td>-</td>
			<td>To attach piano hinge to PVC tubing</td>
		</tr>
		<tr>
			<td>Hardware - set of bolts, nuts, washers 
			Quantity: 60</td>
			<td>-</td>
			<td>-</td>
			<td>Long thin bolts are required to secure bottle platforms around magentic stirrers</td>
		</tr>
		<tr>
			<td>Hardware - set of bolts, nuts, washers 
			Quantity: 30</td>
			<td>-</td>
			<td>-</td>
			<td>Larger shorter bolts are required to build the metal stand</td>
		</tr>
		<tr>
			<td>LED driver, constant voltage, 96W 24VDC UL Listed IP65 Driver Class 2 regulated power supply 
			Quantity: 1</td>
			<td>Magnitude Lighting, LED World</td>
			<td>CVN96L24DC</td>
			<td>Regulates power to the lights</td>
		</tr>
		<tr>
			<td>LED lights, Cinco Bright LED Flex Strip 
			Quantity: 4 m roll</td>
			<td>EvenBright, LED World</td>
			<td>FA128M57-4M-24V-X</td>
			<td>Roll is trimmed into 4 x 1 m lengths and secured inside the PVC tube</td>
		</tr>
		<tr>
			<td>Light meter, handheld with submersible sperical probe 
			Quantity: 1</td>
			<td>LI-COR</td>
			<td>LA-250A</td>
			<td>Calibrate the reactors light intensity</td>
		</tr>
		<tr>
			<td>Light sensors Photosynthetic Light (PAR) Smart Sensor 
			Quantity: 2</td>
			<td>HOBO, Hoskin</td>
			<td>S-LIA-M003</td>
			<td>Only one is required however two would be good practice in case one malfunctioned</td>
		</tr>
		<tr>
			<td>Magnetic stirrers (MIXdrive 1 XS) with external control units and power supply (MIXcontrol eco) 
			Quantity: 3</td>
			<td>2Mag, 2MAG USA</td>
			<td>MF 40300</td>
			<td>Stirrers sit sandwiched in bottle platforms</td>
		</tr>
		<tr>
			<td>Metal plate 
			Imperial: 24&#34; x 8&#34; 
			Metric: 61 cm x 20.3 cm 
			Quantity: 1</td>
			<td>-</td>
			<td>-</td>
			<td>This is a surface on which to secure electronics, it is attached to the back of the reactor</td>
		</tr>
		<tr>
			<td>Pipe, white PVC 
			Imperial: 6&#34; diameter x 42&#34; high 
			Metric: 15.2 cm x 106.7 cm 
			Quantity: 1</td>
			<td>-</td>
			<td>-</td>
			<td>Cut lengthwise in two halves, used to house lights and bottles</td>
		</tr>
		<tr>
			<td>Plastic (HDPE) sheets 
			Imperial: 4&#34; x 4&#34; x 1/4&#34; 
			Metric: 10 cm x 10 cm x 1 cm 
			Quantity: 6</td>
			<td>Inventables</td>
			<td>30291-01</td>
			<td>For bottle platforms which house magentic stirrers</td>
		</tr>
		<tr>
			<td>Rubber stoppers - GL45 size 
			Quantity: 3</td>
			<td>Duran, VWR</td>
			<td>76289-760</td>
			<td>Seals culture vessels</td>
		</tr>
		<tr>
			<td>Screw caps - with aperture and GL45 neck 
			Quantity: 3</td>
			<td>Corning, Capitol Scientific</td>
			<td>1395-45HTSC</td>
			<td>Generates seal of culture vessels</td>
		</tr>
		<tr>
			<td>Slotted angle steel lengths 
			Imperial: 1-1/2&#34; X 48&#34; x 0.074&#34; 
			Metric: 3.8 cm x 122 cm x 0.19 cm 
			Quantity: 6</td>
			<td>Paulin, Home Depot</td>
			<td>142-202</td>
			<td>Makes up the body of the metal stand</td>
		</tr>
		<tr>
			<td>Slotted flat steel lenghts 
			Imperial: 1-3/8&#34; x 48&#34; x 0.074&#34; 
			Metric: 3.5 cm x 122 cm x 0.19 cm 
			Quantity: 3</td>
			<td>Paulin, Home Depot</td>
			<td>142-222</td>
			<td>Makes up the body of the metal stand</td>
		</tr>
		<tr>
			<td>Software - Easy Stand Alone (ESA)</td>
			<td></td>
			<td>https://www.dmxsoft.com/#apps</td>
			<td>AKA LED control software</td>
		</tr>
		<tr>
			<td>Software - Rigamo v3.1</td>
			<td></td>
			<td></td>
			<td>AKA data acquisition software</td>
		</tr>
		<tr>
			<td>Software - Storage Upgrade Tools (SUT)</td>
			<td></td>
			<td>https://store.dmxsoft.com//</td>
			<td></td>
		</tr>
		<tr>
			<td>Stir bar 
			Imperial: 1&#34; x 5/16&#34; 
			Metric: 2.5 cm x 0.8 cm 
			Quantity: 3</td>
			<td>Fisherbrand</td>
			<td>14-513-59</td>
			<td>Stirs culture</td>
		</tr>
		<tr>
			<td>Switch box 
			Quantity: 1</td>
			<td>-</td>
			<td>-</td>
			<td>Turns power on/off to reactor</td>
		</tr>
		<tr>
			<td>Syringe, 10 mL 
			Quantity: Multiple</td>
			<td>-</td>
			<td>-</td>
			<td>Optional if you wish to extract culture</td>
		</tr>
		<tr>
			<td>Tube adaptor fittings, plastic - Stopcock 1-way 
			Quantity: 6</td>
			<td>Masterflex, Cole Palmer</td>
			<td>RK-12023-33</td>
			<td>Close/open culture vessel line</td>
		</tr>
		<tr>
			<td>Tube adaptor fittings, plastic - variety of male and female luer lock fittings 
			Imperial: to fit 1/16&#34; and 1/8&#34; ID tubing 
			Metric: to fit 1.59 mm and 3.18 mm tubing 
			Quantity: Multiple packets</td>
			<td>Masterflex, Cole Palmer</td>
			<td>RK-30800-16; RK-30800-18; RK-45518-26; RK-45501-00; RK-45501-04</td>
			<td>Many different combinations can achieve the same end result, best to order a variety of fittings</td>
		</tr>
		<tr>
			<td>Tube adaptor fittings, plastic - variety of straight connectors 
			Imperial: to fit 1/16&#34; and 1/8&#34; ID tubing 
			Metric: to fit 1.59 mm and 3.18 mm tubing 
			Quantity: Multiple packets</td>
			<td>Masterflex, Cole Palmer</td>
			<td>RK-40616-04</td>
			<td>Many different combinations can achieve the same end result, best to order a variety of fittings</td>
		</tr>
		<tr>
			<td>Tubing, flexible, transparent 
			Imperial: ID=1/16&#34;, OD=1/8&#34; 
			Metric: ID = 1.59 mm, 0D = 3.18 mm 
			Quantity: 4 m</td>
			<td>Masterflex, Cole Palmer</td>
			<td>RK-06422-02</td>
			<td>Line from culture vessel to gas sensor</td>
		</tr>
		<tr>
			<td>Tubing, flexible, transparent 
			Imperial: ID=1/8&#34;, OD=1/4&#34; 
			Metric: ID = 3.18 mm, 0D = 6.35 mm 
			Quantity: 2 m</td>
			<td>Masterflex, Cole Palmer</td>
			<td>RK-06422-05</td>
			<td>Gas sensor standard tubing size</td>
		</tr>
		<tr>
			<td>Tubing, rigid, transparent 
			Imperial: ID=1/16&#34;, OD=1/8&#34; 
			Metric: ID = 1.59 mm, 0D = 3.18 mm 
			Quantiy: 1 m</td>
			<td>Masterflex, Cole Palmer</td>
			<td>RK-06605-27</td>
			<td>Spans rubber stopper allowing gas to exit</td>
		</tr>
		<tr>
			<td>Tubing, rigid, transparent 
			Imperial: ID=1/8&#34;, OD=1/4&#34; 
			Metric: ID = 3.18 mm, 0D = 6.35 mm 
			Quantity: 1 m</td>
			<td>Masterflex, Cole Palmer</td>
			<td>RK-06605-30</td>
			<td>Spans rubber stopper allowing gas to exit</td>
		</tr>
		<tr>
			<td>Zip ties, small 
			Quantity: 1 packet</td>
			<td></td>
			<td></td>
			<td>Secure tube fittings</td>
		</tr>
	</tbody>
</table>

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.

Watch this Scientific Journal Video about Operation of Laboratory Photobioreactors with Online Growth Measurements and Customizable Light Regimes at JoVE.com

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

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, ...

operation-laboratory-photobioreactors-with-online-growth-measurements

Research

JoVE Journal

Biology

2.0K Views.  University of Calgary. 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.

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

Presynaptically Silent Synapses Studied with Light Microscopy

Synaptic plasticity likely underlies the nervous system's ability to learn and remember and may also represent an adaptability that prevents otherwise damaging insults from becoming neurotoxic.  We have been studying a form of presynaptic plasticity that is interesting in part because it is expressed as a digital switching on and off of a presynaptic terminal s ability to release vesicles containing the neurotransmitter glutamate.  Here we demonstrate a protocol for visualizing the activity status of presynaptic terminals in dissociated cell cultures prepared from the rodent hippocampus.  The method relies on detecting active synapses using staining with a fixable form of the styryl dye FM1-43, commonly used to label synaptic vesicles.  This staining profile is compared with immunostaining of the same terminals with an antibody directed against the vesicular glutamate transporter 1 (vGluT-1), a stain designed to label all glutamate synapses regardless of activation status.   We find that depolarizing stimuli induce presynaptic silencing.  The population of synapses that is silent under baseline conditions can be activated by prolonged electrical silencing or by activation of cAMP signaling pathways.

Glutamatergic synapses can switch from an active mode to a silent mode. We demonstrate that presynaptic activity status in dissociated culture of rodent neurons is visualized using a fixable form of the FM1-43 dye to visualize active synapses and immunostaining with vGluT-1 antibody to visualize all glutamate synapses.

Synaptic plasticity likely underlies the nervous system's ability to learn and remember and may also represent an adaptability that prevents otherwise ...

Aseptic Laboratory Techniques: Volume Transfers with Serological Pipettes and Micropipettors

Microorganisms are everywhere - in the air, soil, and human body as well as on inanimate surfaces like laboratory benches and computer keyboards. The ubiquity of microbes creates a copious supply of potential contaminants in a laboratory. To ensure experimental success, the number of contaminants on equipment and work surfaces must be minimized. Common among many experiments in microbiology are techniques involving the measurement and transfer of cultures containing bacterial cells or viral particles. To do so without contacting non-sterile surfaces or contaminating sterile media requires (1) preparing a sterile workspace, (2) precisely setting and accurately reading instruments for aseptic transfer of liquids, and (3) properly manipulating instruments, cultures flasks, bottles and tubes within a sterile field. Learning these procedures calls for training and practice. At first, actions should be slow, deliberate, and controlled with the goal being for aseptic technique to become second nature when working at the bench. Here we present the steps for measuring volumes using serological pipettes and micropipettors within a sterile field created by a Bunsen burner. Volumes range from microliters (&mu;l) to milliliters (ml) depending on the instrument used. Liquids commonly transferred include sterile broth or chemical solutions as well as bacterial cultures and phage stocks. By following these procedures, students should be able to: &#8226;Work within the sterile field created by the Bunsen burner flame. &#8226;Use serological pipettes without compromising instrument sterility.&#8226; Aspirate liquids with serological pipettes, precisely reading calibrated volumes by aligning the meniscus formed by the liquid to the graduation marks on the pipette. &#8226;Keep culture bottles, flasks, tubes and their respective caps sterile during liquid transfers. &#8226;Identify different applications for plastic versus glass serological pipettes. &#8226;State accuracy limitations for micropipettors. &#8226;Precisely and accurately set volumes on micropipettors. &#8226;Know how to properly use the first and second stop on a micropipettor to aspirate and transfer correct volumes.

When working in a laboratory, it is imperative to minimize sources of contamination. Aseptic technique refers to procedures that permit transfer of cultures and reagents while avoiding contact with non-sterile surfaces. Serological pipettes and micropipettors are used to measure precise volumes without compromising sterility of solutions used in experiments.

Microorganisms are everywhere - in the air, soil, and human body as well as on inanimate surfaces like laboratory benches and computer keyboards. The ...

In vivo Clonal Tracking of Hematopoietic Stem and Progenitor Cells Marked by Five Fluorescent Proteins using Confocal and Multiphoton Microscopy

We developed and validated a fluorescent marking methodology for clonal tracking of hematopoietic stem and progenitor cells (HSPCs) with high spatial and temporal resolution to study in vivo hematopoiesis using the murine bone marrow transplant experimental model. Genetic combinatorial marking using lentiviral vectors encoding fluorescent proteins (FPs) enabled cell fate mapping through advanced microscopy imaging. Vectors encoding five different FPs: Cerulean, EGFP, Venus, tdTomato, and mCherry were used to concurrently transduce HSPCs, creating a diverse palette of color marked cells. Imaging using confocal/two-photon hybrid microscopy enables simultaneous high resolution assessment of uniquely marked cells and their progeny in conjunction with structural components of the tissues. Volumetric analyses over large areas reveal that spectrally coded HSPC-derived cells can be detected non-invasively in various intact tissues, including the bone marrow (BM), for extensive periods of time following transplantation. Live studies combining video-rate multiphoton and confocal time-lapse imaging in 4D demonstrate the possibility of dynamic cellular and clonal tracking in a quantitative manner.

In vivo Clonal Tracking of Hematopoietic Stem and Progenitor Cells Marked by Five Fluorescent Proteins using Confocal and Multiphoton Microscopy

Combinatorial 5 fluorescent proteins marking of hematopoietic stem and progenitor cells allows in vivo clonal tracking via confocal and two-photon microscopy, providing insights into bone marrow hematopoietic architecture during regeneration. This method allows non-invasive fate mapping of spectrally-coded HSPCs-derived cells in intact tissues for extensive periods of time following transplantation.

We developed and validated a fluorescent marking methodology for clonal tracking of hematopoietic stem and progenitor cells (HSPCs) with high spatial and ...

A Customizable Chamber for Measuring Cell Migration

Cell migration is a vital part of immune responses, growth, and wound healing. Cell migration is a complex process that involves interactions between cells, the extracellular matrix, and soluble and non-soluble chemical factors (e.g., chemoattractants). Standard methods for measuring the migration of cells, such as the Boyden chamber assay, work by counting cells on either side of a divider. These techniques are easy to use; however, they offer little geometric modification for different applications. In contrast, microfluidic devices can be used to observe cell migration with customizable concentration gradients of soluble factors1,2. However, methods for making microfluidics based assays can be difficult to learn.
Here, we describe an easy method for creating cell culture chambers to measure cell migration in response to chemical concentration gradients. Our cell migration chamber method can create different linear concentration gradients in order to study cell migration for a variety of applications. This method is relatively easy to use and is typically performed by undergraduate students.
The microchannel chamber was created by placing an acrylic insert in the shape of the final microchannel chamber well into a Petri dish. After this, poly(dimethylsiloxane) (PDMS) was poured on top of the insert. The PDMS was allowed to harden and then the insert was removed. This allowed for the creation of wells in any desired shape or size. Cells may be subsequently added to the microchannel chamber, and soluble agents can be added to one of the wells by soaking an agarose block in the desired agent. The agarose block is added to one of the wells, and time-lapse images can be taken of the microchannel chamber in order to quantify cell migration. Variations to this method can be made for a given application, making this method highly customizable.

This protocol details a customizable method to measure cell migration in response to chemoattractants that may also be used to determine the diffusion rate of a drug out of a polymer matrix.

Cell migration is a vital part of immune responses, growth, and wound healing. Cell migration is a complex process that involves interactions between ...

Analysis of the Gap Junction-dependent Transfer of miRNA with 3D-FRAP Microscopy

Small antisense RNAs, like miRNA and siRNA, play an important role in cellular physiology and pathology and, moreover, can be used as therapeutic agents in the treatment of several diseases. The development of new, innovative strategies for miRNA/siRNA therapy is based on an extensive knowledge of the underlying mechanisms. Recent data suggest that small RNAs are exchanged between cells in a gap junction-dependent manner, thereby inducing gene regulatory effects in the recipient cell. Molecular biological techniques and flow cytometric analysis are commonly used to study the intercellular exchange of miRNA. However, these methods do not provide high temporal resolution, which is necessary when studying the gap junctional flux of molecules. Therefore, to investigate the impact of miRNA/siRNA as intercellular signaling molecules, novel tools are needed that will allow for the analysis of these small RNAs at the cellular level. The present protocol describes the application of three-dimensional fluorescence recovery after photobleaching (3D-FRAP) microscopy to elucidating the gap junction-dependent exchange of miRNA molecules between cardiac cells. Importantly, this straightforward and non-invasive live-cell imaging approach allows for the visualization and quantification of the gap junctional shuttling of fluorescently labeled small RNAs in real time, with high spatio-temporal resolution. The data obtained by 3D-FRAP confirm a novel pathway of intercellular gene regulation, where small RNAs act as signaling molecules within the intercellular network.

Here, we describe the application of three-dimensional fluorescence recovery after photobleaching (3D-FRAP) for the analysis of the gap junction-dependent shuttling of miRNA. In contrast to commonly applied methods, 3D-FRAP allows for the quantification of the intercellular transfer of small RNAs in real time, with high spatio-temporal resolution.

Small antisense RNAs, like miRNA and siRNA, play an important role in cellular physiology and pathology and, moreover, can be used as therapeutic agents ...

Cystometric and External Urethral Sphincter Measurements in Awake Rats with Implanted Catheter and Electrodes Allowing for Repeated Measurements

Lower urinary tract function is mainly assessed by means of cystometric bladder function analysis in rodents. Conventional cystometries are usually performed as terminal analysis under urethane anesthesia. It is well known that anesthetic drugs can influence bladder function. Hence, the aim of this technique is to perform cystometric measurements of the urinary bladder and external urethral sphincter in lightly restrained awake rats. For this purpose, a bladder catheter is implanted into the bladder dome. Subsequently, two electrodes are implanted bilateral to the external urethral sphincter and a ground electrode is sutured to a non-responsive skeletal muscle. The bladder catheter and the three electrodes are finally tunneled subcutaneously to the neck region and affixed to a harness. With this technique, the lower urinary tract can be measured at multiple time points in the same animal to assess lower urinary tract function. The main application of this technique is the follow-up of simultaneous urinary bladder and external urethral sphincter function in awake healthy rats and after induction of a disease or injury. Moreover, subsequent lower urinary tract monitoring can be performed during evaluation of the disease/injury and to monitor treatment efficacy.

This protocol first describes the surgical procedure of the permanent implantation of a urinary bladder catheter combined with external urethral sphincter electrodes, and second, the measurement of the function of the urinary bladder and external urethral sphincter in implanted awake animals.

Lower urinary tract function is mainly assessed by means of cystometric bladder function analysis in rodents. Conventional cystometries are usually ...

Analysis of Arabidopsis thaliana Growth Behavior in Different Light Qualities

Plant biologists often need to observe the growth behavior of their chosen species. To this end, the plants need constant environmental and stable light conditions, which are preferably variable in quantity and quality so that studies under different setups can be conducted. These requirements are met by climatic chambers featuring light emitting diodes (LED) lights, which can &#8211; in contrast to fluorescent lights &#8211; be set to different wavelengths. LEDs are energy conserving and emit virtually no heat even at light intensities, which often constitutes a problem with other light sources. The presented protocol provides a step-by-step guidance of how to program a climatic chamber equipped with variable LED lights as well as describing several approaches for in depth analysis of growth phenotypes. Depending on the experimental set-up various characteristics of the growing plants can be observed and analyzed. Here we describe how to determine fresh weight, leaf area, photosynthetic activity, and stomatal density. We demonstrate that in order to obtain reliable data and draw valid conclusions it is mandatory to use a sufficient number of individuals for statistical evaluation. Taking too few plants for this kind of analysis results in high statistical errors and consequently in less clear interpretations of the data.

Analysis of Arabidopsis thaliana Growth Behavior in Different Light Qualities

Here, we present a protocol for studying plant growth behavior and especially phenotypes in a reproducible manner. We show how to provide variable and at the same time stable light conditions. Proper analyses depend on sufficient sample numbers and valid statistical evaluations.

Plant biologists often need to observe the growth behavior of their chosen species. To this end, the plants need constant environmental and stable light ...

Quantitative Analysis of Aspergillus nidulans Growth Rate using Live Microscopy and Open-Source Software

It is well established that colony growth of filamentous fungi, mostly dependent on changes in hyphae/mycelia apical growth rate, is macroscopically estimated on solidified media by comparing colony size. However, to quantitatively measure the growth rate of genetically different fungal strains or strains under different environmental/growth conditions (pH, temperature, carbon and nitrogen sources, antibiotics, etc.) is challenging. Thus, the pursuit of complementary approaches to quantify growth kinetics becomes mandatory in order to better understand fungal cell growth. Furthermore, it is well-known that filamentous fungi, including Aspergillus spp., have distinct modes of growth and differentiation under sub-aerial conditions on solid media or submerged cultures. Here, we detail a quantitative microscopic method for analyzing growth kinetics of&#160;the model fungus Aspergillus nidulans, using live imaging in both submerged cultures and solid media. We capture images, analyze, and quantify growth rates of different fungal strains in a reproducible and reliable manner using an open source, free software for bio-images (e.g., Fiji), in a way that does not require any prior image analysis expertise from the user.

Quantitative Analysis of Aspergillus nidulans Growth Rate using Live Microscopy and Open-Source Software

We present a label-free live imaging protocol using transmitted light microscopy techniques to capture images, analyze and quantify growth kinetics of the filamentous fungus A. nidulans in both submerged cultures and solid media. This protocol can be used in conjunction with fluorescence microscopy.

It is well established that colony growth of filamentous fungi, mostly dependent on changes in hyphae/mycelia apical growth rate, is macroscopically ...

Application of Passive Head Motion to Generate Defined Accelerations at the Heads of Rodents

Exercise is widely recognized as effective for various diseases and physical disorders, including those related to brain dysfunction. However, molecular mechanisms behind the beneficial effects of exercise are poorly understood. Many physical workouts, particularly those classified as aerobic exercises such as jogging and walking, produce impulsive forces at the time of foot contact with the ground. Therefore, it was speculated that mechanical impact might be implicated in how exercise contributes to organismal homeostasis. For testing this hypothesis on the brain, a custom-designed ''passive head motion'' (hereafter referred to as PHM) system was developed that can generate vertical accelerations with controlled and defined magnitudes and modes and reproduce mechanical stimulation that might be applied to the heads of rodents during treadmill running at moderate velocities, a typical intervention to test the effects of exercise in animals. By using this system, it was demonstrated that PHM recapitulates the serotonin (5-hydroxytryptamine, hereafter referred to as 5-HT) receptor subtype 2A (5-HT2A) signaling in the prefrontal cortex (PFC) neurons of mice. This work provides detailed protocols for applying PHM and measuring its resultant mechanical accelerations at rodents' heads.

The present protocol describes a custom-designed ''passive head motion'' system, which reproduces mechanical accelerations at rodents' heads generated during their treadmill running at moderate velocities. It allows dissecting mechanical factors/elements from the beneficial effects of physical exercise.

Exercise is widely recognized as effective for various diseases and physical disorders, including those related to brain dysfunction. However, molecular ...

Characterizing Mechanical Properties of Primary Cell Wall in Living Plant Organs Using Atomic Force Microscopy

The mechanical properties of the primary cell walls determine the direction and rate of plant cell growth and, therefore, the future size and shape of the plant. Many sophisticated techniques have been developed to measure these properties; however, atomic force microscopy (AFM) remains the most convenient for studying cell wall elasticity at the cellular level. One of the most important limitations of this technique has been that only superficial or isolated living cells can be studied. Here, the use of atomic force microscopy to investigate the mechanical properties of primary cell walls belonging to the internal tissues of a plant body is presented. This protocol describes measurements of the apparent Young&#39;s modulus of cell walls in roots, but the method can also be applied to other plant organs. The measurements are performed on vibratome-derived sections of plant material in a liquid cell, which allows (i) avoiding the use of plasmolyzing solutions or sample impregnation with wax or resin, (ii) making the experiments fast, and (iii) preventing dehydration of the sample. Both anticlinal and periclinal cell walls can be studied, depending on how the specimen was sectioned. Differences in the mechanical properties of different tissues can be investigated in a single section. The protocol describes the principles of study planning, issues with specimen preparation and measurements, as well as the method of selecting force-deformation curves to avoid the influence of topography on the obtained values of elastic modulus. The method is not limited by sample size but is sensitive to cell size (i.e., cells with a large lumen are difficult to examine).

Studies of cell wall biomechanics are essential for understanding plant growth and morphogenesis. The following protocol is proposed to investigate thin primary cell walls in the internal tissues of young plant organs using atomic force microscopy.

The mechanical properties of the primary cell walls determine the direction and rate of plant cell growth and, therefore, the future size and shape of the ...