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, 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 0–2400 µmol 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.
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't possible15. A single method is not suited to all experimental applications—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 8–10% 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 0.1–3 g/L12,21,22. Small volumes of dilute suspensions yield little dry biomass, <10 mg. After combustion, ash may only weigh 1 mg. Therefore, depending on the culture density, this method requires volumes between 5–100 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 2–3 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'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.
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 3.2–3.6). According to the manufacturer's instruction, the gas sensor packing liquid should be filled to the "tracing screw for liquid level" 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 (<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 400–700 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 µmolphotons 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 2–3-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.
The authors have nothing to disclose.
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.
Aluminum channels Imperial: 0.90” x 39.37” Metric: 2.3 cm x 100 cm Quantity: 4 |
LED World | AC-AR1-1M | Required as a heat sink |
Bungee cords, small Quantity: 5 |
– | – | To secure bottles |
Computer – desktop/laptop Quantity: 1 |
– | – | – |
Data Logger, HOBO U30 USB Weather Station Quantity: 1 |
HOBO, Hoskin | U30-NRC-VIA-10-S100-000 | Records light sensor information |
Digital interface module, Rigamo, 4-channel Quantity: 1 |
Ritter | N/A | This is to transmit gas sensor data to the computer |
DMX decoder, 12~24 VDC, DMX-CV-4X5A Quantity: 1 |
LITECH, LED World | LT-840-6A | Transmit messages which alter the light pattern |
DMX lighting controller, SUSHI-RB-RJ Quantity: 1 |
Arcolis, Nicolaudie America Inc. | SUSHI-RB-RJ DMX | Encodes the lighting program |
Gas sensor packing liquid (Silox) Quantity: 1 L |
Ritter | https://www.ritter.de/en/data-sheets/silox | |
Gas sensor, volumetric Quantity: 3 |
Ritter | MGC-1 V3.4 PMMA (https://www.ritter.de/downloads/mgc-milligascounter-en) | Measures oxygen production |
Glass bottles, round 1 L with GL45 neck Quantity: 3 |
Corning, Capitol Scientific | 1395-1L | Culture vessels |
Hardware – end caps for slotted steel Quantity: 10 |
Paulin, Home Depot | 142-612 | To cover sharp edges of slotted steel |
Hardware – eye hooks Quantity: 6 |
– | – | To secure bottles |
Hardware – metal corner braces (large) Imperial: 4" x 4" Metric: 10 cm x 10 cm Quantity: 8 |
– | – | Larger brackets to construct metal stand |
Hardware – metal corner braces (small) Imperial: 2 1/2" x 2 1/2" Metric: 6.4 cm x 6.4 cm Quantity: 6 |
– | – | Small brackets to connect bottle platforms to PVC pipe |
Hardware – metal corner gussets Imperial: 3" x 3" Metric: 7.6 cm x 7.6 cm Quantity: 6 |
Paulin, Home Depot | 142-616 | Flat brackets to construct metal stand |
Hardware – piano hinge Imperial: 36" Metric: 91 cm Quantity: 1 |
– | – | Connects two halves of PVC pipe |
Hardware – rivets Quantity: 40 |
– | – | To attach piano hinge to PVC tubing |
Hardware – set of bolts, nuts, washers Quantity: 60 |
– | – | Long thin bolts are required to secure bottle platforms around magentic stirrers |
Hardware – set of bolts, nuts, washers Quantity: 30 |
– | – | Larger shorter bolts are required to build the metal stand |
LED driver, constant voltage, 96W 24VDC UL Listed IP65 Driver Class 2 regulated power supply Quantity: 1 |
Magnitude Lighting, LED World | CVN96L24DC | Regulates power to the lights |
LED lights, Cinco Bright LED Flex Strip Quantity: 4 m roll |
EvenBright, LED World | FA128M57-4M-24V-X | Roll is trimmed into 4 x 1 m lengths and secured inside the PVC tube |
Light meter, handheld with submersible sperical probe Quantity: 1 |
LI-COR | LA-250A | Calibrate the reactors light intensity |
Light sensors Photosynthetic Light (PAR) Smart Sensor Quantity: 2 |
HOBO, Hoskin | S-LIA-M003 | Only one is required however two would be good practice in case one malfunctioned |
Magnetic stirrers (MIXdrive 1 XS) with external control units and power supply (MIXcontrol eco) Quantity: 3 |
2Mag, 2MAG USA | MF 40300 | Stirrers sit sandwiched in bottle platforms |
Metal plate Imperial: 24" x 8" Metric: 61 cm x 20.3 cm Quantity: 1 |
– | – | This is a surface on which to secure electronics, it is attached to the back of the reactor |
Pipe, white PVC Imperial: 6" diameter x 42" high Metric: 15.2 cm x 106.7 cm Quantity: 1 |
– | – | Cut lengthwise in two halves, used to house lights and bottles |
Plastic (HDPE) sheets Imperial: 4" x 4" x 1/4" Metric: 10 cm x 10 cm x 1 cm Quantity: 6 |
Inventables | 30291-01 | For bottle platforms which house magentic stirrers |
Rubber stoppers – GL45 size Quantity: 3 |
Duran, VWR | 76289-760 | Seals culture vessels |
Screw caps – with aperture and GL45 neck Quantity: 3 |
Corning, Capitol Scientific | 1395-45HTSC | Generates seal of culture vessels |
Slotted angle steel lengths Imperial: 1-1/2" X 48" x 0.074" Metric: 3.8 cm x 122 cm x 0.19 cm Quantity: 6 |
Paulin, Home Depot | 142-202 | Makes up the body of the metal stand |
Slotted flat steel lenghts Imperial: 1-3/8" x 48" x 0.074" Metric: 3.5 cm x 122 cm x 0.19 cm Quantity: 3 |
Paulin, Home Depot | 142-222 | Makes up the body of the metal stand |
Software – Easy Stand Alone (ESA) | https://www.dmxsoft.com/#apps | AKA LED control software | |
Software – Rigamo v3.1 | AKA data acquisition software | ||
Software – Storage Upgrade Tools (SUT) | https://store.dmxsoft.com// | ||
Stir bar Imperial: 1" x 5/16" Metric: 2.5 cm x 0.8 cm Quantity: 3 |
Fisherbrand | 14-513-59 | Stirs culture |
Switch box Quantity: 1 |
– | – | Turns power on/off to reactor |
Syringe, 10 mL Quantity: Multiple |
– | – | Optional if you wish to extract culture |
Tube adaptor fittings, plastic – Stopcock 1-way Quantity: 6 |
Masterflex, Cole Palmer | RK-12023-33 | Close/open culture vessel line |
Tube adaptor fittings, plastic – variety of male and female luer lock fittings Imperial: to fit 1/16" and 1/8" ID tubing Metric: to fit 1.59 mm and 3.18 mm tubing Quantity: Multiple packets |
Masterflex, Cole Palmer | RK-30800-16; RK-30800-18; RK-45518-26; RK-45501-00; RK-45501-04 | Many different combinations can achieve the same end result, best to order a variety of fittings |
Tube adaptor fittings, plastic – variety of straight connectors Imperial: to fit 1/16" and 1/8" ID tubing Metric: to fit 1.59 mm and 3.18 mm tubing Quantity: Multiple packets |
Masterflex, Cole Palmer | RK-40616-04 | Many different combinations can achieve the same end result, best to order a variety of fittings |
Tubing, flexible, transparent Imperial: ID=1/16", OD=1/8" Metric: ID = 1.59 mm, 0D = 3.18 mm Quantity: 4 m |
Masterflex, Cole Palmer | RK-06422-02 | Line from culture vessel to gas sensor |
Tubing, flexible, transparent Imperial: ID=1/8", OD=1/4" Metric: ID = 3.18 mm, 0D = 6.35 mm Quantity: 2 m |
Masterflex, Cole Palmer | RK-06422-05 | Gas sensor standard tubing size |
Tubing, rigid, transparent Imperial: ID=1/16", OD=1/8" Metric: ID = 1.59 mm, 0D = 3.18 mm Quantiy: 1 m |
Masterflex, Cole Palmer | RK-06605-27 | Spans rubber stopper allowing gas to exit |
Tubing, rigid, transparent Imperial: ID=1/8", OD=1/4" Metric: ID = 3.18 mm, 0D = 6.35 mm Quantity: 1 m |
Masterflex, Cole Palmer | RK-06605-30 | Spans rubber stopper allowing gas to exit |
Zip ties, small Quantity: 1 packet |
Secure tube fittings |